Supported by R&#252;diger Foundation grants in 2008 &#38; 2010 (BVS), BONFOR/Gerok Scholarship O-137.0015 (BVS), BONFOR/Gerok Scholarship O-137.0019 (FT), Deutsche Forschungsgemeinschaft/ DFG (BVS) STA 1135/2-1, Chinese Scholarship Council No. 2008627116 (ZL) and an unrestricted grant by Geuder AG, Heidelberg (Fig. 2). Members of H. Skottman&#39;s laboratory, University of Tampere Finland are gratefully acknowledged for providing hES derived RPE shown in Fig 2.&#160;

Ethics of animal handling in ophthalmic research: We obtained approval from the Ethics committee of the Medical Faculty, University of Bonn, and adhere to the guidelines stated by The Association for Research in Vision and Ophthalmology (ARVO). Moreover, all procedures were approved by the state regulatory authorities of North Rhine-Westphalia. Animals were held indoors in a specialized facility in an air-conditioned room with temperatures between 18 - 20 &#176;C, exposure to regular daylight, in standardized individual cages with free access to food and water.

Note: To ensure the animals operative affinity, an animal health score sheet is followed which includes the following definitive animal exclusion criteria: 20% weight loss compared to weight on admission; apparent cyanosis of the animal; animal shivers, has cramps or cannot move in coordination; ataxia/ paresthesia, e.g., paralyses; apathy; extreme auto mutilation (skin wounds, severed limbs).

1. Instrument Sterilization
<ol>
	<li>Place reusable instruments in ultrasonic bath.</li>
	<li>Add 500 ml distilled water and 2 ml of instrument disinfectant.</li>
	<li>Clean instruments using sweep function for 15 min.</li>
	<li>Remove instruments from ultrasonic bath and rinse thoroughly using distilled water for 5 min.</li>
	<li>Insert instruments in autoclave and use the standard program (sterilization of instruments under 121 &#186;C for 20 min).</li>
</ol>
2. Instrument Preparation
<ol>
	<li>Establish and maintain a sterile field, by working in a closed room, wearing surgical scrubs, mask and hair cover. Disinfect hands prior to wearing sterile surgical gloves. For detailed approach see 28.</li>
	<li>Place sterilized instruments on a sterile drape.</li>
	<li>Place 1 ml syringe filled with 40 mg triamcinolone attached to a 27 G needle for injection, 10 ml syringe with Balance salt solution (BSS), and 5 ml syringe of lubricant on drape.</li>
	<li>Place 3-0 silk, 7-0 vicryl, ocular sticks (to stop conjunctival/ scleral bleeding), twister gauze sponges, wound closure strips (to fixate the vitrectomy tip tubing), and chandelier endoillumination fiber wire on a drape.</li>
	<li>Unwrap 25 G chandelier endoilluminator and connect to light machine using sterile techniques (see step 2.1). Connect vitrectomy set including high speed vitrector and Venturi cassette to vitrectomy machine using sterile techniques (see step 2.1).</li>
	<li>Open 500 ml BSS bottle and connect solution to Venturi cassette according manufacturer&#39;s instructions.</li>
</ol>
3. Preparation of Anesthesia and Positioning of the Animal
<ol>
	<li>Weigh animal to ensure accurate medication dosage.</li>
	<li>Prepare intramuscular (IM) anesthesia using 1 syringe with 27 G needle containing 0.35 mg/kg ketamine and 0.25 mg/kg medetomidine for start. Flip the syringe to mix.</li>
	<li>Prepare 2 syringes with 1/3 the doses to maintain the anesthesia during the operation.</li>
	<li>Prepare syringe containing 20 ml of 5% glucose solution and 18 G needle for subcutaneous injection as an intravenous infusion alternative.</li>
	<li>Give 3 x 1 drop of mydriatic eye drops prior to vitrectomy for pupil dilatation.</li>
	<li>Cover rabbit with blanket to calm before anesthesia injection, inject in the hind limb (gluteal muscle) and massage around injection site for 30 sec. 
	Note: The first shot of IM anesthesia lasts about 1 - 2 hr. depending on the rabbit&#39;s size, drug tolerance, fat layer, stress and body temperature. The first sign of the anesthesia fading away is a nystagmus (must be monitored by the surgeon), subsequent injections last about 30 - 45 min.</li>
	<li>Confirm proper anesthesia, by verifying hypnosis, hyporeflexia, analgesia and muscle relaxation of the animal.</li>
	<li>Give subcutaneous injection (Pt. 3.3) in neck skin fold, once the rabbit is unconscious.</li>
	<li>Add methylcellulose lubricant every 5 - 10 min in operated eye, add lubricant and tape lids in non-operated eye.</li>
	<li>Place the covered rabbit on the surgical table draped with a cover such as a cotton blanket in an optimal position (nose slightly elevated through a mold of the blanket, so it is level with ocular surface) under surgical microscope. Align eye perpendicular to microscope objective.</li>
	<li>Ensure proper body core temperature using rectal thermometer (normothermia 39 &#177;&#160;1 &#186;C)29.</li>
	<li>Cut eyelashes using scissors (some ointment on blade) to reduce postoperative infections.</li>
	<li>Disinfect the eye using 2 - 3 drops of 0.1 g/ml Povidone-Iodine topically for 1 min and rinse with sterile BSS.</li>
	<li>Cover eye with sterile drape with pre-cut opening in the middle for the eye and then cover with (sticky) surgical incision drape 12 x 17 cm.</li>
</ol>
4. Vitrectomy
<ol>
	<li>Proptose and secure eye with 3-0 silk using inverted caliper and, perform a conjunctival peritomy.
	<ol>
		<li>Incise the conjunctiva with a vannas scissor close to limbus but far enough from the blood vessels (~1 mm distance).</li>
		<li>Dissect the conjunctiva by creating a &#34;T-cut&#34;. First enlarge the peritomy with the scissor parallel to limbus and then incise the conjunctiva vertically in form of a &#34;T&#34; for about 6 - 7 mm. Carefully separate the conjunctiva bluntly.</li>
	</ol>
	</li>
	<li>Perform a sclerotomy using a 23 G microvitreoretinal (MVR) blade at 8 o&#39;clock on right eye/OD (4 o&#39;clock on left eye/OS) by carefully inserting the sharp tip of the blade in direction towards the optic nerve. Slowly retract the blade in the same direction and avoid enlarging the sclerotomy.</li>
	<li>Insert and suture custom side port-infusion cannula27 using 7-0 silk suture and set intraocular pressure (IOP) at 24 mmHg.</li>
	<li>Perform a sclerotomy with 25 G flat head trocar at 2 o&#39;clock on OD (10 o&#39;clock on OS) similar to step 4.2.</li>
	<li>Insert 25 G chandelier light into flat head trocar, fixate with sticky tape and turn on light source at ca. 30%.</li>
	<li>If needed, remove edematous corneal epithelium using a #20 scalpel for better intraocular visualization.</li>
	<li>Perform a sclerotomy similar to step 4.2 at 10 o&#39;clock on OD (2 o&#39;clock on OS), (pre-) place u-shaped 7-0 sutures around sclerotomy without tying the knot, and insert vitrectomy cutter tip.</li>
	<li>Start vitrectomy30 around the entry port, then continue over the optic disc and the fibrae medullares using high speed vitrector by cutting the vitreous humor into small pieces at max. 2,000 - 3,000 cuts/min, aspirating at max. 200 mmHg using the stated parameter setup of the vitrectomy machine (Table 1)</li>
	<li>Perform a posterior vitreous detachment (PVD) by separating the vitreous humor from the retina by holding the high speed vitrector over the posterior pole and (if feasible gently) superior of the disc31 while aspirating only at max. 200 mmHg without cutting.</li>
	<li>Inject ca. 50 &#181;l (20 mg) triamcinolone or diluted fluorescein (ca. 0.1 mg/ml) intravitreally to visualize and facilitate (near total) removal of the floating vitreous over the posterior pole and midperiphery during vitrectomy. Avoid crossing over under the lens. Indentation to shave peripheral vitreous by a (skilled) assistant is recommended if gas tamponade is desired.</li>
	<li>Add 20 units/ml heparin and 0.5 mg epinephrine to final concentration of 0.001 mg/ml into the BSS infusion solution in parallel or after step 4.10. 
	Note: As heparin/epinephrine are not injected intraocular their effects are delayed depending on infusion flow rate.</li>
</ol>
5. Loading Shooter
Note: The work described herein does not fall under the tenets of the Declaration of Helsinki; it did not involve human patients. Here, standard RPE cells were isolated from fetal human eyes, cultured and differentiated on uncoated 10-&#181;m-thick polyester (PET) inserts according to our previously published protocol14. A permission to work with the human fetal material was obtained from the ethics committee of the University of Bonn. Alternatively, hES-RPE were shipped from the Skottman lab (manuscript in prep.), where they were cultured according to the technique described by Vaajasaari et al.32; for these cells a permission has been obtained from the R. Koch Institute, Berlin, Germany.
<ol>
	<li>Rinse cell culture prior to preparation of the implant 3x with ophthalmic grade BSS.</li>
	<li>Fill a standard cell culture dish (100 x 20 mm) with 10 ml ophthalmic grade BSS.</li>
	<li>Add the cell culture insert into the BSS and center the dish under a light microscope.</li>
	<li>Punch out a 2.4 x 1.1 mm implant with a blunt, oval, custom-made needle to obtain a flat, bean-shaped substrate with two long edges and two round edges.</li>
	<li>Gently flood the needle through the second port with BSS to flush out the implant into the BSS filled custom made loading station (Figure 1).</li>
	<li>Optionally cut one round end of implant (&#60;0.5 mm), just to obtain a third edge.</li>
	<li>Make sure that the implant is in the right orientation by ensuring that the monolayer is upside on the cell carrier. To change positioning carefully use two scalpels.</li>
	<li>Push the implant gently and completely into the shooter instrument using the needle holder until all of the implant is secured inside of the tip. The plunger should remain retracted.</li>
	<li>Keep the &#34;loaded&#34; shooter tip in loading station under BSS until the moment of implantation.</li>
</ol>
6. Implantation
<ol>
	<li>Approach neural retina with extendible 41 G subretinal injection needle connected to a gastight syringe (ensure that all air bubbles have been evacuated from tubing!).</li>
	<li>Inject BSS (with calcium and magnesium/ CM) subretinally and thereby create a bleb retinal detachment (bRD) of about 2 - 3 disc diameter (DD). Two bRD per eye can be raised safely.</li>
	<li>Enlarge retinotomy to 1.5 mm with vertical 23 G VR-scissors. The subretinal space is now accessible for implantation or further maneuvering.</li>
	<li>Extend sclerotomy (precisely) with a 1.4 mm incision knife to 20 G approach.</li>
	<li>Attempt passing through the sclerotomy using a 20 G shooter dummy, enlarge as needed to ensure smooth, yet snug transition of the loaded shooter.</li>
	<li>Pass with the loaded shooter27 through sclerotomy ideally at 24 mmHg.</li>
	<li>Approach retinotomy edge and eject the implant subretinally from an epiretinal position.</li>
	<li>Adjust the implant with half-closed 23 G scissors, forceps or 41 G needle to make sure it is positioned well under the retina- reasonably away from the retinotomy.</li>
</ol>
7. Ending Operation
<ol>
	<li>Remove 25 G chandelier and infusion cannula.</li>
	<li>Suture all sclerotomies.</li>
	<li>Inject 25 &#181;l (10 mg) triamcinolone by the 8 o&#39;clock sclerotomy (prior to suturing last sclerotomy).</li>
	<li>Check/adjust IOP by palpation and inject BSS via 30 G needle/syringe, if needed.</li>
	<li>Suture conjunctiva with 7-0 vicryl.</li>
	<li>Remove proptosing 3-0 silk sling slowly (Avoid deep orbital venous plexus!).</li>
	<li>Add dexamethasone/antibiotic ointment under lid.</li>
	<li>Position for 1 hr covered with blanket with operated eye facing up (w/o gas), or down (with air/gas).</li>
	<li>Do not leave animal unattended until it regains sufficient consciousness to maintain sternal recumbence.</li>
	<li>Do not transport rabbit before the anesthesia fades completely, this can be speeded up by injecting medetomidine reversing agent equal to amount of medetomidine given.</li>
</ol>
8. Post-operative Animal Care
<ol>
	<li>Keep rabbits under appropriate conditions (temperature, light, food, water, space, etc.) and close monitoring in a specialized facility.</li>
	<li>Ensure that animal is well rested, i.e., no extended periods of food or water deprivation.</li>
	<li>Look for any wounds or injuries, especially on injection sites.</li>
	<li>Keep wounds dry to prevent infections. Give antibiotics when infection is suspected: dexamethasone 1 mg/g, neomycin sulfate 3,500 I.U./g, polymyxin B sulfate 6,000 I.U./g ointment was applied twice daily for 1 week postoperative onto the ocular surface.</li>
	<li>Add dexamethasone/antibiotic ointment for next 7 days postoperative twice daily for better ocular surface regeneration and reduced post-operative pain.</li>
	<li>Give systemic analgesics (Carprofene 4 mg/kg twice daily) for initial 48 hr.</li>
	<li>Do not leave an animal unattended until it has regained sufficient consciousness to maintain sternal recumbence.</li>
	<li>Do not return an animal that has undergone surgery to the company of other animals until fully recovered.</li>
	<li>Do not expose animals to unnecessary distress.</li>
</ol>
9. SLO/SD-OCT Guidance
<ol>
	<li>Prepare and inject intramuscular (IM) anesthesia using 1 syringe with 27 G needle containing 0.175 mg/kg ketamine and 0.125 mg/kg medetomidine for start. Flip the syringe to mix.</li>
	<li>Add a lubricant at least every 5 min to moisturize eye and maintain clear SD-OCT imaging, optionally a custom contact lens may be used.</li>
	<li>Attach a steel platform to the headrest to stabilize the animal in the required position.</li>
	<li>Place rabbit on steel platform, placing its eye perpendicular to probe.</li>
	<li>Hold the rabbit&#39;s head from the lower cheek bone (mandibula) avoiding the trachea.</li>
	<li>Tilt animal&#39;s head by roughly 45&#186; towards the SD-OCT probe to optimize the viewing angle on the implant.</li>
	<li>Use a 30-degree lens and the following parameters for optimal OCT imaging [HS]: 30 degrees settings for single line scans with ART mode set to 100 (averaging) and 20 x 20 degree settings for volume scans with ART mode set to 15; high resolution mode is not required.</li>
	<li>Use SLO infrared reflectance imaging to find the focal plane of the implant (Fig. 2A); optimal focus is reached when (all) the implant edges are sharp14.</li>
	<li>Add dexamethasone 1 mg/g, neomycin sulfate 3,500 I.U./g, polymyxin B sulfate 6,000 I.U./g ointment under lid when finished.</li>
</ol>

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<li>Hong, S. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Physiologic+characteristics+of+cold+perfluorocarbon-induced+hypothermia+during+partial+liquid+ventilation+in+normal+rabbits%5BTitle%5D)+AND+%22Anesth+Analg%22%5BJournal%5D)">Physiologic characteristics of cold perfluorocarbon-induced hypothermia during partial liquid ventilation in normal rabbits.</a> Anesth Analg. 94, 157-162 (2002).</li>
<li>Machemer, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+development+of+pars+plana+vitrectomy%3A+a+personal+account%5BTitle%5D)+AND+%22Graefes+Arch+Clin+Exp+Ophthalmol%22%5BJournal%5D)">The development of pars plana vitrectomy: a personal account.</a> Graefes Arch Clin Exp Ophthalmol. 233, 453-468 (1995).</li>
<li>Los, L. I., van Luyn, M. J., Nieuwenhuis, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Organization+of+the+rabbit+vitreous+body%3A+lamellae%2C+Cloquet%27s+channel+and+a+novel+structure%2C+the+%27alae+canalis+Cloqueti%27%5BTitle%5D)+AND+%22Exp+Eye+Res%22%5BJournal%5D)">Organization of the rabbit vitreous body: lamellae, Cloquet's channel and a novel structure, the 'alae canalis Cloqueti'.</a> Exp Eye Res. 69, 343-350 (1999).</li>
<li>Vaajasaari, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Toward+the+defined+and+xeno-free+differentiation+of+functional+human+pluripotent+stem+cell-derived+retinal+pigment+epithelial+cells%5BTitle%5D)+AND+%22Mol+Vis%22%5BJournal%5D)">Toward the defined and xeno-free differentiation of functional human pluripotent stem cell-derived retinal pigment epithelial cells.</a> Mol Vis. 17, 558-575 (2011).</li>
<li>Iverson, D. A., Katsura, H., Hartzer, M. K., Blumenkranz, M. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Inhibition+of+intraocular+fibrin+formation+following+infusion+of+low-molecular-weight+heparin+during+vitrectomy%5BTitle%5D)+AND+%22Arch+Ophthalmol%22%5BJournal%5D)">Inhibition of intraocular fibrin formation following infusion of low-molecular-weight heparin during vitrectomy.</a> Arch Ophthalmol. 109, 405-409 (1991).</li>
<li>Thieltges, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Subretinal+implantation+of+human+embryonic+stem+cell+derived+RPE+on+ultrathin+polyester+carriers+in+rabbits%5BTitle%5D)+AND+%22Investigative+Ophthalmology+%26+Visual+Science%22%5BJournal%5D)">Subretinal implantation of human embryonic stem cell derived RPE on ultrathin polyester carriers in rabbits.</a> Investigative Ophthalmology & Visual Science. 56, 1824(2015).</li>
<li>Kamao, H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Characterization+of+human+induced+pluripotent+stem+cell-derived+retinal+pigment+epithelium+cell+sheets+aiming+for+clinical+application%5BTitle%5D)+AND+%22Stem+Cell+Reports%22%5BJournal%5D)">Characterization of human induced pluripotent stem cell-derived retinal pigment epithelium cell sheets aiming for clinical application.</a> Stem Cell Reports. 2, 205-218 (2014).</li>
<li>Seiler, M. J., Aramant, R. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Intact+sheets+of+fetal+retina+transplanted+to+restore+damaged+rat+retinas%5BTitle%5D)+AND+%22Invest+Ophthalmol+Vis+Sci%22%5BJournal%5D)">Intact sheets of fetal retina transplanted to restore damaged rat retinas.</a> Invest Ophthalmol Vis Sci. 39, 2121-2131 (1998).</li>
<li>Stanzel, B. V., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((SD-OCT+Complements+Histology+in+Evaluation+of+Potential+Bruch%27s+Membrane+Prosthetics%5BTitle%5D)+AND+%22Invest+Ophthalmol+Vis+Sci%22%5BJournal%5D)">SD-OCT Complements Histology in Evaluation of Potential Bruch's Membrane Prosthetics.</a> Invest Ophthalmol Vis Sci. 51, 5241(2010).</li>
<li>Marmor, M. F., Abdul-Rahim, A. S., Cohen, D. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+effect+of+metabolic+inhibitors+on+retinal+adhesion+and+subretinal+fluid+resorption%5BTitle%5D)+AND+%22Invest+Ophthalmol+Vis+Sci%22%5BJournal%5D)">The effect of metabolic inhibitors on retinal adhesion and subretinal fluid resorption.</a> Invest Ophthalmol Vis Sci. 19, 893-903 (1980).</li>
<li>Frambach, D. A., Marmor, M. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+rate+and+route+of+fluid+resorption+from+the+subretinal+space+of+the+rabbit%5BTitle%5D)+AND+%22Invest+Ophthalmol+Vis+Sci%22%5BJournal%5D)">The rate and route of fluid resorption from the subretinal space of the rabbit.</a> Invest Ophthalmol Vis Sci. 22, 292-302 (1982).</li>
<li>Kita, M., Negi, A., Marmor, M. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Lowering+the+calcium+concentration+in+the+subretinal+space+in+vivo+loosens+retinal+adhesion%5BTitle%5D)+AND+%22Invest+Ophthalmol+Vis+Sci%22%5BJournal%5D)">Lowering the calcium concentration in the subretinal space in vivo loosens retinal adhesion.</a> Invest Ophthalmol Vis Sci. 33, 23-29 (1992).</li>
<li>Marmor, M. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Control+of+subretinal+fluid%3A+experimental+and+clinical+studies%5BTitle%5D)+AND+%22Eye%22%5BJournal%5D)">Control of subretinal fluid: experimental and clinical studies.</a> Eye. 4 (Pt 2), 340-344 (1990).</li>
<li>Faude, F., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Facilitation+of+artificial+retinal+detachment+for+macular+translocation+surgery+tested+in+rabbit%5BTitle%5D)+AND+%22Invest+Ophthalmol+Vis+Sci%22%5BJournal%5D)">Facilitation of artificial retinal detachment for macular translocation surgery tested in rabbit.</a> Invest Ophthalmol Vis Sci. 42, 1328-1337 (2001).</li>
<li>Szurman, P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Ultrastructural+Changes+after+Artificial+Retinal+Detachment+with+Modified+Retinal+Adhesion%5BTitle%5D)+AND+%22Invest.+Ophthalmol.+Vis.+Sci%22%5BJournal%5D)">Ultrastructural Changes after Artificial Retinal Detachment with Modified Retinal Adhesion.</a> Invest. Ophthalmol. Vis. Sci. 47, 4983-4989 (2006).</li>
<li>Fang, X. Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Effect+of+Ca%282%2B%29-free+and+Mg%282%2B%29-free+BSS+Plus+solution+on+the+retinal+pigment+epithelium+and+retina+in+rabbits%5BTitle%5D)+AND+%22Am.J.Ophthalmol%22%5BJournal%5D)">Effect of Ca(2+)-free and Mg(2+)-free BSS Plus solution on the retinal pigment epithelium and retina in rabbits.</a> Am.J.Ophthalmol. 131, 481-488 (2001).</li>
<li>Ivert, L., Kjeldbye, H., Gouras, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Long-term+effects+of+short-term+retinal+bleb+detachments+in+rabbits%5BTitle%5D)+AND+%22Graefes+Arch.Clin.Exp.Ophthalmol%22%5BJournal%5D)">Long-term effects of short-term retinal bleb detachments in rabbits.</a> Graefes Arch.Clin.Exp.Ophthalmol. 240, 232-237 (2002).</li>
</ol>

RB, BVS, ZL, NE and Geuder AG have filed a European patent application on the shooter. NB is an employee of Geuder AG. Publication fees for this video-article were&#160;paid by Geuder AG.

Using a rabbit model, a safe and reproducible method is presented for transvitreal delivery of cultured RPE on cell carriers into the subretinal space with a custom-designed shooter instrument. The described method offers a short/optimized surgical technique for easy learning, as it involves standard techniques in vitrectomy with subretinal maneuvers. Outcome is greatly facilitated by a clean vitreoretinal interface, and intraocular infusion that avoids fluid turbulence over the implantation site, inducing bleb retinal detachment (bRD) at low IOP, preventing retina and sclera damage through dryness, and appropriate positioning of the rabbit.
We caution however, as several intra-operative complications may occur any time, hindering the success of the implantation, for instance intra ocular bleedings, anesthesia fading off during vital steps such as the implantation, collapsing of bRD due to instrument manipulation or ocular hypotony, rabbit death due to excessive doses of anesthesia, low blood pressure during long operation causing hypoxic brain damage, or hyperthermia. Yet these complications decrease with time as they are quickly tackled and resolved by increasing experience of the surgical team.
Some complications could be reduced by following a few simple, yet crucial steps. Lubricant should be added every 5 - 10 min to prevent corneal, scleral and conjunctival damage during the operation, and to maintain a clear intraocular media, as dried/blackened sclera may be a cause of wound dehiscence, which in turn leads to ocular hypotony and/ or intraoperative leakage from sclerotomies. Heparin should be added to prevent the formation of a fibrin film that makes particularly subretinal implantation challenging and simultaneously adding epinephrine to reduce bleeding under heparin16. Too long heparin/ epinephrine exposure times (&#62;1 hour) should be avoided to prevent corneal edema by endothelial decompensation33, hypertensive crisis or intraoperative fatality. Meticulous vitreous removal should be performed at instrument (entry) port to avoid retinal and/ or choroidal detachments. Intraocular instruments should be pointed towards posterior pole to avoid lens touch (causes iatrogenic cataract formation) or (entry site) retinal damage. An intraocular side-port infusion cannula should be used, as it attenuates the jet stream around the implantation area, thus preventing uncontrolled tearing of the retinotomy, and collapse of the bRD. bRD induction in the midline (vertical axis from optic nerve) or close to optic medullar fibers should be avoided to prevent extensive iatrogenic retinal detachments. Finally, last but not least bRD should be induced at low IOP, to avoid subretinal BSS injection using excessive flow rates which may lead to a retinal damage (e.g., by stretching).
Many study variables such as cell carrier variants, fetal, adult or stem cell derived RPE cell sources, choices for immunosuppressants, etc., can be explored 14,26,27,34. Further improvement such as serum-free RPE culture methods, characterization of xenoRPE in subretinal space, removal of the host RPE layer14 or strategies for implant anchorage are current work in progress.
To date the described techniques have been used on 5 different rabbit strains, including chinchilla bastard, Chinchilla bastard/KBL hybrids, New Zealand White/Red Cross, New Zealand White (albino) and Dutch belted. Both male and female rabbits were operated on, with rabbits at least 1.5 kg or 2 months of age (depending on species). Most surgeries were on pigmented rabbits (chinchilla bastard or chinchilla bastard hybrids) with weights between 2.5 - 3 kg.
All of the rabbit strains we have had the opportunity to work with seem to have some peculiarities. Given the exclusive availability of pigmented rabbits of the chinchilla bastard strain in Germany in 2009-13, we have collected the most experience with these animals. Unfortunately it is no longer available, since breeding has been discontinued, but compares very well to New Zealand White/Red Cross except for the more advantageous thicker sclera and larger eye volumes in the latter. Chinchilla bastard hybrids have significant intraoperative fibrin formation and require heparin/epinephrine use as outlined above to ensure successful subretinal maneuvers. This protocol has also been performed in non-pigmented albino rabbits (New Zealand White), however particularly bRD creation and subretinal implantation is more challenging given reduced contrast appreciation. The feasibility of inducing a posterior vitreous detachment did not seem rabbit strain dependent in our hands.
Transvitreal subretinal delivery is likely the future surgical strategy of choice given it is the most common route nowadays clinically to access the retina. As a result many other groups have presented such techniques for cultured RPE on carrier supports in animal work11,15,23,35. Aramant et al.36 have an instrument, which places rather than pushes their hydrogel-encapsulated soft implant to its subretinal target site. The design of Thumann et al. utilizes a hollow spatula, which releases the carrier-supported graft by floating it off through fluid injection 19. Both former strategies require subretinal insertion of the instrument, which in our view is more prone to complications, when compared to an epiretinally appositioned instrument. Montezuma et al.22 described a subretinal inserter instrument for the delivery of subretinal chip implants in pigs but no further work has been published since to the best of our knowledge. We have been able to extend the described technique with some modification to pig.
Our preferred cell carriers are 10 micron thick polyester terephthalate (PET) membranes. From a surgical perspective, this material has favorable stiffness and elasticity parameters, in addition to its broad versatility during cell culture experiments. We found similar experiences with expanded tetrafluoroethylene (ePTFE) 37 or nanofiber membranes electrospun from PET, poly-lactic/capronolactic acid (PLCL) or poly--lactic-co-glycolic acid (PLGA), as well as composite nanofiber (PLGA or PET) and ultrathin PET 26. When PET membranes are used with our metallic shooter instrument, they do have an occasional tendency to exhibit electrostatic charge, which challenges their ejection from the shooter 27. Ultrathin polyimide membranes could in our hands not be implanted in the subretinal space with the protocol outlined above (manuscript in preparation).
Marmor et al. have systematically studied spontaneous resorption of subretinal fluid in iatrogenic localized retinal detachments38-41. Even following manipulation in the subretinal space these were found to be reabsorbed by postoperative day 4 in uneventful surgeries. Laser retinopexy is not performed to secure the edges of the retinotomy. Although counterintuitive when compared to human surgery, air/ gas tamponade is not required. Unless meticulous removal of peripheral vitreous can be achieved, particularly in the superior quadrant, this may in fact result in giant retinal tears originating from the retinotomy site. It is only recommended to perform fluid air exchange with subsequent 20% SF6 gas tamponade to salvage intraoperative iatrogenic retinal detachments or in case a particular implant position needs to be secured.
Although the mechanically induced ablation of the neural retina can cause RPE and photoreceptor damage in rabbits42,43, its extent varies greatly (even with regular BSS) depending on factors such as IOP, syringe type used, injection volume with thereby induced retinal stretching, etc. We have also tested the often recommended Ca/Mg-free BSS facilitated detachment 42-44, but found that it causes intraoperative lens opacification (particularly with elevated temperature), and significantly delays or even impairs retinal re-attachment 27. Slow subretinal injection of 20 - 30 &#181;l volume of regular BSS with a 100 &#181;l syringe is therefore recommended; injection needle movements should be minimal so the retinotomy seals around it and prevent Bruch&#39;s membrane damage. Some of the iatrogenic damage may be resolved by RPE wound healing, and the observed relative preservation of ONL thickness after reattachment, suggests that the RPE/photoreceptor complex can tolerate this impairment, as also described by others45.
Cell-based therapeutics or retinal prosthetics require preclinical animal testing prior to regulatory approval and commencing human safety studies. The former vary from country to country. The rabbit model described here may serve as a cost-efficient and less challenging platform for establishing or even carrying out all requirements by regulatory authorities. Moreover, it may subsequently serve for training of surgeons in eventual multicenter clinical trials or further improvements of the technique along the way.

Age-related macular degeneration (AMD) is the most common cause of visual impairment in adults aged 50 or older in industrially developed countries, as it causes loss of central vision. About 15% of these patients suffer from the &#34;wet&#34; form of the disease, in which neovascularization originates from the choroid and disrupts retinal function1. This variant can be treated by a highly effective therapy with repeated intra-vitreal injections of antiangiogenic drugs2. However, the vast majority of patients (~85%) suffer from the dry form, which is characterized by extracellular deposits (e.g., drusen) under the retinal pigment epithelium (RPE). These deposits cause RPE dysfunction leading to retinal atrophy in the macula. Given the lack of any curative therapeutic options, AMD evolved into an intensively developing research field, where many different curative therapeutic approaches are being tested. Surgical RPE replacement is one attractive future possibility to defeat this debilitating disease3.
Autologous subretinal RPE transplantation replaces dysfunctional or lost RPE in macula, and has the potential to restore its physiological function4-9. This surgical technique had a breakthrough with the development of RPE differentiation protocols from human embryonic stem cells (hESC) and induced pluripotent stem cells (iPSC), giving the scientist an unlimited cell source of RPE for transplantation10. RPE transplantation is now recognized as an attractive first-in-human application for stem cell derived therapeutics. The eye offers excellent surgical access and sophisticated in vivo monitoring tools11-13.
To transplant the RPE, one way is with a minimally invasive delivery using a cell suspension, alternatively, to better preserve RPE characteristics and transplant function, arti&#64257;cial carrier substrates (scaffolds) for RPE replacement are being considered4,14,15. Large animal models are required for preclinical validation, yet detailed technical information on animal handling and surgical technique is missing to date16-23.
We and others11,24 despite some evidence to the contrary25, suggest the use of a rigid yet elastic carrier substrate as it provides safer handling, preserves monolayer integrity and functionality. Over time we have tested several custom-designed instruments and ancillary techniques for the implantation of cell-carrier supported RPE transplants into the subretinal space (SRS). We utilized intraoperative video recordings, in vivo scanning laser ophthalmoscopy combined with spectral domain optical coherence tomography (SLO/ SD-OCT), and histology to evaluate the implantation success14,26,27. Here we provide our current recommendation for subretinal RPE implants in rabbits, which were tested in 5 different rabbit strains, 7 cell carrier materials and 4 RPE cell sources in over 150 procedures.

<table><tbody><tr><td>s30 ultrasonic cleaning unit</td><td>Elmasonic</td><td>100 4631</td><td>2.75 L</td></tr><tr><td>DE-23&amp;nbsp; autoclave</td><td>Systec</td><td>C 2209</td><td>23 L</td></tr><tr><td>Syringe&amp;nbsp;</td><td>BD&amp;nbsp;</td><td>300013&lt;br/&gt; 300995&amp;nbsp;&lt;br/&gt; 301285&lt;br/&gt; 300294&lt;br/&gt; 300330&amp;nbsp;</td><td>1 ml x3&lt;br/&gt; 2 ml x3&lt;br/&gt; 5 ml x1&lt;br/&gt; 10 ml x1&lt;br/&gt; 20 ml x1</td></tr><tr><td>Needle&amp;nbsp;</td><td>BD</td><td>305196&lt;br/&gt; 305136</td><td>18 G x1&amp;nbsp;&lt;br/&gt; 27 G x5</td></tr><tr><td>Scalpel</td><td>Feather</td><td>2975#20</td><td>blade#20 x3</td></tr><tr><td>Surgical drape</td><td>HARTMANN&lt;br/&gt; LOHMANN &amp;amp; RAUSCHER</td><td>277 502&lt;br/&gt; 25 440</td><td>60 x 40 cm x2&lt;br/&gt; 12 x 17 cm</td></tr><tr><td>Ocular sticks</td><td>LOHMANN &amp;amp; RAUSCHER</td><td>16 516</td><td>66 x 5 mm</td></tr><tr><td>Twister gauze sponges</td><td>HARTMANN</td><td>481 274</td><td>x2</td></tr><tr><td>Closure strips</td><td>HARTMANN</td><td>540 686</td><td>x4</td></tr><tr><td>Opmi Visu CS Microscope</td><td>Zeiss</td><td>N/a</td><td>incl. fundus imaging system BIOM II</td></tr><tr><td>Chandelier endoillumination</td><td>Geuder</td><td>G-S03503 +&lt;br/&gt; G-S03504</td><td>25 G incl. trocar</td></tr><tr><td>Light machine</td><td>Geuder</td><td>G-26033</td><td>Xenotron III</td></tr><tr><td>Vitrectomy machine</td><td>Geuder</td><td>G-60000</td><td>MegaTRON S4 S4/ HPS</td></tr><tr><td>Vitrector</td><td>Geuder</td><td>G-46301</td><td>MACH2 vitreous cutter 23 G</td></tr><tr><td>Venturi cassette&amp;nbsp;</td><td>Geuder</td><td>G-60700</td><td></td></tr><tr><td>Sideport-infusion cannula</td><td>Geuder</td><td>custom&amp;nbsp;</td><td>23 G x1</td></tr><tr><td>3-0 silk suture</td><td>ETHICON</td><td>V546G</td><td>x1</td></tr><tr><td>Caliper</td><td>Geuder</td><td>G-19135</td><td>x2</td></tr><tr><td>Vannas scissors</td><td>Geuder</td><td>G-19777</td><td>x1</td></tr><tr><td>Sclerotomie blade</td><td>Ziemer</td><td>21-2301</td><td>1x 23 G&lt;br/&gt; 1x 20 G</td></tr><tr><td>7-0 silk suture</td><td>ETHICON</td><td>EH6162H</td><td>x1</td></tr><tr><td>Needle holder</td><td>Geuder</td><td>G-32320</td><td>x2</td></tr><tr><td>Iris forceps</td><td>Geuder</td><td>G-18910</td><td>x1</td></tr><tr><td>Colibri forceps</td><td>Geuder</td><td>G-18950</td><td>x1</td></tr><tr><td>Extendible subretinal injection needle</td><td>DORC</td><td>1270.EXT</td><td>41 G</td></tr><tr><td>VR scissor</td><td>Geuder</td><td>G-36542</td><td>25 G</td></tr><tr><td>Grieshaber forceps holder</td><td>Alcon</td><td>712.00.41</td><td>23 G</td></tr><tr><td>Curved scissor forceps tips</td><td>Alcon</td><td>723.52</td><td>23 G</td></tr><tr><td>Implant loading station</td><td>Dow Corning</td><td>3097358-1004</td><td>SYLGARD 184 Silicone Elastomer Kit</td></tr><tr><td>Blunt oval implant trephine&amp;nbsp;</td><td>Geuder</td><td>custom-made&amp;nbsp;</td><td>2.4 x 1.1 mm&amp;nbsp;</td></tr><tr><td>Shooter dummy</td><td>Geuder</td><td>G-32227</td><td>x1</td></tr><tr><td>Shooter</td><td>Geuder</td><td>G-S03443</td><td>x1</td></tr><tr><td>Flute needle</td><td>DORC</td><td>1281.SD</td><td>20 G (Vacuum)</td></tr><tr><td>Manual microliter syringe</td><td>Hamilton</td><td>24535</td><td>100 &amp;micro;l</td></tr><tr><td>Tissue culture plates</td><td>Greiner bio-one</td><td>&amp;nbsp;664160</td><td>100 x 20 mm</td></tr><tr><td>Spectralis Multi-Modality&lt;br/&gt; Imaging System</td><td>Heidelberg&lt;br/&gt; Engineering</td><td>N/a</td><td>Spectralis HRA&lt;br/&gt; &amp;nbsp;+ OCT</td></tr><tr><td>&lt;strong&gt;Drugs and solutions&lt;/strong&gt;</td><td></td><td></td><td></td></tr><tr><td>&lt;strong&gt;Name&lt;/strong&gt;</td><td>&lt;strong&gt;Company&lt;/strong&gt;</td><td>&lt;strong&gt;Active agent&lt;/strong&gt;</td><td>&lt;strong&gt;Comments&lt;/strong&gt;</td></tr><tr><td>Mucadont-IS</td><td>Merz Hygiene</td><td>virucidal instrument disinfectant&amp;nbsp;</td><td>2 L</td></tr><tr><td>Mucocit&amp;nbsp;T</td><td>Merz Hygiene</td><td>Aldehyde-free instrument disinfectant</td><td>2 L</td></tr><tr><td>Ketamin 10%</td><td>WDT</td><td>Ketamine&amp;nbsp;</td><td>10 ml (100 mg/ml)</td></tr><tr><td>Domitor</td><td>Orion Pharma</td><td>Medetomidine hydrochloride</td><td>10 ml (1 mg/ml)</td></tr><tr><td>Antisedan</td><td>Orion Pharma</td><td>Atipamezole hydrochloride</td><td>10 ml (5 mg/ml)</td></tr><tr><td>Neosynephrin POS 10%</td><td>URSAPHARM</td><td>Phenylephrine HCl</td><td>10 ml</td></tr><tr><td>Mydriacyl</td><td>Alcon</td><td>Tropicamid</td><td>10 ml (5 mg/ml)</td></tr><tr><td>Methocel 2%</td><td>Omni Vision</td><td>hydroxypropyl methylcellulose</td><td>10 g</td></tr><tr><td>PURI CLEAR</td><td>ZEISS</td><td>Balance salt solution (BSS)&amp;nbsp;</td><td>500 ml</td></tr><tr><td>Glucose 5%&amp;nbsp;&amp;nbsp;</td><td>B.Braun</td><td>Glucose 5%&amp;nbsp; solution</td><td>100 ml</td></tr><tr><td>Heparin-Natrium-25,000</td><td>Ratiopharm</td><td>Heparin</td><td>5 ml (2,500 unit/ml)</td></tr><tr><td>Suprarenin</td><td>SANOFI</td><td>Epinephrine</td><td>1 ml (1 mg/ml)</td></tr><tr><td>Triamcinolone</td><td>University of Bonn pharmacy</td><td>preservative-free Triamcinolone&amp;nbsp;</td><td>1 ml (40 mg/ml)&amp;nbsp;</td></tr><tr><td>Isoptomax eye ointment</td><td>Alcon</td><td>dexamethasone 1 mg/g&lt;br/&gt; neomycin sulfate 3,500 IU/g&lt;br/&gt; polymyxin B sulfate 6,000 IU/g</td><td>10 ml</td></tr><tr><td>Betaisodona</td><td>Mundipharma</td><td>Povidon-Iod</td><td>30 ml (1 g/10 ml)</td></tr><tr><td>Optive</td><td>ALLERGAN</td><td>sodium carboxymethylcellulose glycerol</td><td>10 ml</td></tr></tbody></table>

a step by step protocol for subretinal surgery in rabbits

Age related macular degeneration (AMD), retinitis pigmentosa, and other RPE related diseases are the most common causes for irreversible loss of vision in adults in industrially developed countries. RPE transplantation appears to be a promising therapy, as it may replace dysfunctional RPE, restore its function, and thereby vision.
Here we describe a method for transplanting a cultured RPE monolayer on a scaffold into the subretinal space (SRS) of rabbits. After vitrectomy xenotransplants were delivered into the SRS using a custom made shooter consisting of a 20-gauge metallic nozzle with a polytetrafluoroethylene (PTFE) coated plunger. The current technique evolved in over 150 rabbit surgeries over 6 years. Post-operative follow-up can be obtained using non-invasive and repetitive in vivo imaging such as spectral domain optical coherence tomography (SD-OCT) followed by perfusion-fixed histology.
The method has well-defined steps for easy learning and high success rate. Rabbits are considered a large eye animal model useful in preclinical studies for clinical translation. In this context rabbits are a cost-efficient and perhaps convenient alternative to other large eye animal models.

The results from the described method for subretinal implantation are shown in Table 2. Engraftment under the retina had a success rate of ca. 61% when a core vitrectomy was performed and rose up to 76%, when posterior vitreous detachment was induced. These numbers include ca. 21% of animals who died either intraoperatively or in the first 3 postoperative days. This technique can be used to implant two scaffolds on different retinal areas in one eye simultaneously.
The rabbits underwent postoperative in vivo follow up using SD-OCT and histologic processing as described by Stanzel et al.27 (Figure 2). Fig. 2A&#160;shows scanning laser ophthalmoscope infrared reflectance image of implanted cultured RPE on a polyester membrane (PET) after an uncomplicated transplantation. The halo around implant corresponds to photoreceptor atrophy. Fig. 2B&#160;shows corresponding SD-OCT, notice retinal, mainly outer nuclear layer (ONL) thinning, hyper reflective band on SD-OCT above implant, while the neural retina adjacent to the implant shows near-normal reflection bands. These results suggest an atraumatic delivery. Fig. 2C&#160;shows Hematoxiline/Eosin (H/E) stain of the implant which shows subretinal scarring and ONL atrophy around the retinotomy site likely as a result of iatrogenic manipulation, a contiguous yet irregular pigmented layer over PET. Bruch&#39;s membrane underneath the implant also appeared to be contiguous, and the choriocapillaris contains some scattered erythrocytes. These morphologic result are comparable to the SD-OCT and strengthen the thesis of an atraumatic delivery.
<img alt="Figure 1" src="/files/ftp_upload/53927/53927fig1.jpg" /> 
Figure 1: Loading Shooter with Human iPSC-RPE Cultured on PET Cell Carrier. A) Shows punching out an implant using custom made needle. B) Bean-shaped implant cut from cell culture. C) Positioning of implant before loading. D) Loading of shooter with implant. <a href="https://www.jove.com/files/ftp_upload/53927/53927fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/53927/53927fig2.jpg" /> 
Figure 2: Cultured Human hES-RPE on PET Cell Carrier 4 Weeks in Rabbit Subretinal Space. A) Shows SLO infrared reflectance image, the green line demarcates the cross-section shown in Fig. 2B. B) Corresponding SD-OCT. C) H/E stain, see text or26,27 for details. <a href="https://www.jove.com/files/ftp_upload/53927/53927fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table fo:keep-together.within-page="1">
	<colgroup>
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:97px;">Parameter</td>
			<td style="width:131px;">Used settings</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Vitrectomy</td>
			<td>6,000 cuts/min</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Vacuum</td>
			<td>200 mmHg</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Rise time</td>
			<td>1 sec</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">&#160;Air</td>
			<td>24 mmHg</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Irrigation&#160;</td>
			<td>24 cmH2O</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Diathermy</td>
			<td>30%</td>
		</tr>
	</tbody>
</table>
Table 1: Parameter Setup of the Vitrectomy Machine.
<table fo:keep-together.within-page="1">
	<tbody>
		<tr>
			<td>Vitrectomy</td>
			<td>Implant</td>
			<td>Rabbits operated</td>
			<td>Successful implant</td>
			<td>Failed implant</td>
			<td>Death</td>
			<td>Success rate%</td>
		</tr>
		<tr>
			<td>Core vitrectomy</td>
			<td>PET</td>
			<td>30</td>
			<td>19</td>
			<td>4</td>
			<td>7</td>
			<td>63.33</td>
		</tr>
		<tr>
			<td></td>
			<td>PET + RPE</td>
			<td>70</td>
			<td>42</td>
			<td>12</td>
			<td>16</td>
			<td>60</td>
		</tr>
		<tr>
			<td>PVD, &#177;&#160;plasmin w/PPV</td>
			<td>PET</td>
			<td>28</td>
			<td>21</td>
			<td>2</td>
			<td>5</td>
			<td>75</td>
		</tr>
		<tr>
			<td></td>
			<td>PET + RPE</td>
			<td>22</td>
			<td>17</td>
			<td>2</td>
			<td>3</td>
			<td>77.27</td>
		</tr>
	</tbody>
</table>
Table 2: Summary of the Last 150 Operations Including Method and Implant Type.

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A Step by Step Protocol for Subretinal Surgery in Rabbits

Retinal pigment epithelium (RPE) replacement strategies and gene-based therapy are considered for several retinal degenerative conditions. For clinical translation, large eye animal models are required to study surgical techniques applicable in patients. Here we present a rabbit model for subretinal surgery geared towards RPE transplantation, which is versatile and cost-efficient.

Age related macular degeneration (AMD), retinitis pigmentosa, and other RPE related diseases are the most common causes for irreversible loss of vision in ...

a-step-by-step-protocol-for-subretinal-surgery-in-rabbits

Research

JoVE Journal

Medicine

15.4K Views.  University of Bonn.  Netvliespigmentepitheel (RPE) vervangende strategieën en gen-gebaseerde therapie in aanmerking komen voor een aantal retinale degeneratieve aandoeningen. Voor klinische toepassing worden grote oog dierlijke modellen moeten chirurgische technieken bij patiënten toepassing bestuderen. Hier presenteren we een konijnenmodel voor subretinale chirurgie gericht RPE transplantatie, die veelzijdig en kostenefficiënt. 

Video: A Step by Step Protocol for Subretinal Surgery in Rabbits

Novel Photoacoustic Microscopy and Optical Coherence Tomography Dual-modality Chorioretinal Imaging in Living Rabbit Eyes

Photoacoustic ocular imaging is an emerging ophthalmic imaging technology that can noninvasively visualize ocular tissue by converting light energy into sound waves and is currently under intensive investigation. However, most reported work to date is focused on the imaging of the posterior segment of the eyes of small animals, such as rats and mice, which poses challenges for clinical human translation due to small eyeball sizes. This manuscript describes a novel photoacoustic microscopy (PAM) and optical coherence tomography (OCT) dual-modality system for posterior segment imaging of the eyes of larger animals, such as rabbits. The system configuration, system alignment, animal preparation, and dual-modality experimental protocols for in vivo, noninvasive, label-free chorioretinal imaging in rabbits are detailed. The effectiveness of the method is demonstrated through representative experimental results, including retinal and choroidal vasculature obtained by the PAM and OCT. This manuscript provides a practical guide to reproducing the imaging results in rabbits and advancing photoacoustic ocular imaging in larger animals.

This manuscript describes the novel setup and operating procedure of a photoacoustic microscopy and optical coherence tomography dual-modality system for noninvasive, label-free chorioretinal imaging of larger animals, such as rabbits.

Photoacoustic ocular imaging is an emerging ophthalmic imaging technology that can noninvasively visualize ocular tissue by converting light energy into ...

Engineering

The Rabbit Blood-shunt Model for the Study of Acute and Late Sequelae of Subarachnoid Hemorrhage: Technical Aspects

Early brain injury and delayed cerebral vasospasm both contribute to unfavorable outcomes after subarachnoid hemorrhage (SAH). Reproducible and controllable animal models that simulate both conditions are presently uncommon. Therefore, new models are needed in order to mimic human pathophysiological conditions resulting from SAH.
This report describes the technical nuances of a rabbit blood-shunt SAH model that enables control of intracerebral pressure (ICP). An extracorporeal shunt is placed between the arterial system and the subarachnoid space, which enables examiner-independent SAH in a closed cranium. Step-by-step procedural instructions and necessary equipment are described, as well as technical considerations to produce the model with minimal mortality and morbidity. Important details required for successful surgical creation of this robust, simple and consistent ICP-controlled SAH rabbit model are described.

The experimental intracranial pressure-controlled blood shunt subarachnoid hemorrhage (SAH) model in the rabbit combines the standard procedures &#8212; subclavian artery cannulation and transcutaneous cisterna magna puncture, which enables close mimicking of human pathophysiological conditions after SAH. We present step-by-step instructions and discuss key surgical points for successful experimental SAH creation.

Early brain injury and delayed cerebral vasospasm both contribute to unfavorable outcomes after subarachnoid hemorrhage (SAH). Reproducible and ...

Scleral Cross-linking Using Riboflavin and Ultraviolet-A Radiation for Prevention of Axial Myopia in a Rabbit Model

Myopic individuals, especially those with severe myopia, are at higher-than-normal risk of cataract, glaucoma, retinal detachment and chorioretinal abnormalities. In addition, pathological myopia is a common irreversible cause of visual impairment and blindness1-3. Our study demonstrates the effect of scleral crosslinking using riboflavin and ultraviolet-A radiation on the development of axial myopia in a rabbit model. The axial length of the eyeball was measured by A-scan ultrasound in New Zealand white rabbits aged 13 days (male and female). The eye then underwent 360&#176; conjunctival peritomy with scleral crosslinking, followed by tarsorrhaphy. Axial elongation was induced in 13 day-old New Zealand rabbits by suturing their right eye eyelids (tarsorrhaphy). The eyes were divided into quadrants, and every quadrant had two scleral irradiation zones, each with an area of 0.2 cm&#178; and a radius of 4 mm. Crosslinking was performed by dropping 0.1% dextran-free riboflavin-5-phosphate onto the irradiation zones 20 sec before ultraviolet-A irradiation and every 20 sec during the 200 sec irradiation time. UVA radiation (370 nm) was applied perpendicular to the sclera at 57 mW/cm&#178; (total UVA light dose, 57 J/cm&#178;). Tarsorrhaphies were removed on day 55, followed by repeated axial length measurements. This study demonstrates that scleral crosslinking with riboflavin and ultraviolet-A radiation effectively prevents occlusion-induced axial elongation in a rabbit model.

We demonstrate the effect of scleral crosslinking with riboflavin and UVA on an axial elongation rabbit eye. Axial elongation was induced in 13 day-old New Zealand rabbits (male and female) by suturing their right eye eyelids (tarsorrhaphy).

Myopic individuals, especially those with severe myopia, are at higher-than-normal risk of cataract, glaucoma, retinal detachment and chorioretinal ...

Use of Rabbit Eyes in Pharmacokinetic Studies of Intraocular Drugs

The intraocular route of drug administration enables the delivery of high concentrations of therapeutic drugs, while minimizing their systemic absorption. Several drugs are administered into the anterior chamber or vitreous, and the intraocular injection has been effective in curing various intraocular diseases. Rabbit eyes have been widely used for ophthalmic research, as the animal is easy to handle and economical compared to other mammals, and the size of a rabbit eye is similar to that of a human eye. Using a 30 G needle, drugs can be injected into the intracameral and intravitreal spaces of rabbit eyes. The eyeballs are then frozen until analysis, and can be divided into the aqueous humor, vitreous, and retina/choroid. The vitreous and retina/choroid samples can be homogenized and solubilized before analysis. Then, immunoassays can be performed to measure the concentrations of intraocular drugs in each compartment. Appropriate pharmacokinetic models can be used to calculate several parameters, such as the half-life and maximum concentration of the drug. Rabbit eyes can be a good model for pharmacokinetic studies of intraocular drugs.

Rabbits are widely used to study the pharmacokinetics of intraocular drugs. We describe a method for conducting pharmacokinetic studies of intraocular drugs using rabbit eyes.

The intraocular route of drug administration enables the delivery of high concentrations of therapeutic drugs, while minimizing their systemic absorption. ...

Development of a Refined Protocol for Trans-scleral Subretinal Transplantation of Human Retinal Pigment Epithelial Cells into Rat Eyes

Degenerative retinal diseases such as age-related macular degeneration (AMD) are the leading cause of irreversible vision loss worldwide. AMD is characterized by the degeneration of retinal pigment epithelial (RPE) cells, which are a monolayer of cells functionally supporting and anatomically wrapping around the neural retina. Current pharmacological treatments for the non-neovascular AMD (dry AMD) only slow down the disease progression but cannot restore vision, necessitating studies aimed at identifying novel therapeutic strategies. Replacing the degenerative RPE cells with healthy cells holds promise to treat dry AMD in the future. Extensive preclinical studies of stem cell replacement therapies for AMD involve the transplantation of stem cell-derived RPE cells into the subretinal space of animal models, in which the subretinal injection technique is applied. The approach most frequently used in these preclinical animal studies is through the trans-scleral route, which is made difficult by the lack of direct visualization of the needle end and can often result in retinal damage. An alternative approach through the vitreous allows for direct observation of the needle end position, but it carries a high risk of surgical traumas as more eye tissues are disturbed. We have developed a less risky and reproducible modified trans-scleral injection method that uses defined needle angles and depths to successfully and consistently deliver RPE cells into the rat subretinal space and avoid excessive retinal damage. Cells delivered in this manner have been previously demonstrated to be efficacious in the Royal College of Surgeons (RCS) rat for at least 2 months. This technique can be used not only for cell transplantation but also for delivery of small molecules or gene therapies.

Subretinal injection has been widely applied in preclinical studies of stem cell replacement therapy for age-related macular degeneration. In this visualized article, we describe a less risky, reproducible and precisely modified subretinal injection technique via the trans-scleral approach to deliver cells into rat eyes.

Degenerative retinal diseases such as age-related macular degeneration (AMD) are the leading cause of irreversible vision loss worldwide. AMD is ...

Neuroscience

Subretinal Implantation of RPE on a Carrier in Minipigs: Guidelines for Preoperative Preparations, Surgical Techniques, and Postoperative Care

Degenerative disorders of the retina (including age-related macular degeneration), which originate primarily at or within the retinal pigmented epithelial (RPE) layer, lead to a progressive disorganization of the retinal anatomy and the deterioration of visual function. The substitution of damaged RPE cells (RPEs) with in vitro cultured RPE cells using a subretinal cell carrier has shown potential for re-establishing the anatomical structure of the outer retinal layers and is, therefore, being further studied. Here, we present the principles of a surgical technique that allows for the effective subretinal transplantation of a cell carrier with cultivated RPEs into minipigs. The surgeries were performed under general anesthesia and included a standard lens-sparing three-port pars plana vitrectomy (PPV), subretinal application of a balanced salt solution (BSS), a 2.7 mm retinotomy, implantation of a nanofibrous cell carrier into the subretinal space through an additional 3.0 mm sclerotomy, fluid-air exchange (FAX), silicone oil tamponade, and closure of all the sclerotomies. This surgical approach was used in 29 surgeries (18 animals) over the past 8 years with a success rate of 93.1%. Anatomic verification of the surgical placement was carried out using in vivo fundus imaging (fundus photography and optical coherence tomography). The recommended surgical steps for the subretinal implantation of RPEs on a carrier in minipig eyes can be used in future preclinical studies using large-eye animal models.

The subretinal implantation of retinal pigmented epithelium (RPE) is one of the most promising approaches for the treatment of degenerative retinal diseases. However, the performance of preclinical studies on large-eye animal models remains challenging. This report presents guidelines for the subretinal transplantation of RPE on a cell carrier into minipigs.

Degenerative disorders of the retina (including age-related macular degeneration), which originate primarily at or within the retinal pigmented epithelial ...

Bioengineering

Arterial Pouch Microsurgical Bifurcation Aneurysm Model in the Rabbit

Endovascular treatment for intracranial aneurysms gained importance over the past decades, consequently there is an increased need of testing endovascular devices. Animal models respecting rheological, hemodynamic and aneurysm wall conditions are highly warranted. Therefore, the aim of the present study was to design a novel standardized and reproducible surgical technique to create autologous arterial pouch bifurcation aneurysms with non-modified and modified wall conditions in rabbits.
Bifurcation aneurysms were created by end-to-side anastomosis of the right on the left common carotid artery, both serving as parent arteries for the arterial pouch, which was microsurgically sewn on. Grafts were taken from the proximal right common carotid artery, either for the control (n = 7, immediate autologous re-implantation) or modified (n = 7, incubated with 100 international units elastase for 20 minutes before autologous re-implantation) group. Pouch and parent artery patency were controlled by fluorescence angiography immediately after creation. At follow-up (28 days), all rabbits underwent contrast enhanced magnetic resonance angiography and fluorescence angiography followed by aneurysm harvesting, macroscopic and histological evaluation.
A total of 16 female New Zealand White rabbits were operated upon. Two animals died prematurely. At follow-up, 85.72% of all aneurysms remained patent. Both groups revealed an increase in aneurysm size over time; this was more pronounced in the control group (6.48 &#177; 1.81 mm3 at time of creation vs. 19.85 &#177; 6.40 mm3 at follow-up, p = 0.037) than in the modified group (8.03 &#177; 1.08 mm3 at time of creation vs. 20.29 &#177; 6.16 mm3 at follow-up, p = 0.054).
Our findings demonstrate the adequacy of this new rabbit model which allows for the creation of bifurcation aneurysms with different wall conditions in a microsurgical approach. Given the excellent long-term patency and the property of aneurysm growth over time, this model may serve as an important tool for preclinical evaluation of novel endovascular therapies.

Developing and testing endovascular devices for intracranial aneurysm treatment is still of great importance. Most aneurysm models used today miss either the important characteristics of an arterial degenerated wall or the hemodynamics of a true bifurcation. Therefore, we aimed to design a novel arterial pouch bifurcation model in rabbits.

Endovascular treatment for intracranial aneurysms gained importance over the past decades, consequently there is an increased need of testing endovascular ...

Establishment of a Severe Dry Eye Model Using Complete Dacryoadenectomy in Rabbits

Dry eye disease (DED) is a complex disease with multiple etiologies and variable symptoms, having ocular surface inflammation as its key pathophysiologic step. Despite advances in our understanding of DED, significant knowledge gaps remain. Advances are limited in part due to the lack of informative animal models. The authors recently reported on a method of DED induced by injecting all orbital lacrimal gland (LG) tissues with the lectin concanavalin A. Here, we report a novel model of aqueous-deficient DED based on the surgical resection of all orbital LG (dacryoadenectomy) tissues. Both methods use rabbits because of their similarity to human eyes in terms of the size and structure of the ocular surface. One week after removal of the nictitating membrane, the orbital superior LG was surgically removed under anesthesia, followed by removal of the palpebral superior LG, and finally removal of the inferior LG. Dacryoadenectomy induced severe DED, evidenced by a marked reduction in the tear break up time test and the Schirmer&#39;s tear test, and significantly increased tear osmolarity and rose bengal staining. Dacryoadenectomy-induced DED lasted at least eight weeks. There were no complications and animals tolerated the procedure well. The technique can be mastered relatively easily by those with adequate surgical experience and appreciation of the relevant rabbit anatomy. Since this model recapitulates the features of human aqueous-deficient DED, it is suitable for studies of ocular surface homeostasis, DED, and candidate therapeutics.

A novel approach is presented to induce chronic dry eye disease in rabbits by surgically removing all orbital lacrimal glands. This method, distinct from those previously reported, produces a stable, reproducible model of aqueous deficient dry eye well suited to study tear physiology and pathophysiology and ocular therapeutics.

Dry eye disease (DED) is a complex disease with multiple etiologies and variable symptoms, having ocular surface inflammation as its key pathophysiologic ...

Development of a Surgical Technique for Subretinal Implants in Rats

Retinal degeneration, such as age-related macular degeneration (AMD), is a leading cause of blindness worldwide. A myriad of approaches have been undertaken to develop regenerative medicine-based therapies for AMD, including stem cell-based therapies. Rodents as animal models for retinal degeneration are a foundation for translational research, due to the broad spectrum of strains that develop retinal degeneration diseases at different stages. However, mimicking human therapeutic delivery of subretinal implants in rodents is challenging, due to anatomical differences such as lens size and vitreous volume. This surgical protocol aims to provide a guided method for transplanting implants into the subretinal space in rats. A user-friendly comprehensive description of the critical steps has been included. This protocol has been developed as a cost-efficient surgical procedure for reproducibility across different preclinical studies in rats. Proper miniaturization of a human-sized implant is required prior to conducting the surgical experiment, which includes adjustments to the dimensions of the implant. An external approach is used instead of an intravitreal procedure to deliver the implant to the subretinal space. Using a small sharp needle, a scleral incision is performed in the temporal superior quadrant, followed by paracentesis to reduce intraocular pressure, thereby minimizing resistance during the surgical implantation. Next, a balanced salt solution (BSS) injection through the incision is carried out to achieve focal retinal detachment (RD). Lastly, insertion and visualization of the implant into the subretinal space are conducted. Post-operative assessment of the subretinal placement of the implant includes imaging by spectral domain optical coherence tomography (SD-OCT). Imaging follow-ups ascertain the subretinal stability of the implant, before the eyes are harvested and fixated for histological analysis.

The present protocol describes the scleral approach for subretinal device implantation, a feasible surgical technique for implementation in animal models of retinal diseases in research.

Retinal degeneration, such as age-related macular degeneration (AMD), is a leading cause of blindness worldwide. A myriad of approaches have been ...

Subretinal Transplantation of Human Embryonic Stem Cell Derived-retinal Pigment Epithelial Cells into a Large-eyed Model of Geographic Atrophy

Geographic atrophy (GA), the late stage of dry age-related macular degeneration is characterized by loss of the retinal pigment epithelial (RPE) layer, which leads to subsequent degeneration of vital retinal structures (e.g., photoreceptors) causing severe vision impairment. Similarly, RPE-loss and decrease in visual acuity is seen in long-term follow up of patients with advanced wet age-related macular degeneration (AMD) receiving intravitreal anti-vascular endothelial growth factor (VEGF) treatment. Therefore, on the one hand, it is fundamental to efficiently derive RPE cells from an unlimited source that could serve as replacement therapy. On the other hand, it is important to assess the behavior and integration of the derived cells in a model of the disease entailing surgical and imaging methods as close as possible to those applied in humans. Here, we provide a detailed protocol based on our previous publications that describes the generation of a preclinical model of GA using the albino rabbit eye, for evaluation of the human embryonic stem cell derived retinal pigment epithelial cells (hESC-RPE) in a clinically relevant setting. Differentiated hESC-RPE are transplanted into naive eyes or eyes with NaIO3-induced GA-like retinal degeneration using a 25 G transvitreal pars plana technique. Evaluation of degenerated and transplanted areas is performed by multimodal high-resolution non-invasive real-time imaging.

Retinal pigment epithelial cells could serve as a cell-replacement therapy for the advanced form of dry age-related macular degeneration. This protocol describes the generation of a large-eyed model of geographic atrophy and the subretinal transplantation of human embryonic stem cell-derived retinal pigment epithelial cells into this model of disease.

Geographic atrophy (GA), the late stage of dry age-related macular degeneration is characterized by loss of the retinal pigment epithelial (RPE) layer, ...

A Step by Step Protocol for Subretinal Surgery in Rabbits

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