The authors wish to thank Drs. David Calkins (Vanderbilt University) and James Morgan (Cardiff University) for sharing their expertise and for helpful advice towards developing this procedure. This study was supported by grants from the Canadian Institutes of Health Research (A.D.P.). Y.A.I. and N.B. are the recipients of postdoctoral fellowships from the Fonds de recherche du Qu&#233;bec-Sant&#233; (FRQS). N.B. was awarded a H.H. Jasper scholarship from the Groupe de Recherche sur le Syst&#232;me Nerveux Central (GRSNC). A.D.P. is a Chercheur Boursier National FRQS.

The following procedure was performed in compliance with the guidelines of the Canadian Council on Animal Care for the Use of Experimental Animals and the Statement for the Use of Animals in Ophthalmic and Visual Research from the Association for Research in Vision and Ophthalmology (ARVO).
1. Preparation of the Microneedle for Anterior Intracameral Injection
<ol>
	<li>With a puller, generate a microneedle from a borosilicate glass capillary</li>
	<li>Under a microscope, use a sharp blade to carefully create an opening at the tip of the microneedle. The resulting opening should have an elliptical shape with a major and minor axis diameter of approximately 190 &#181;m and 70 &#181;m, respectively. Measure the area of the prepared microneedle opening by first acquiring images with a ruler placed under the microscope followed by quantification using image analysis software.</li>
	<li>In the micropipette bevelling system, place the microneedle at a 20 degree angle relative to the bevelling plate so that the microneedle opening is touching the plate. Bevel for approximately 10 min until the edges are flat and smooth. Add a few drops of distilled water to aid in the process.</li>
	<li>Rotate the microneedle to bevel the two edges surrounding the opening until the tip is sharp.</li>
	<li>Clean all debris and water from the microneedle tip opening using an aerosol duster.</li>
	<li>Carefully examine the finished microneedle under a microscope. Discard microneedles with fractures to minimize the risk of the microneedle breaking during surgery.</li>
	<li>Sterilize the microneedle by rinsing first with ethanol, then with sterile balanced salt solution (BSS).</li>
</ol>
2. Preparation of the Magnetic Microbead Solution
Note: The magnetic microbeads used in this study are coated with epoxy groups. To prevent any adverse effects, such as clumping of the beads and unwanted molecular interactions, these epoxy groups must first be removed from the microbeads before proceeding with the injection surgery.
<ol>
	<li>Removal of Epoxy Groups from Magnetic Beads
	<ol>
		<li>Prepare a solution of 0.02 M sodium hydroxide (NaOH, MW 39.997 g/mol) in 10x Tris buffer (MW 121.14 g/mol).</li>
		<li>Gently vortex the stock of magnetic microbead solution (4.5 &#181;m diameter, 4 x 108 beads/ml) until the beads are evenly suspended in solution.</li>
		<li>Quickly pipet 1 ml of magnetic bead solution into 50 ml of 0.02 M NaOH in 10x Tris buffer.</li>
		<li>Rotate for 24 hr&#160;at RT&#160;to remove the epoxy groups from the beads.</li>
		<li>Collect the beads by securing a magnet to the bottom of the tube. Orient the tube horizontally to ensure that all the beads are attracted to the magnet. Rotate for an additional 4 hr&#160;at RT.</li>
		<li>With a micropipette, carefully remove the supernatant without disturbing the pellet of beads.</li>
		<li>Gently vortex the pellet in 50 ml of 10x Tris buffer until the beads are well suspended.</li>
		<li>Repeat steps 2.1.4 to 2.1.6.</li>
	</ol>
	</li>
	<li>Concentration and Resuspension of Magnetic Microbeads in Sterile Balanced Salt Solution</li>
</ol>
Note: It is necessary to concentrate the stock of magnetic bead solution in sterile balanced salt solution (BSS) to reach a final concentration of 1.6 x 106 beads/&#181;l so that 2.4 x 106 beads can be injected into the anterior chamber in a final volume of 1.5 &#181;l, which is appropriate for the small mouse eye.
<ol>
	<li>Wash the beads in 5 ml of ultra-pure laboratory grade water by gently vortexing for 2 min.</li>
	<li>Collect the beads by attracting them to the bottom of the tube with a magnet.</li>
	<li>With a micropipette, carefully remove the water without disturbing the pellet of beads.</li>
	<li>Repeat steps 2.2.1 to 2.2.3 three more times.</li>
	<li>In a laminar flow hood, wash the beads with 500 &#181;l of BSS by pipetting up and down. Perform the remaining steps in this section under sterile conditions in a laminar flow hood.</li>
	<li>Collect the beads by attracting them to the bottom of the tube with a magnet.</li>
	<li>With a micropipette, carefully remove the BSS without disturbing the pellet of beads.</li>
	<li>Repeat steps 2.2.5 to 2.2.7 three more times.</li>
	<li>Resuspend by pipetting the beads up and down in 250 &#181;l of BSS.</li>
	<li>Ensure that the bead solution is well homogenized. Then, quickly aliquot 25 &#181;l of the suspension into sterile 0.5 ml tubes. The final concentration of the stock bead solution is 1.6 x 106 beads/&#181;l.</li>
	<li>Store at 4 &#176;C.</li>
</ol>
3. Induction of Ocular Hypertension
Note: Section 3 is a two-person operation. In the event that a particular action is to be performed by a specific person, the appropriate person is identified. In general, Person 1 handles the mouse under the microscope while Person 2 is responsible for manipulating the microsyringe pump. The total duration of the surgical procedure should be less than 10 min (steps 3.9 to 3.17).
<ol>
	<li>Perform procedures in adult C57BL/6 mice. House mice in a standard environment with access to food and water ad libitum. In this paper, use female C57BL/6 mice between 3 and 4.5 months of age. However, this protocol can be adapted to males and mice of different ages, as well as other mouse strains, including transgenic and knockout mice.</li>
	<li>Measure baseline IOP in awake mice prior to anaesthesia and microbead injection using a calibrated rebound tonometer. Apply one drop of proparacaine hydrochloride on the cornea.
	<ol>
		<li>Gently restrain the mouse by holding the skin between the ears. Place the mouse on the benchtop so that the animal is comfortable and the eyes are accessible. Hold the tonometer perpendicular to the corneal surface and take at least three sets of ten consecutive readings per eye to obtain an IOP average. The measuring of IOP in awake mice is preferred to bypass anaesthesia-related effects on IOP. Alternatively, IOP can be measured in anesthetised mice after step 3.4.</li>
	</ol>
	</li>
	<li>Prepare a stock mouse cocktail anaesthesia mixture composed of 20 mg/ml ketamine, 2 mg/ml xylazine, and 0.4 mg/ml acepromazine.</li>
	<li>Induce anaesthesia in the mouse by intraperitoneal administration of the cocktail mixture (1 &#181;l/g of body weight). The use of an injectable anaesthetic cocktail is preferred over gas anaesthetics (e.g., isoflurane) because it allows flexibility when handling the mouse head as the animal is not connected to an inhalation mask. In addition, the longer recovery period required with an injectable anaesthetic ensures that microbeads settle at the iridocorneal angle without dislodging back into the anterior chamber.</li>
	<li>Administer 0.05 mg per kg of body weight of buprenorphine subcutaneously.</li>
	<li>Treat the eye with a tropicamide eye drop to induce pupil dilation. Due to the small size of the murine anterior chamber, the pupil must be dilated to easily visualize the positioning and advancement of the microneedle during injection.</li>
	<li>Apply topical ointment on the contralateral eye (un-operated) to avoid drying of the cornea during the procedure.</li>
	<li>Attach a clean microneedle to the injection assembly of the microsyringe pump. Replace the microneedle after every operation to avoid cross-animal contamination.</li>
	<li>Person 1: Transfer the anesthetised mouse to the operating platform. Under the microscope, ensure that the pupil is fully dilated and that the ocular muscles are relaxed so that there is no eye movement. The absence of eye movements ensures stability during the injection. Gently wipe the tropicamide eye drop from the eye using absorbent swabs.</li>
	<li>Person 2: Mix the magnetic microbead solution by pipetting up and down.</li>
	<li>Using the microsyringe pump, immediately load the microneedle (prepared in section 1) with 1.5 &#181;l of the homogenized magnetic microbead solution (2.4 x 106 beads). Ensure that air bubbles are absent at the tip of the microneedle. After the microneedle is loaded, carry out steps 3.12 to 3.13 as quickly as possible so that the magnetic microbead solution remains in a homogeneous suspension.</li>
	<li>Position the loaded microneedle at a 45&#176; angle, placed anteriorly relative to the limbus. Person 1: support the eye using plastic forceps. Ensure that the angle between the microneedle and the plastic forceps is approximately 90&#176;.</li>
	<li>Person 2: With the loaded microneedle, gently puncture the cornea so that the tip of the microneedle enters the anterior chamber. Ensure that the loaded microneedle remains at a 45&#176; angle relative to the limbus during the puncture. Avoid any contact with the lens or the iris. Ensure that the microneedle does not enter the posterior chamber. Person 1: continue to support the eye using plastic forceps.</li>
	<li>Person 1: Without moving the mouse head, place the magnet beside the eye, opposite to the microneedle tip, to attract the magnetic beads into the anterior chamber and minimize contact of the beads with the inner surface of the cornea. Person 2: Using the microsyringe pump, inject 1.5 &#181;l of the magnetic bead solution into the anterior chamber. The microbead solution is injected over a period of 15 to 30 sec. Person 1: Continue to hold the magnet opposite to the microneedle tip during the entire duration of the injection.</li>
	<li>Person 2: Once the full volume of beads has been injected, slowly withdraw the microneedle from the eye. Person 1: To avoid reflux of the microbeads, continue to attract the magnetic beads towards the anterior chamber by holding the magnet next to the eye for an additional 30 to 60 sec.</li>
	<li>Person 1: Using the magnet, attract the beads to the iridocorneal angle. Ensure that the beads form an evenly distributed ring around the circumference of the anterior chamber. During this step, avoid attracting beads to the cornea as they have a tendency to stick to the inner surface of the cornea upon contact.</li>
	<li>Treat the operated eye with an antibiotic eye drop to minimize the risk of infection.</li>
	<li>Allow the mouse to recover on a heat pad until fully awake (~3 to 4 hr). Place the mouse so that the operated eye is facing upwards. This positioning will prevent accumulation of the injected beads to the inner surface of the cornea by gravitational pull. In addition, this position will decrease the potential for infection as the operated eye will not be in contact with any bedding and/or other materials that may be present in the cage.&#160;In the event that an animal displays any sign of distress, administer additional doses of buprenorphine as needed.&#160;</li>
	<li>Allow mice to recover from the procedure for at least 2 days before taking IOP measurements. Measure IOP as described in 3.2. Monitor IOP at least once a week or more frequently, as needed, and at the same time of the day to minimize circadian-related fluctuations.</li>
	<li>For IOP measurements, use the contralateral eye from the operated mouse as an internal control. Alternatively, use measurements from intact, non-operated or sham-operated mice as na&#239;ve controls.</li>
</ol>
4. Assessment of Retinal Ganglion Cell Soma and Axon Survival
<ol>
	<li>Perfuse mice subjected to ocular hypertension by intracardial injection of 0.1 M phosphate buffer solution (PBS) immediately followed by ice-cold 4% paraformaldehyde (PFA).</li>
	<li>Perfuse intact, non-operated mice as described in 4.1 and use them as uninjured controls. The use of contralateral eyes from operated mice is not recommended to assess RGC survival because changes in contralateral eyes following injury have been reported15,16 and can confound data interpretation. Alternatively, use sham-operated eyes injected with 1.5 &#181;l of BSS as controls.</li>
	<li>Using microscissors, carefully cut the connective tissue around the eye to isolate it from the eye socket. Separate the optic nerve from the eye by cutting it at the level of the optic nerve head.</li>
	<li>Using a 30 G needle, make a hole in the cornea to allow penetration of the fixative solution into the eye. Place the eye in 4% PFA and incubate for 1 hr at 4 &#176;C for additional fixation.</li>
	<li>Place the optic nerve in a solution containing 2% PFA and 2.5% glutaraldehyde in 0.1 M sodium cacodylate (MW: 214 g/mol) and incubate O/N&#160;at 4 &#176;C for additional fixation.</li>
</ol>
5. Quantification of RGC Soma Density on Flat-mounted Retinas
Note: The following procedure is adapted from a protocol by Nadal-Nicolas et al. 17 and outlines the quantification of RGCs using an antibody against the brain-specific homeobox/POU domain protein 3A (Brn3a) on retinal flat-mounts. Alternative methods for labeling RGCs can also be used including immunohistochemistry with an antibody against RNA-binding protein with multiple splicing (RBPMS) or retrograde labelling with Fluorogold or DiI.
<ol>
	<li>Under a dissecting microscope, remove the anterior part of the eye by making an incision along the entire limbus until the cornea detaches easily. Remove the cornea and lens. Carefully detach the retina from the eye by making cuts along the ora serrata and optic nerve.</li>
	<li>Prepare a retinal flat-mount by making four small equidistant incisions from the periphery of the retina towards the optic nerve to clearly delineate the four retinal quadrants. Using a small brush, gently remove any remaining vitreous from the retina. The removal of as much vitreous humour as possible is crucial for obtaining a clean and strong immunohistochemical signal.</li>
	<li>Carefully transfer the retina to a 48-well flat bottom culture plate containing 0.5% Triton X-100 in PBS so that the retina is free-floating. Make sure the ganglion cell layer is facing upwards.</li>
	<li>Place the culture plate at -70 &#176;C for 15 min. After thawing the retinas, further permeabilize by washing twice in fresh 2% Triton X-100 in PBS.</li>
	<li>Dilute the Brn3a antibody to 0.3 - 0.5 &#181;g/ml with PBS containing 2% Triton X-100 and 2% normal donkey serum. Incubate the retina in the Brn3a primary antibody solution by gently shaking O/N at 4 &#176;C. The retinas should be fully immersed in 150 to 200 &#181;l of solution at all times.</li>
	<li>Dilute the donkey anti-goat IgG secondary antibody to 2 &#181;g/ml with PBS containing 2% Triton X-100 and incubate the retina in this solution for 2 hr&#160;at RT with gentle shaking. The retina should be fully immersed in the antibody solution at all times. Cover the culture plate with aluminum foil for the remaining steps to prevent photobleaching.</li>
	<li>Using a brush, carefully transfer the retina to a slide so that the tissue lies as flat as possible. Air dry for 10 min. Mount using anti-fade mounting medium.</li>
	<li>Examine the retinal flat-mounts under a fluorescent microscope. Quantify the number of Brn3a positive RGCs in three non-overlapping areas per retinal quadrant as described.18</li>
</ol>
6. Quantification of RGC Axons on Optic Nerve Cross Sections&#160;
<ol>
	<li>Collect the optic nerve from step 4.5 and incubate in 2% osmium tetroxide (OsO4, MW 254.23 g/mol) for 2 hr. 
	Caution: Due to its high toxicity, osmium tetroxide should be handled in a fume hood with appropriate laboratory attire.</li>
	<li>Dehydrate the optic nerve by immersing it in increasing concentrations of ethanol (50%, 70%, 90%, 95%, and 100%) for 15 min each.</li>
	<li>Prepare the epoxy resin using the following recipe: 15.72 ml Embed-812, 6.45 ml dodecenyl succinic anhydride, 7.83 ml nadic methyl anhydride, 0.45 ml DMP-30.</li>
	<li>Sequentially incubate the optic nerve in solutions composed of the following ratios of epoxy resin to propylene oxide (MW 58.08 g/mol): 0:1, 1:1, 0.75:0.25, and 1:0. The optic nerve is incubated in each solution at RT for an O/N period.</li>
	<li>Incubate optic nerves embedded in 100% epoxy resin (last step from 6.4) at 60 &#176;C for an additional 48 hr.</li>
	<li>Generate semi-thin optic nerve cross sections (0.75 &#181;m) using a microtome.</li>
	<li>Stain optic nerve cross sections with 1% toluidine blue.</li>
	<li>Quantify the RGC axons in five non-overlapping areas of each optic nerve section as described 19.</li>
</ol>

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The authors declare that they have no competing financial interests.

The video technique presented here provides detailed step-by-step instructions on how to perform intracameral injection of magnetic microbeads to effectively and reproducibly induce IOP elevation in mice. This procedure results in sustained IOP increase that does not require additional injections and promotes detectable RGC soma and axon loss within the first 3 weeks of ocular hypertension induction.Elevated IOP is a major risk factor for developing glaucoma in humans. Therefore, this is a valuable murine ocular hypertension-dependent glaucoma model that has potential for a wide range of applications.
A common drawback associated with the injection of microbeads into the anterior chamber relates to bead reflux through the injection site when the needle is withdrawn, which often results in only partial obstruction of aqueous outflow and increased variability. To address this issue, several important modifications were implemented. First, the careful preparation of a clean, sharp glass microneedle with a facetted bevel is essential for the successful injection of the microbeads. A properly prepared microneedle enables controlled and smooth penetration of the cornea with minimal application of pressure to the delicate ocular surface. The small corneal puncture prevents backflow of microbeads. In addition, the fine microneedle reduces the risk of damaging nearby structures such as the iris and the lens, which could result in non-disease related inflammation. Secondly, the application of a handheld magnet to strategic ocular areas during and after the injection is another critical aspect of this technique. During the injection, the magnet is used to draw the magnetic microbeads to the anterior chamber preventing reflux of the microbeads when the microneedle is withdrawn. After the injection, the magnet is then used to direct the microbeads to the iridocorneal angle to block aqueous humour outflow.
Another problem often encountered in microbead occlusion models is that repeated bead injections are often necessary to achieve sustained IOP elevation 10,11. This might be the result of microbeads dislodging from the iridocorneal angle with time. The combination of a handheld magnet, as described above, and the positioning of the mouse post-operatively greatly improves the outcome. The use of injectable anaesthetics, which allow flexibility to move the head during the procedure and require a longer post-operative recovery period, is favoured. Placement of the mouse with the operated eye facing upwards for a couple of hours after the surgery contributes to the settlement of microbeads at the iridocorneal angle and decreases the risk of dislodging back into the anterior chamber.
Ensuring that the number of injected beads is relatively consistent is another critical step to minimize inter-animal variations. Since the microbeads settle at the bottom of the tube, it is necessary to fully homogenize the microbead solution and withdraw the appropriate volume into the microneedle in a timely manner. Injection of fewer beads into the anterior chamber could result in incomplete blockage of the aqueous humour drainage structures, which is likely to result in poor or variable IOP elevation. Of note, although the ultimate purpose of the microbead injection is to elevate IOP, caution should be taken when IOP measurements from awake mice are higher than the peak values reported in this study (~25 mmHg). Extremely high IOPs increase the risk of ischemic damage and may also cause pain to the animal. The elevation of IOP should be considered as one of many factors to assess the success of the surgery. As such, the outcome of the procedure should be gauged based on several parameters including IOP elevation, RGC soma death, and axon loss.
Although the protocol described here results in most microbeads successfully settling at the angle, a potential limitation of this model is that those beads that remain floating in the anterior chamber might interfere with live retinal imaging through the cornea, as well as electrophysiological or behavioral assays which require effective passage of light. Another important aspect to consider when utilizing this microbead occlusion model is that the extent of IOP elevation and subsequent RGC degeneration varies with the age and genetic background of the operated mouse [4]. Therefore, the extent of IOP elevation and the timeline of RGC degeneration will need to be determined for each specific transgenic mouse line and/or age range.
A feature of this model is that elevated IOP results in the gradual loss of RGC death during the first three weeks after microbead injection, and significant RGC death is detected at 3 weeks after the procedure. Hence, this model enables the examination of early and/or subtle changes that occur in this disease, prior to overt RGC soma and axon loss. A significant increase in RGC death was not observed between 3 and 6 weeks after induction of ocular hypertension. In fact, RGC soma and axon loss remained stable at ~22 - 25% between 3 and 6 weeks in spite of successful and sustained IOP elevation at these time points. A longer duration of sustained IOP may be required for additional RGC loss to occur in C57BL/6 mice, which appear to be more resistant to RGC damage compared to other mouse strains.5 Additional modifications to the protocol presented here, including adjustment of bead size and additional injections, might be required to study RGC loss at later time points. Therefore, our protocol is ideal for studies focused on early pathophysiological changes that correlate with modest RGC neurodegeneration which are relevant to onset and early progression in human glaucoma.

Glaucoma is a progressive and irreversible blinding condition that will affect an estimated 80 million people worldwide by 20201. In glaucoma patients, vision loss is caused by the selective death of retinal ganglion cells (RGCs), the output neurons that transmit visual information from the retina to the brain. Glaucoma is an age-related neurodegenerative disease with many risk factors of which the most common is elevated intraocular pressure (IOP). Indeed, IOP is the only modifiable risk factor in glaucoma and current treatments focus solely on managing eye pressure. However, multiple genetic, cellular, and environmental factors affect the onset and progression of this disease. Therefore, understanding the various mechanisms that ultimately contribute to neuronal death is essential to develop effective treatments for glaucoma.
Animal models of glaucoma are essential to study disease pathophysiology and to identify and test promising therapeutics. The increasing availability of transgenic mouse lines including conditional knockout strains and mice carrying genetically-encoded fluorescent tracers has propelled the need for inducible murine glaucoma models. Several rodent models of glaucoma have been developed over the years (reviewed in2,3). In many of these models, glaucoma is induced by disrupting aqueous humour dynamics, resulting in the elevation of IOP. Occlusion models, in which microbeads or other substances are injected into the anterior chamber of the eye to block aqueous drainage, have gained popularity in recent years partly due to their relative ease to increase IOP4-14.
The microbead occlusion model of glaucoma, first carried out in primates 12, rabbits 8, and rats 4,9,11, was recently adapted for use in mice 5,6,10. In these studies, the intracameral injection of polystyrene microbeads, alone or in combination with a viscoelastic material, resulted in IOP elevation leading to subsequent RGC death 6,10. However, reflux when the needle is withdrawn from the eye and dislodging of microbeads from the iridocorneal angle are common problems that arise during the procedure. To minimize these drawbacks, magnets have been used to attract the magnetic microbeads to the iridocorneal angle of the eye 4,9.
The protocol described here is a modified procedure based on previous studies 9,10 that uses magnetic microbeads and a handheld magnet adapted to the mouse eye (Figure 1). Several important modifications have been introduced in our protocol to ensure effective and reproducible IOP increase in mice. First, the injection of microbeads is done using a carefully prepared glass microneedle with a facetted bevel. The resulting smooth surfaces of the microneedle as well as its sharpened tip ensures that minimal damage is inflicted as it punctures the cornea. The use of this glass microneedle also results in increased control when the microneedle tip enters the anterior chamber, thereby reducing the risk of damaging nearby structures such as the iris and the lens. In addition, the tiny injection lesion facilitates corneal self-repair and reduces unwanted injury-related effects.
Second, the injection of magnetic microbeads and the use of a handheld magnet allow precise control to attract the beads to the iridocorneal angle in the small mouse eye. Magnetic microbeads that are 4.5 &#181;m in diameter were used because this microbead size did not clog the prepared microneedle opening and importantly, once injected, these microbeads effectively blocked the drainage of aqueous humor. This approach not only reduces reflux of the injected microbeads, but also ensures that a maximum number of microbeads accumulates in the target area to effectively block aqueous humour drainage. Furthermore, this strategy also reduces the number of beads floating in the anterior chamber avoiding contact with other structures, such as the iris and the lens, and preventing passage to the posterior chamber. Collectively, these modifications ensure that the microbead injection surgery is performed with relative ease and in a timely manner resulting in a highly reproducible, effective, and sustained induction of ocular hypertension in mice.

<table>
	<tbody>
		<tr>
			<td>Puller</td>
			<td>Narishige</td>
			<td>PC-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Thin Wall Glass Capillaries</td>
			<td>World Precision Instruments</td>
			<td>TW150F-4</td>
			<td>Capillary has an outer diameter of 1.5 mm and inner diameter of 1.12 mm</td>
		</tr>
		<tr>
			<td>Stereo Microscope</td>
			<td>Zeiss</td>
			<td>MZ9.5</td>
			<td>Zoom factor range of 2.5 to 6.0. Microscope used for needle-making and the micro-bead injection surgery.</td>
		</tr>
		<tr>
			<td>Footswitch</td>
			<td>Linemaster</td>
			<td>T-91-SE</td>
			<td></td>
		</tr>
		<tr>
			<td>Stainless Steel Blade</td>
			<td>Feather</td>
			<td>No. 11</td>
			<td></td>
		</tr>
		<tr>
			<td>Microelectrode Beveler</td>
			<td>Science Products</td>
			<td>BV-10</td>
			<td></td>
		</tr>
		<tr>
			<td>Aerosol Duster</td>
			<td>Fisher</td>
			<td>23-022-523</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Hydroxide</td>
			<td>Fisher Scientific</td>
			<td>BP359-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris Base</td>
			<td>Fisher Scientific</td>
			<td>BP152-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Vortex</td>
			<td>Fisher Scientific</td>
			<td>12-812</td>
			<td></td>
		</tr>
		<tr>
			<td>Dynabeads M-450 Epoxy</td>
			<td>Life Technologies</td>
			<td>14011</td>
			<td>Magnetic beads are 4.5 &#181;m in diameter. Stock solution is at a concentration of 4 x 108 beads/mL. Store at 4&#176;C.</td>
		</tr>
		<tr>
			<td>Mini-Tube Rotators</td>
			<td>Fisher Scientific</td>
			<td>05-450-127</td>
			<td></td>
		</tr>
		<tr>
			<td>3 Handheld Magnets</td>
			<td>Geomag</td>
			<td></td>
			<td>0.45 Tesla. Magnet used for microbead preparation and microbead injection surgery.</td>
		</tr>
		<tr>
			<td>25 mL serological pipet</td>
			<td>Costar</td>
			<td>4489</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipet</td>
			<td>Drummond</td>
			<td>4-000-101</td>
			<td></td>
		</tr>
		<tr>
			<td>Biological Containment Hood</td>
			<td>Biostad</td>
			<td>377355</td>
			<td></td>
		</tr>
		<tr>
			<td>Balanced salt solution (BSS)</td>
			<td>Alcon</td>
			<td>0065-0800-25</td>
			<td></td>
		</tr>
		<tr>
			<td>P1000 Micropipet</td>
			<td>Gilson</td>
			<td>F123602</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microtube 1.5 mL</td>
			<td>Sarstedt</td>
			<td>72.690</td>
			<td></td>
		</tr>
		<tr>
			<td>P200 Micropipet</td>
			<td>Gilson</td>
			<td>F123601</td>
			<td></td>
		</tr>
		<tr>
			<td>0.2 mL PCR tube</td>
			<td>Sarstedt</td>
			<td>72737.002</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td></td>
			<td></td>
			<td>Controlled substance</td>
		</tr>
		<tr>
			<td>Xylazine</td>
			<td>Bayer Healthcare</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Acepromazine</td>
			<td>Vetoquinol</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>U-100 Insulin Syringe</td>
			<td>Becton Dickinson and Company</td>
			<td>329461</td>
			<td></td>
		</tr>
		<tr>
			<td>Balance</td>
			<td>Ohaus</td>
			<td>CS 200</td>
			<td></td>
		</tr>
		<tr>
			<td>Buprenorphine</td>
			<td></td>
			<td></td>
			<td>Controlled substance</td>
		</tr>
		<tr>
			<td>Tropicamide ophthalmic solution</td>
			<td>Alcon</td>
			<td>0998-0355-15</td>
			<td>1% Mydriacyl</td>
		</tr>
		<tr>
			<td>Manual Microsyringe Pump with Digital Display</td>
			<td>World Precision Instruments</td>
			<td>DMP</td>
			<td></td>
		</tr>
		<tr>
			<td>Manual Micromanipulator</td>
			<td>World Precision Instruments</td>
			<td>M3301R</td>
			<td></td>
		</tr>
		<tr>
			<td>Platform</td>
			<td>Fisher Scientific</td>
			<td>14-673-52</td>
			<td>8 x 8 inch</td>
		</tr>
		<tr>
			<td>Absorbent swabs</td>
			<td>Kettenbach</td>
			<td>30601</td>
			<td></td>
		</tr>
		<tr>
			<td>P20 Micropipet</td>
			<td>Gilson</td>
			<td>F123600</td>
			<td></td>
		</tr>
		<tr>
			<td>Plastic forcep</td>
			<td>Euroband</td>
			<td>1001</td>
			<td>Ensure forcep is plastic and has a flat surface to avoid damaging the eye</td>
		</tr>
		<tr>
			<td>Fluoroquinolone ophthalmic solution</td>
			<td>Alcon</td>
			<td></td>
			<td>Vigamox</td>
		</tr>
		<tr>
			<td>Heating pad</td>
			<td>Sunbeam</td>
			<td>E12107-834</td>
			<td></td>
		</tr>
		<tr>
			<td>Tonometer</td>
			<td>iCare</td>
			<td>TV02</td>
			<td>TONOLAB rebound tonometer&#160;</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehyde, Para</td>
			<td>Fisher Scientific</td>
			<td>T353-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Dissection tools</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Small brush</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Glutaraldehyde solution</td>
			<td>Sigma-Aldrich</td>
			<td>G7651</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Cacodylate, tryhydrate</td>
			<td>Canemco and Marivec</td>
			<td>124-65-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Brn-3a antibody (C-20)</td>
			<td>Santa Cruz Biotechnology</td>
			<td>sc-31984</td>
			<td></td>
		</tr>
		<tr>
			<td>Tissue Culture Plate, 48 well</td>
			<td>Falcon</td>
			<td>353078</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Fisher Scientific</td>
			<td>BP151-500</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey Serum</td>
			<td>Sigma-Aldrich</td>
			<td>D9663</td>
			<td></td>
		</tr>
		<tr>
			<td>Donkey anti-Goat IgG (H+L) Secondary Antibody, Alexa Fluor 594 conjugate</td>
			<td>Life Technologies</td>
			<td>A-11058</td>
			<td></td>
		</tr>
		<tr>
			<td>Aluminum foil</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope Slides</td>
			<td>Fisher Scientific</td>
			<td>12-550-15</td>
			<td></td>
		</tr>
		<tr>
			<td>Slow fade Gold antifade reagent</td>
			<td>Life Technologies</td>
			<td>S36936</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover Glass</td>
			<td>Fisher Scientific</td>
			<td>12-548-5E</td>
			<td></td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Osmium tetroxide 2% aqueous solution</td>
			<td>Electron Microscopy Sciences</td>
			<td>3294949</td>
			<td></td>
		</tr>
		<tr>
			<td>Embed-812</td>
			<td>Electron Microscopy Sciences</td>
			<td>14900</td>
			<td></td>
		</tr>
		<tr>
			<td>Dodecenyl succinic anhydride</td>
			<td>Electron Microscopy Sciences</td>
			<td>13710</td>
			<td></td>
		</tr>
		<tr>
			<td>Nadic methyl anhydride</td>
			<td>Electron Microscopy Sciences</td>
			<td>19000</td>
			<td></td>
		</tr>
		<tr>
			<td>DMP-30</td>
			<td>Electron Microscopy Sciences</td>
			<td>13600</td>
			<td></td>
		</tr>
		<tr>
			<td>Propylene oxide</td>
			<td>Sigma-Aldrich</td>
			<td>110205-1L</td>
			<td></td>
		</tr>
		<tr>
			<td>Embedding mold-Dykstra</td>
			<td>Electron Microscopy Sciences</td>
			<td>70907</td>
			<td></td>
		</tr>
		<tr>
			<td>Porter-Blum ultra-microtome</td>
			<td>Sorvall</td>
			<td>MT-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Toluidine blue O (Certified Biological Stain)</td>
			<td>Fisher-Scientific</td>
			<td>T161-25</td>
			<td></td>
		</tr>
	</tbody>
</table>

a magnetic microbead occlusion model to induce ocular hypertension-dependent glaucoma in mice

The use of rodent models of glaucoma has been essential to understand the molecular mechanisms that underlie the pathophysiology of this multifactorial neurodegenerative disease. With the advent of numerous transgenic mouse lines, there is increasing interest in inducible murine models of ocular hypertension. Here, we present an occlusion model of glaucoma based on the injection of magnetic microbeads into the anterior chamber of the eye using a modified microneedle with a facetted bevel. The magnetic microbeads are attracted to the iridocorneal angle using a handheld magnet to block the drainage of aqueous humour from the anterior chamber. This disruption in aqueous dynamics results in a steady elevation of intraocular pressure, which subsequently leads to the loss of retinal ganglion cells, as observed in human glaucoma patients. The microbead occlusion model presented in this manuscript is simple compared to other inducible models of glaucoma and also highly effective and reproducible. Importantly, the modifications presented here minimize common issues that often arise in occlusion models. First, the use of a bevelled glass microneedle prevents backflow of microbeads and ensures that minimal damage occurs to the cornea during the injection, thus reducing injury-related effects. Second, the use of magnetic microbeads ensures the ability to attract most beads to the iridocorneal angle, effectively reducing the number of beads floating in the anterior chamber avoiding contact with other structures (e.g., iris, lens). Lastly, the use of a handheld magnet allows flexibility when handling the small mouse eye to efficiently direct the magnetic microbeads and ensure that there is little reflux of the microbeads from the eye when the microneedle is withdrawn. In summary, the microbead occlusion mouse model presented here is a powerful investigative tool to study neurodegenerative changes that occur during the onset and progression of glaucoma.

The injection of magnetic microbeads into the anterior chamber of adult mice described in this protocol resulted in a robust and reproducible elevation of IOP. One week after the procedure, IOP increased from 10 &#177; 0.6 mm Hg (mean &#177; S.E.M.), the average baseline IOP in contralateral eyes, to 19 &#177; 0.5 mm Hg in hypertensive eyes (Student&#39;s t-test; ***p &#60;0.001, n = 12, Table 1, Figure 2). IOP stabilized thereafter and remained elevated at an average of 20 mm Hg for at least 6 weeks, the longest time-point examined in this study. The average peak IOP in microbead-injected eyes at 2, 3, and 6 weeks after surgery was 25 mm Hg. The vast majority of treated mice developed sustained high IOP, therefore this protocol does not require a second injection of microbeads.
To assess the time-course of RGC loss in this model, RGC soma were first quantified by immunostaining with Brn3a, a RGC-specific marker 17. The number of Brn3a-positive cells was quantified on flat-mounted retinas at 1, 2, 3 and 6 weeks after induction of ocular hypertension. Although a significant IOP elevation was detected as early as 1 week after microbead injection, no significant loss of RGC soma was observed within the first 2 weeks of the procedure (Figure 3). Substantial RGC death (22%), however, was evident at 3 weeks (2,430 &#177; 67 RGCs/mm2, mean &#177; S.E.M., n = 12) and 6 weeks (2,350 &#177; 74 RGCs/mm2, n = 10) post-induction of ocular hypertension, compared to intact control eyes from un-operated mice (3,141 &#177; 49 RGCs/mm2, n = 23) (ANOVA, p &#60;0.001).
Dysfunction and degeneration of RGC axons is a cardinal feature of glaucoma. Therefore, axonal loss was examined at 3 and 6 weeks after microbead injection by quantification of RGC axons in optic nerve cross-sections stained with toluidine blue (Figure 4). A substantial loss of RGC axons (25%) was observed at 3 weeks (28,401 &#177; 702 axons/nerve, mean &#177; S.E.M., n = 5) and 6 weeks (29,426 &#177; 948 axons/ nerve, n = 6) after injection of microbeads compared to intact optic nerves from un-operated eyes (39,467 &#177; 137 axons/nerve, n = 4) (ANOVA, p &#60;0.001). Collectively, these data demonstrate that injection of magnetic microbeads into the mouse anterior chamber leads to reproducible and sustained IOP elevation that results in RGC soma and axon degeneration.
<table>
	<tbody>
		<tr>
			<td colspan="2">Time after OHT surgery</td>
			<td>N</td>
			<td colspan="3">Mean IOP (mmHg) &#177; S.E.M</td>
			<td colspan="2">Peak IOP (mmHg)</td>
		</tr>
		<tr>
			<td></td>
			<td></td>
			<td></td>
			<td>Contralateral</td>
			<td>Glaucoma</td>
			<td>Difference</td>
			<td>Contralateral</td>
			<td>Glaucoma</td>
		</tr>
		<tr>
			<td colspan="2">1 week</td>
			<td>12</td>
			<td>10 &#177; 0.4</td>
			<td>19 &#177; 0.5</td>
			<td>9 &#177; 0.6</td>
			<td>12 &#177; 0.4</td>
			<td>22 &#177; 0.6</td>
		</tr>
		<tr>
			<td colspan="2">2 weeks</td>
			<td>13</td>
			<td>11 &#177; 0.5</td>
			<td>20 &#177; 0.8</td>
			<td>9 &#177; 0.5</td>
			<td>12 &#177; 0.9</td>
			<td>25 &#177; 0.7</td>
		</tr>
		<tr>
			<td colspan="2">3 weeks</td>
			<td>10</td>
			<td>11 &#177; 0.8</td>
			<td>20 &#177; 0.7</td>
			<td>10 &#177; 0.9</td>
			<td>13 &#177; 0.2</td>
			<td>25 &#177; 0.9</td>
		</tr>
		<tr>
			<td colspan="2">6 weeks</td>
			<td>12</td>
			<td>12 &#177; 0.5</td>
			<td>20 &#177; 0.6</td>
			<td>9 &#177; 0.7</td>
			<td>13 &#177; 0.5</td>
			<td>24 &#177; 0.6</td>
		</tr>
	</tbody>
</table>
Table 1. Elevation of Intraocular Pressure in the Murine Magnetic Microbead Occlusion Model. In awake female C57 BL/6 mice, IOP was measured using a calibrated rebound tonometer. Operated eyes displayed an increase in IOP detected at one week post-surgery that remained elevated for at least six weeks after the procedure.
<img alt="Figure 1" src="/files/ftp_upload/53731/53731fig1.jpg" /> 
Figure 1. Workflow of the Steps Involved in the Murine Magnetic Microbead Occlusion Model of Glaucoma. Step-by-step outline of all the procedures performed before, during, and after the surgery. <a href="https://www.jove.com/files/ftp_upload/53731/53731fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53731/53731fig2.jpg" /> 
Figure 2. Increase in Intraocular Pressure in the Murine Magnetic Microbead Occlusion Model. In awake female C57 BL/6 mice, IOP was measured using a calibrated rebound tonometer. The IOPs of microbead-injected eyes were significantly elevated at one week post-surgery (ANOVA, p &#60;0.001). The IOPs remained significantly elevated relative to the contralateral eye of injected mice for at least 6 weeks (ANOVA, p &#60;0.001). (Intact: n = 12; 1 week: n = 12, 2 weeks: n = 13, 3 weeks: n = 10, 6 weeks: n = 12). <a href="https://www.jove.com/files/ftp_upload/53731/53731fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/53731/53731fig3.jpg" /> 
Figure 3. Retinal Ganglion Cell Death in the Murine Magnetic Microbead Occlusion Model. RGCs were visualized by immunostaining of flat-mounted retinas using Brn3a in intact control retinas (A) and glaucomatous retinas at 3 and 6 weeks after microbead injection to induce ocular hypertension (OHT) (B, C). Scale bars: 20 &#181;m. (D) Quantitative analysis confirmed that microbead injection resulted in significant RGC soma loss at 3 and 6 weeks after the procedure compared to control eyes. The density of RGC soma in intact, non-glaucomatous C57/BL6 mice is shown as reference (white bars, 100% survival). Values are expressed as the mean &#177; S.E.M. (Intact: n = 23; 1 week: n = 6, 2 weeks: n = 6, 3 weeks: n = 12, 6 weeks: n = 10, ANOVA, *** p &#60;0.001). <a href="https://www.jove.com/files/ftp_upload/53731/53731fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" src="/files/ftp_upload/53731/53731fig4.jpg" /> 
Figure 4. Axonal Degeneration in the Murine Magnetic Microbead Occlusion Model. RGC axons were visualized by staining of optic nerve cross-sections with toluidine blue in intact control (A) and glaucomatous retinas at 3 and 6 weeks after microbead injection to induce ocular hypertension (OHT) (B, C). Scale bars: 10 &#181;m. (D) Quantitative analysis confirmed that microbead injection resulted in significant RGC axon loss at 3 and 6 weeks after the procedure compared to control eyes. The density of RGC axons in intact, non-glaucomatous C57/BL6 mice is shown as reference (white bars, 100% survival). Values are expressed as the mean &#177; S.E.M. (Intact: n = 4; 3 weeks: n = 5, 6 weeks: n = 6, ANOVA, *** p &#60;0.001). <a href="https://www.jove.com/files/ftp_upload/53731/53731fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about A Magnetic Microbead Occlusion Model to Induce Ocular Hypertension-Dependent Glaucoma in Mice at JoVE.com

A Magnetic Microbead Occlusion Model to Induce Ocular Hypertension-Dependent Glaucoma in Mice

Here, we present a protocol to induce ocular hypertension in the murine eye that results in the loss of retinal ganglion cells as observed in glaucoma. Magnetic microbeads are injected into the anterior chamber and attracted to the iridocorneal angle using a magnet to block the outflow of aqueous humour.

The use of rodent models of glaucoma has been essential to understand the molecular mechanisms that underlie the pathophysiology of this multifactorial ...

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19.2K Views.  CRCHUM and University of Montreal. Here, we present a protocol to induce ocular hypertension in the murine eye that results in the loss of retinal ganglion cells as observed in glaucoma. Magnetic microbeads are injected into the anterior chamber and attracted to the iridocorneal angle using a magnet to block the outflow of aqueous humour.

Video: A Magnetic Microbead Occlusion Model to Induce Ocular Hypertension-Dependent Glaucoma in Mice

Modeling Stroke in Mice - Middle Cerebral Artery Occlusion with the Filament Model

Stroke is among the most frequent causes of death and adult disability, especially in highly developed countries. However, treatment options to date are very limited. To meet the need for novel therapeutic approaches, experimental stroke research frequently employs rodent models of focal cerebral ischaemia. Most researchers use permanent or transient occlusion of the middle cerebral artery (MCA) in mice or rats.
Proximal occlusion of the middle cerebral artery (MCA) via the intraluminal suture technique (so called filament or suture model) is probably the most frequently used model in experimental stroke research. The intraluminal MCAO model offers the advantage of inducing reproducible transient or permanent ischaemia of the MCA territory in a relatively non-invasive manner. Intraluminal approaches interrupt the blood flow of the entire territory of this artery. Filament occlusion thus arrests flow proximal to the lenticulo-striate arteries, which supply the basal ganglia. Filament occlusion of the MCA results in reproducible lesions in the cortex and striatum and can be either permanent or transient. In contrast, models inducing distal (to the branching of the lenticulo-striate arteries) MCA occlusion typically spare the striatum and primarily involve the neocortex. In addition these models do require craniectomy. In the model demonstrated in this article, a silicon coated filament is introduced into the common carotid artery and advanced along the internal carotid artery into the Circle of Willis, where it blocks the origin of the middle cerebral artery. In patients, occlusions of the middle cerebral artery are among the most common causes of ischaemic stroke. Since varying ischemic intervals can be chosen freely in this model depending on the time point of reperfusion, ischaemic lesions with varying degrees of severity can be produced. Reperfusion by removal of the occluding filament at least partially models the restoration of blood flow after spontaneous or therapeutic (tPA) lysis of a thromboembolic clot in humans.
In this video we will present the basic technique as well as the major pitfalls and confounders which may limit the predictive value of this model.

Filamentous occlusion of the Middle cerebral artery is a common model for studying ischemic stroke in mice.

Stroke is among the most frequent causes of death and adult disability, especially in highly developed countries. However, treatment options to date are ...

Pressure Controlled Ventilation to Induce Acute Lung Injury in Mice

Murine models are extensively used to investigate acute injuries of different organs systems (1-34). Acute lung injury (ALI), which occurs with prolonged mechanical ventilation, contributes to morbidity and mortality of critical illness, and studies on novel genetic or pharmacological targets are areas of intense investigation (1-3, 5, 8, 26, 30, 33-36). ALI is defined by the acute onset of the disease, which leads to non-cardiac pulmonary edema and subsequent impairment of pulmonary gas exchange (36). We have developed a murine model of ALI by using a pressure-controlled ventilation to induce ventilator-induced lung injury (2). For this purpose, C57BL/6 mice are anesthetized and a tracheotomy is performed followed by induction of ALI via mechanical ventilation. Mice are ventilated in a pressure-controlled setting with an inspiratory peak pressure of 45 mbar over 1 - 3 hours. As outcome parameters, pulmonary edema (wet-to-dry ratio), bronchoalveolar fluid albumin content, bronchoalveolar fluid and pulmonary tissue myeloperoxidase content and pulmonary gas exchange are assessed (2). Using this technique we could show that it sufficiently induces acute lung inflammation and can distinguish between different treatment groups or genotypes (1-3, 5). Therefore this technique may be helpful for researchers who pursue molecular mechanisms involved in ALI using a genetic approach in mice with gene-targeted deletion.

A murine model for ventilator induced lung injury is an important tool to study an acute lung injury in vivo. Here, we report an easy applicable in situ model for acute lung injury using high-pressure mechanical ventilation to induce acute failure of the lung.

Murine models are extensively used to investigate acute injuries of different organs systems (1-34). Acute lung injury (ALI), which occurs with prolonged ...

Use of a Hanging Weight System for Coronary Artery Occlusion in Mice

Murine studies of acute injury are an area of intense investigation, as knockout mice for different genes are becoming increasingly available 1-38. Cardioprotection by ischemic preconditioning (IP) remains an area of intense investigation. To further elucidate its molecular basis, the use of knockout mouse studies is particularly important 7, 14, 30, 39. Despite the fact that previous studies have already successfully performed cardiac ischemia and reperfusion in mice, this model is technically very challenging. Particularly, visual identification of the coronary artery, placement of the suture around the vessel and coronary occlusion by tying off the vessel with a supported knot is technically difficult. In addition, re-opening the knot for intermittent reperfusion of the coronary artery during IP without causing surgical trauma adds additional challenge. Moreover, if the knot is not tied down strong enough, inadvertent reperfusion due to imperfect occlusion of the coronary may affect the results. In fact, this can easily occur due to the movement of the beating heart.
Based on potential problems associated with using a knotted coronary occlusion system, we adopted a previously published model of chronic cardiomyopathy based on a hanging weight system for intermittent coronary artery occlusion during IP 39. In fact, coronary artery occlusion can thus be achieved without having to occlude the coronary by a knot. Moreover, reperfusion of the vessel can be easily achieved by supporting the hanging weights which are in a remote localization from cardiac tissues.
We tested this system systematically, including variation of ischemia and reperfusion times, preconditioning regiments, body temperature and genetic backgrounds39. In addition to infarct staining, we tested cardiac troponin I (cTnI)
as a marker of myocardial infarction in this model. In fact, plasma levels of cTnI correlated with infarct sizes (R2=0.8). Finally, we could show in several studies that this technique yields highly reproducible infarct sizes during murine IP and myocardial infarction6, 8, 30, 40, 41. Therefore, this technique may be helpful for researchers who pursue molecular mechanisms involved in cardioprotection by IP using a genetic approach in mice with targeted gene deletion. Further studies on cardiac IP using transgenic mice may consider this technique.

A murine model for myocardial ischemia and ischemic preconditioning is an important tool study cardioprotective mechanisms in vivo. Here, we report an easy applicable in situ model for cardiac IP using a hanging-weight system for coronary artery occlusion.

Murine studies of acute injury are an area of intense investigation, as knockout mice for different genes are becoming increasingly available 1-38. ...

The CYP2D6 Animal Model: How to Induce Autoimmune Hepatitis in Mice

Autoimmune hepatitis is a rare but life threatening autoimmune disease of the liver of unknown etiology1,2. In the past many attempts have been made to generate an animal model that reflects the characteristics of the human disease 3-5. However, in various models the induction of disease was rather complex and often hepatitis was only transient3-5. Therefore, we have developed a straightforward mouse model that uses the major human autoantigen in type 2 autoimmune hepatitis (AIH-2), namely hCYP2D6, as a trigger6. Type 1 liver-kidney microsomal antibodies (LKM-1) antibodies recognizing hCYP2D6 are the hallmark of AIH-27,8. Delivery of hCYP2D6 into wildtype FVB or C57BL/6 mice was by an Adenovirus construct (Ad-2D6) that ensures a direct delivery of the triggering antigen to the liver. Thus, the ensuing local inflammation generates a fertile field9 for the subsequent development of autoimmunity. A combination of intravenous and intraperitoneal injection of Ad-2D6 is the most effective route to induce a long-lasting autoimmune damage to the liver (section 1). Here we provide a detailed protocol on how autoimmune liver disease is induced in the CYP2D6 model and how the different aspects of liver damage can be assessed. First, the serum levels of markers indicating hepatocyte destruction, such as aminotransferases, as well as the titers of hCYP2D6 antibodies are determined by sampling blood retroorbitaly (section 2). Second, the hCYP2D6-specific T cell response is characterized by collecting lymphocytes from the spleen and the liver. In order to obtain pure liver lymphocytes, the livers are perfused by PBS via the portal vein (section 3), digested in collagen and purified over a Percoll gradient (section 4). The frequency of hCYP2D6-specific T cells is analyzed by stimulation with hCYP2D6 peptides and identification of IFN&gamma;-producing cells by flow cytometry (section 5). Third, cellular infiltration and fibrosis is determined by immunohistochemistry of liver sections (section 6). Such analysis regimen has to be conducted at several times after initiation of the disease in order to prove the chronic nature of the model. The magnitude of the immune response characterized by the frequency and activity of hCYP2D6-specific T and/or B cells and the degree of the liver damage and fibrosis have to be assessed for a subsequent evaluation of possible treatments to prevent, delay or abrogate the autodestructive process of the liver.

Infection of mice with an Adenovirus expressing the major human autoantigen cytochrome P450 2D6 (hCYP2D6) recognized by sera of patients suffering from type 2 autoimmune hepatitis results in a persistent form of autoimmune-mediated liver disease characterized by extensive hepatitis, fibrosis and generation of a CYP2D6-specific immune response.

Autoimmune hepatitis is a rare but life threatening autoimmune disease of the liver of unknown etiology1,2. In the past many attempts have been made to ...

Bilateral Common Carotid Artery Occlusion as an Adequate Preconditioning Stimulus to Induce Early Ischemic Tolerance to Focal Cerebral Ischemia

There is accumulating evidence, that ischemic preconditioning - a non-damaging ischemic challenge to the brain - confers a transient protection to a subsequent damaging ischemic insult. We have established bilateral common carotid artery occlusion as a preconditioning stimulus to induce early ischemic tolerance to transient focal cerebral ischemia in C57Bl6/J mice. In this video, we will demonstrate the methodology used for this study.

There is accumulating evidence, that ischemic preconditioning (PC) &#x2013; a non-damaging ischemic challenge to the brain - confers a transient protection to a subsequent damaging ischemic insult. We established bilateral common carotid artery occlusion (BCCAO) as a preconditioning stimulus to induce early ischemic tolerance (IT) to transient focal cerebral ischemia (induced by middle cerebral artery occlusion, MCAO) in C57Bl6/J mice.

There is accumulating evidence, that ischemic preconditioning - a non-damaging ischemic challenge to the brain - confers a transient protection to a ...

Reproducable Paraplegia by Thoracic Aortic Occlusion in a Murine Model of Spinal Cord Ischemia-reperfusion

Background 
Lower extremity paralysis continues to complicate aortic interventions. The lack of understanding of the underlying pathology has hindered advancements to decrease the occurrence this injury.&#160;The current model demonstrates reproducible lower&#160;extremity paralysis following thoracic aortic occlusion.
Methods 
Adult male C57BL6 mice were anesthetized with isoflurane. Through a cervicosternal incision the aorta was exposed. The descending thoracic aorta and left subclavian arteries were identified without entrance into pleural space. Skeletonization of these arteries was followed by immediate closure (Sham) or occlusion for 4 min (moderate ischemia) or 8 min (prolonged ischemia). The sternotomy and skin were closed and the mouse was transferred to warming bed for recovery.&#160; Following recovery, functional analysis was obtained at 12 hr intervals until 48 hr.
Results 
Mice that underwent sham surgery showed no observable hind&#160;limb deficit. Mice subjected to moderate ischemia for 4 min had minimal functional deficit at 12 hr followed by progression to complete paralysis at 48 hr. Mice subjected to prolonged ischemia had an immediate paralysis with no observable hind-limb movement at any point in the postoperative period. There was no observed intraoperative or post&#160;operative mortality.
Conclusion 
Reproducible lower extremity paralysis whether immediate or delayed can be achieved in a murine model. Additionally, by using a median sternotomy and careful dissection, high survival rates, and reproducibility can be achieved.

The lack of mechanistic understanding of spinal cord ischemia-reperfusion injury has hindered further adjuncts to prevent paraplegia following high risk aortic operations. Thus, the development of animal models is imperative. This manuscript demonstrates reproducible lower extremity paralysis following thoracic aortic occlusion in a murine model.

Background
Lower extremity paralysis continues to complicate aortic interventions. The lack of understanding of the underlying pathology has hindered ...

Experimental Glaucoma Induced by Ocular Injection of Magnetic Microspheres

Progress in understanding the pathophysiology, and providing novel treatments for glaucoma is dependent on good animal models of the disease. We present here a protocol for elevating intraocular pressure (IOP) in the rat, by injecting magnetic microspheres into the anterior chamber of the eye. The use of magnetic particles allows the user to manipulate the beads into the iridocorneal angle, thus providing a very effective blockade of fluid outflow from the trabecular meshwork. This leads to long-lasting IOP rises, and eventually neuronal death in the ganglion cell layer (GCL) as well as optic nerve pathology, as seen in patients with the disease. This method is simple to perform, as it does not require machinery, specialist surgical skills, or many hours of practice to perfect. Furthermore, the pressure elevations are very robust, and reinjection of the magnetic microspheres is not usually required unlike in some other models using plastic beads. Additionally, we believe this method is suitable for adaptation for the mouse eye.

We present a method for inducing elevated intraocular pressure (IOP), by injecting magnetic microspheres into the rat eye, to model glaucoma. This leads to strong pressure rises, and extensive neuronal death. This protocol is easy to perform, does not require repeat injections, and produces stable long-lasting IOP rises.

Progress in understanding the pathophysiology, and providing novel treatments for glaucoma is dependent on good animal models of the disease. We present ...

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 ...

Surgical Approach for Middle Cerebral Artery Occlusion and Reperfusion Induced Stroke in Mice

Stroke is a leading cause of death worldwide and continues to be one of the major causes of long-term adult disabilities. About 87% of strokes are ischemic in origin and occur in the territory of the middle cerebral artery (MCA). Currently the only Food and Drug Administration (FDA) approved drug for the treatment of this devastating disease is tissue plasminogen activator (tPA). However, tPA has a small therapeutic window for administration (3 - 6 hr), and is only effective in 4% of the patients who actually receive it. Current research focuses on understanding the pathophysiology of stroke in order to find potential therapeutic targets. Thus, reliable models are crucial, and the MCA occlusion (MCAo) model (also termed the intraluminal filament or suture model) is deemed to be the most clinically relevant surgical model of ischemic stroke, and is fairly non-invasive and easily reproducible. Typically the MCAo model is used with rodents, especially with mice due to all the genetic variations available for this species. Here we describe (and present in the video) how to successfully perform the MCAo model (with reperfusion) in mice to generate reliable and reproducible data.

In order to understand the pathophysiology of stroke, it is important to use reliable models. This paper will describe one of the most frequently used stroke models in mice, termed the middle cerebral artery occlusion (MCAo) model (also termed the intraluminal filament or suture model) with reperfusion.

Stroke is a leading cause of death worldwide and continues to be one of the major causes of long-term adult disabilities. About 87% of strokes are ...

A Closed-chest Model to Induce Transverse Aortic Constriction in Mice

Research on cardiac hypertrophy and heart failure is frequently based on pressure overload mouse models induced by TAC. The standard procedure is to perform a partial thoracotomy to visualize the transverse aortic arch. However, the surgical trauma caused by the thoracotomy in open-chest models changes the respiratory physiology as the ribs are dissected and left unattached after chest closure. To prevent this, we established a minimally invasive, closed chest approach via lateral thoracotomy. Herein we approach the aortic arch via the 2nd intercostal space without entering the chest cavities, leaving the mouse with a less traumatic injury to recover from. We perform this operation using standard laboratory settings for open chest TAC procedures with equal survival rates. Apart from maintaining physiological breathing patterns due to the closed chest approach, the mice seem to benefit by showing rapid recovery, as the less invasive technique appears to facilitate a fast healing process and to reduce immune response after trauma.

Here, we present a protocol of Transverse Aortic constriction (TAC) via a lateral thoracotomy. This technique is a minimally invasive, closed chest surgical procedure aiming to simulate pressure overload and heart failure in mice utilizing standard TAC laboratory settings.

Research on cardiac hypertrophy and heart failure is frequently based on pressure overload mouse models induced by TAC. The standard procedure is to ...