This work was funded by the following funding: NEI R01EY033103 (MSS), Foundation Fighting Blindness (MSS),the Shulsky Foundation (MSS), the Joseph Albert Hekimian Fund (MSS), the Juliette RP Vision Foundation (YVL), Research to Prevent Blindness (unrestricted grant to the Wilmer Eye Institute at Johns Hopkins University and the Cullen Eye Institute at Baylor College of Medicine). We thank Dr. Malia Edwards (Johns Hopkins University School of Medicine) for kindly providing training on the microscope.

All animal experiments were carried out in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals (NIH Publications No. 8023, revised 1978) and the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research. All procedures were approved by the Johns Hopkins University Animal Care and Use Committee (approval M021M459).
1. Animals
<ol>
	<li>Use Rho::EGFP mice (aged P3-P6) as donors of retinal cell suspensions. 
	NOTE: This strain was a kind gift from Dr. T. Wensel, Baylor College of Medicine.</li>
	<li>Use OPN1LW-EGFP/NRL-/- mice14 (aged P3) as donors of retinal sheets.</li>
	<li>Use adult retinal degeneration Rd1/NS mice with immune deficiency (either sex) as recipients.</li>
	<li>House all mice in cages under a 12:12 h light-dark cycle with access to water and food as needed.</li>
</ol>
2. Collect neural retina (Figure 1)
NOTE: All the following steps were performed under sterile conditions. Supplier information of the research tools and products is provided in the Table of Materials.
<ol>
	<li>Donor mice preparation: Euthanize donor mice with an overdose of carbon dioxide and confirm euthanasia with cervical dislocation.</li>
	<li>Isolate mice eyeballs. To do so follow the steps below.
	<ol>
		<li>Carefully open pups&#39; eyelids using micro scissors to expose the eyeball.</li>
		<li>Use smooth forceps to grab the optic nerve and pull out the eyeball with a pair of forceps.</li>
	</ol>
	</li>
	<li>Isolate neural retina 
	NOTE: This step is performed under a dissection microscope.
	<ol>
		<li>Incise a hole in the center of the cornea using a 25 G sterile needle.</li>
		<li>Cut the cornea, through the hole, in half and enlarge the incision to the sclera and RPE, then remove the sclera and RPE.</li>
		<li>Use micro-toothed forceps to gently remove the lens and the vitreous to isolate the neural retina.</li>
		<li>Proceed with section 3 to prepare donor retinal suspension or section 4 to prepare donor retinal sheet, as required.</li>
	</ol>
	</li>
</ol>
3. Prepare donor retinal suspension (Figure 1)
<ol>
	<li>Incubate the neural retina in Papain solution at 37 &#176;C for 20-30 min until no cell clumps are detected. 
	NOTE: Gently pipette the cell mixture 15x every 10 min.</li>
	<li>Follow manufacturer&#39;s instructions of the Papain Dissociation Kit to collect single cells.</li>
</ol>
4. Prepare donor retinal sheets (Figure 1)
NOTE: This step follows section 2 and is performed under a dissection microscope.
<ol>
	<li>Put the isolated neural retina cup into a 35 mm Petri dish with prechilled sterile PBS.</li>
	<li>Use micro scissors to cut the neural retina cup into multiple retinal sheets (around 1 mm x 2 mm).</li>
</ol>
5. Prepare recipient mice
<ol>
	<li>Anesthetize recipient mice with intraperitoneal injection of ketamine (100 mg/kg body weight) and xylazine hydrochloride (20 mg/kg body weight).</li>
	<li>Confirm the surgical plane of anesthesia by ensuring the loss of blink and pain reflexes, regular breathing, and respiration.</li>
	<li>Assess the anesthetic depth by the lack of response to the tail pinch or pedal reflexes withdrawal.</li>
	<li>Always recheck the anesthetic depth during the operative procedure and adjust the anesthetic depth if the animal is responsive to pain. Keep the mice on the pre-warmed surgery table to prevent hypothermia.</li>
	<li>Dilate recipient pupils with 1% (wt/vol) tropicamide eye drops 5 min before the surgery. A well-dilated pupil can facilitate transpupillary visualization under the operating microscope.</li>
	<li>Disinfect the operating mouse eye and surrounding ocular tissues with povidone-iodine, wash with sterilized PBS.</li>
	<li>Apply 0.5% Proparacaine hydrochloride on the mouse eye for analgesia.</li>
</ol>
6. Subretinal transplantation of retinal grafts (Figure 2)
NOTE: All the following steps were performed under aseptic conditions. Surgical tools were autoclaved (use instrument trays to protect the tool tips and take the plunger out of the micro syringes to prevent them getting stuck in the lumen). Supplier information of the research tools and products is provided in the Table of Materials.
<ol>
	<li>Anterior chamber paracentesis: Penetrate peripheral cornea into anterior chamber with an insulin needle to allow aqueous humor to passively egress, to prevent the intraocular pressure (IOP) spikes during transplantation. 
	NOTE: A successful paracentesis is indicated by the efflux of aqueous humor through the cornea hole.</li>
	<li>Put a drop of sodium hyaluronate and a glass coverslip (5 mm diameter) on top of the cornea to enable transpupillary visualization of the fundus under the surgical scope.</li>
	<li>Use toothed forceps to expose the injection locus (1 mm post-corneal limbus) by pressing either the superior or inferior eye wall, as required by the experiment, toward the center of transpupillary vision.</li>
	<li>Use a microinjection needle to perpendicularly penetrate the sclera.</li>
	<li>Carefully rotate the needle direction tangentially to allow complete penetration of the sclera-choroid-RPE complex into the subretinal space.</li>
	<li>Insert the needle tangentially to access the terminal injection locus. 
	NOTE: Verify the complete positioning of the needle&#39;s bevel by transpupillary vision guidance. Use the superficial retinal vessels as an anatomical reference for accurate needle positioning within the subretinal space.</li>
	<li>Inject retinal grafts 
	NOTE: The presence of small bubbles preloaded in the syringe can facilitate validation of the accurate subretinal positioning of donor cells. Elevated IOP resulting from the subretinal injection increases the risk of donor cell reflux.
	<ol>
		<li>If the cornea becomes cloudy24,25, an indicator of high IOP, keep the needle in the subretinal space for at least 3 min until the cornea clears, an indicator that the IOP has reduced sufficiently. 
		NOTE: In rare cases, the IOP may take longer to normalize, and it is recommended keeping the needle in place until the clarity of the cornea is restored even if this would take 8-10 min. Observe under a surgical microscope to prevent damage to retinal structures.</li>
	</ol>
	</li>
	<li>Grasp the injection hole edge with micro-toothed forceps and quickly withdraw the needle. 
	NOTE: The criteria for successful subretinal delivery of a retinal cell suspension and sheet are a constant bleb and a visible white sheet in the subretinal space, respectively.</li>
	<li>Clean the surgical area of the eye with artificial tears and apply ointment if necessary.</li>
	<li>Post-surgery administration
	<ol>
		<li>Place transplanted mice in a pre-warmed (37 &#176;C) clean recovery cage and carefully monitor for distress.</li>
		<li>Apply a drop of topical 0.5% Proparacaine Hydrochloride on the surgical eye 2-3 times if distress occurs.</li>
		<li>Return mice to the house cage after the mice are completely alert and mobile. Place a small number of food pellets and a gel cup in the cage if the mice have trouble reaching the food hopper. 
		NOTE: Keep the eye surface humid until anesthesia wears off to prevent cataract formation.</li>
	</ol>
	</li>
</ol>
7. Multimodal confocal scanning laser ophthalmoscopy (cSLO) imaging
<ol>
	<li>Two months post-transplantation, anesthetize recipient Rd1/NS mice (n=5) by intraperitoneal injection of ketamine (100 mg/kg body weight) and xylazine hydrochloride (20 mg/kg body weight) for imaging.</li>
	<li>Dilate pupils with 1% (wt/vol) tropicamide eye drops.</li>
	<li>Perform standard 55&#176; multimodal imaging using the confocal scanning laser ophthalmoscope cSLO system22.</li>
	<li>Obtain MR and SD-OCT images using 30 frames of ART averaging.</li>
</ol>
8. Histological staining
<ol>
	<li>Prepare frozen slides of transplanted Rd1/NS mice eyes as previously reported 14.</li>
	<li>Block and penetrate the frozen slides of transplanted retinae using a mixture of 5% goat serum and 1% Triton X100 in PBS.</li>
	<li>Incubate samples with primary antibodies over night at 4 &#176;C and rinse in PBS (5 min, 3x). The primary antibodies include goat anti-GFP-FITC (1:200), rabbit anti-recoverin (1:1000), and rabbit anti-S-opsin (1:500).</li>
	<li>Incubate the samples with secondary antibody (goat anti-rabbit Cy3, 1:500) and counterstained with DAPI. 
	NOTE: The suppliers of primary and secondary antibodies are listed in Table of Materials.</li>
</ol>

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MSS is/was a paid advisor to Revision Therapeutics, Johnson &#38; Johnson, Third Rock Ventures, Bayer Healthcare, Novartis Pharmaceuticals, W. L. Gore &#38; Associates, Deerfield, Trinity Partners, Kala Pharmaceuticals, and Acucela. MSS receives sponsored research support from Bayer. These arrangements have been reviewed and approved by the Johns Hopkins University in accordance with its conflict-of-interest policies. KVL is named as inventors on patent applications at University of Colorado. MSS and YVL are named as inventors on patent applications at Johns Hopkins University.

Subretinal transplantation in mice is technically challenging due to the small size of mouse eyes. In this study, we developed a simple and reproducible platform for subretinal transplantation in mouse recipients. The platform allows consistency in donor viability protection, successful subretinal delivery, and ensures a low complication rate.
The subretinal transplantation technique depicted here was developed based on the trans-scleral route, where the injection needle penetrates the outer layers (sclera-choroid-RPE complex) of the eyewall. Compared to the trans-corneal19,20 and trans-vitreous21,26,27&#160;approach, the trans-scleral approach directly enters the subretinal space without penetrating the anterior segment structures and neural retina, thus enabling a harmless transplantation and reducing the risk of donor cell reflux through the penetrating hole. A challenge of the trans-scleral approach is to ensure the needle tip propagates along the subretinal space before arriving at the terminal delivery site, while avoiding potential disturbance of the adjacent RPE and choroid layers. It is challenging to achieve these goals through a traditional trans-scleral approach28,29 without direct visual guidance. In this study, we developed a transpupillary vision-guided platform using a surgical microscope and external illumination of the retina to facilitate subretinal transplantation in mouse models. The platform allows the operator to track the surgical process and the recipients&#39; retinal condition by providing real-time visualization of the recipient fundus under the surgical microscope. For example, successful penetration of the sclera-choroid-RPE complex can be simply validated by a relatively bright reflectance of the needle tip via transpupillary visualization. Importantly, when guided by transpupillary visualization, the operator can accurately modulate the angle and the depth of injection according to the relative position between the needle and anatomic structures in the recipient retina (e.g., retinal vessels and optic nerve head). Moreover, accurate administration of materials using this technique can minimize RPE, choroid trauma, and other surgical complications. Indeed, we found hemorrhages can be minimized by avoiding maneuvers in close proximity to retinal blood vessels with the help of transpupillary visualization.
Reflux of donor cells is a major cause of surgical failure post-injection29,30. There are multiple factors involved in promoting donor cell reflux. The high intraocular pressure (IOP) of the recipient eyes caused by the injected donor cell mixtures is an important factor that promotes the reflux of cells out of the eye. Our protocol reduces the recipients&#39; IOP by anterior chamber penetration at the beginning of the surgery and enables the IOP to lower spontaneously by aqueous fluid egress before pulling out the vitreous needle from the vitreous cavity. A second factor is the unsealed injection tunnel in the recipients&#39; eyes. To prevent graft reflux back through the injection tunnel, we use a small size micro-injection needle (34G) to create a self-sealing tunnel when penetrating the eyewall and toothed micro-forceps to hold the edges of the external tunnel opening when pulling out the needle. In conventional transplantation, refluxed grafts are hardly detected because they may occur rapidly or dissipate quickly. The sodium hyaluronate, upon application of the coverslip, almost invariably slides off the cornea to cover most of the limbus and sclera including the sclerotomy site. Following cell injection, the sodium hyaluronate at the sclerotomy site can help to slow down the bulk flow of refluxed contents, if any, and help to make them visually detectable under the surgical microscope. Moreover, the absence of reflux can also be verified by tracing the constancy of the subretinal bleb for cell suspensions and the intraocular location of the retinal sheet using our protocol, which could be useful for laboratories where optical coherence tomography (OCT) is unavailable.
While a substantial number of photoreceptor cells survived in the transplanted retinal sheets, no discernible cellular orientation, or sheet orientation, could be detected. Although delivery of 3D retinal sheets to large animals has been established26,31,32, the subretinal space in mice, coupled with the lack of intraoperative retinal imaging capability, makes it very challenging to ensure the maintenance of sheet orientation intraoperatively as well as in the immediate postoperative period during which the transplanted cells or sheets may become unstable as the mouse regains ambulation.
We describe a trans-scleral surgical platform with direct transpupillary vision guidance for the subretinal transplantation in mouse recipients. This platform enables high accuracy of injection and a low rate of surgical complications. This platform enables precise delivery of known doses of cells, is relatively easy to learn, and facilitates subretinal delivery in addition to intra-retinal or intravitreal injections for different types of therapeutic agents including gene therapy.

Transplantation of photoreceptor and retinal pigmented epithelial (RPE) cells provides potential therapy for retinal degenerative diseases such as age-related macular degeneration (AMD), Stargardt disease, and retinitis pigmentosa (RP)1,2,3,4,5,6,7. To replenish or replace the diseased photoreceptors and RPE cells in degenerated retinae, the subretinal space is particularly well suited as a transplantation target given the laminar anatomy of host photoreceptors and RPE cells. While surgical procedures of sub-retinal transplantation of RPE cells are well-established in large animals8,9,10 and clinical trials11,12,13, challenges facing photoreceptor transplantation research include the scarcity of transgenic large animal models and the limited understanding of in-depth synaptogenesis mechanisms involved in neuronal transplantation, among other concerns. Genetically modified murine models, with various types of retinal degeneration mutants, provide useful tools for studying molecular mechanisms in the context of transplantation and steering the development of effective cell replacement therapies at the preclinical stage14,15,16,17,18.
Unlike the relatively large eye and small crystalline lens in large animals (e.g., pig, monkey), the small size and large crystalline lens of mouse eyes make them difficult surgical targets, especially for subretinal transplantation in which physical space constraints and limited direct visualization are the core challenges.
Current approaches can be classified into three major types based on the injection route. First, in the case of the trans-corneal approach, the needle is passed through the cornea into the vitreous cavity and then into the subretinal space19,20. Successful subretinal delivery can be achieved using this method but the damage to anterior segment structures (i.e., cornea, iris, lens) is a major risk that may severely impede downstream in vivo analysis. Second, in the case of the trans-vitreous approach, the needle enters the vitreous cavity through the pars plana and then into the subretinal space21. This approach is widely used in humans and large animals. However, there is a potential risk of lens damage in rodents as the lens occupies a larger relative volume of the vitreous cavity. Notably, both trans-corneal and trans-vitreous protocols require penetration of the neural retina to arrive at the subretinal space, which causes damage to the host retina and increases the risk of donor cell reflux through the penetration hole. Third, in the case of the trans-scleral approach22,23, the needle penetrates through the scleral-choroid-RPE complex and directly enters the subretinal space. This approach reduces the potential trauma of anterior segment structures and the vitreous cavity. However, without direct visualization of the host fundus, surgical failures caused by trans-retinal punctures, RPE detachment, and choroid hemorrhage are commonly detected.
Here, we developed a trans-scleral surgical platform with direct transpupillary vision guidance for subretinal transplantation in mouse recipients. Validation of the viability of donor retinal sheets and cell suspensions before transplantation was performed. Successful delivery of the donor cells into the subretinal space of a retina degeneration mouse model was confirmed. Only rare surgical complications were detected. In addition, transplanted photoreceptors in the retinal grafts survived and showed evidence of advanced maturation into &#34;adult&#34; rods and cones two months post-transplantation.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Artificial tears</td>
			<td>CareAll</td>
			<td>P31447-04</td>
			<td></td>
		</tr>
		<tr>
			<td>Coverslips (5mm in diameter)</td>
			<td>Deckglaser</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat anti-GFP (FITC)</td>
			<td>Abcam</td>
			<td>Ab6662</td>
			<td></td>
		</tr>
		<tr>
			<td>Goat-anti-rabbit Cy3&#160;</td>
			<td>Invitrogen</td>
			<td>A10520</td>
			<td></td>
		</tr>
		<tr>
			<td>Insulin syringe (30G)</td>
			<td>Easy Touch</td>
			<td>08496-3015-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Ketamine</td>
			<td>VETone Zetamine</td>
			<td>AH2017J</td>
			<td></td>
		</tr>
		<tr>
			<td>Live Dead Viability Kit</td>
			<td>Thermo Fisher Scientific</td>
			<td>L3224</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro scissor</td>
			<td>Harvard Apparatus</td>
			<td>72-8503</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro smooth forceps</td>
			<td>ASICO</td>
			<td>AE-4360</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro toothed forceps</td>
			<td>World Precision Instruments</td>
			<td>555041FT</td>
			<td></td>
		</tr>
		<tr>
			<td>Microinjection needle (26G)</td>
			<td>Hamilton</td>
			<td>7804-03</td>
			<td></td>
		</tr>
		<tr>
			<td>Microinjection needle (34G)</td>
			<td>Hamilton</td>
			<td>207434</td>
			<td></td>
		</tr>
		<tr>
			<td>Microinjection syringe</td>
			<td>Hamilton</td>
			<td>7633-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Papain dissociation kit</td>
			<td>Worthington Biochemical</td>
			<td>LK003150</td>
			<td></td>
		</tr>
		<tr>
			<td>Petri-dish (35 mm)</td>
			<td>Thermo Fisher Scientific</td>
			<td>FB012920</td>
			<td></td>
		</tr>
		<tr>
			<td>Povidone-iodine (10%)</td>
			<td>Betadine Solution</td>
			<td>N/A</td>
			<td></td>
		</tr>
		<tr>
			<td>Proparacaine Hydrochloride (0.5%)</td>
			<td>Keeler</td>
			<td>AX0500</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit anti-recoverin</td>
			<td>Millipore</td>
			<td>Ab5585</td>
			<td></td>
		</tr>
		<tr>
			<td>Rabbit anti-S-opsin</td>
			<td>Millipore</td>
			<td>Ab5407</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hyaluronate</td>
			<td>Johnson &#38; Johnson Vision&#160;</td>
			<td>10-2400-11</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile cotton swabs</td>
			<td>Puritan</td>
			<td>25-806 2PC</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile needle (25G)</td>
			<td>BD PrecisionGlide Needle</td>
			<td>305122</td>
			<td></td>
		</tr>
		<tr>
			<td>Sterile towel drapes</td>
			<td>Dynarex</td>
			<td>4410</td>
			<td></td>
		</tr>
		<tr>
			<td>Surgical materials/reagents</td>
			<td></td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Tropicamide ophthalmic solution</td>
			<td>Henry Schein</td>
			<td>112-7192</td>
			<td></td>
		</tr>
		<tr>
			<td>Xylazine</td>
			<td>AnaSed Injection</td>
			<td>N/A</td>
			<td></td>
		</tr>
	</tbody>
</table>

transpupillary-guided trans-scleral transplantation of subretinal grafts in a retinal degeneration mouse model

Transplantation of photoreceptor cells and retinal pigment&#160;epithelial (RPE) cells provide a potential therapy for retinal degeneration diseases. Subretinal transplantation of therapeutic donor cells into mouse recipients is challenging due to the limited surgical space allowed by the small volume of the mouse eye. We developed a trans-scleral surgical transplantation platform with direct transpupillary vision guidance to facilitate the subretinal delivery of exogenous cells in mouse recipients. The platform was tested using retinal cell suspensions and three-dimensional retinal sheets collected from rod-rich Rho::EGFP mice and cone-rich OPN1LW-EGFP;NRL-/- mice, respectively. Live/dead assay showed low cell mortality for both forms of donor cells. Retinal grafts were successfully delivered into the subretinal space of a mouse model of retinal degeneration, Rd1/NS, with minimum surgical complications as detected by multimodal confocal scanning laser ophthalmoscope (cSLO) imaging. Two months post-transplantation, histological staining demonstrated evidence of advanced maturation of the retinal grafts into &#39;adult&#39; rods and cones (by robust Rho::EGFP, S-opsin, and OPN1LW:EGFP expression, respectively) in the subretinal space. Here, we provide a surgical platform that can enable highly accurate subretinal delivery with a low rate of complications in mouse recipients. This technique offers precision and relative ease of skill acquisition. Furthermore, the technique could be used not only for studies of subretinal cell transplantation but also for other intraocular therapeutic studies including gene therapies.

Retinal grafts are successfully delivered into the subretinal space and survived in vivo. 
The performance of our subretinal transplantation platform was assessed in Rd1/NS recipient&#160;mice aged 6-8 weeks when their residual retinae were severely thinned due to the almost complete outer nuclear layer (ONL) degeneration. Given the fragility of the retina, the scarcity of cone photoreceptors, and the relatively poor viability of cells in suspension compared to sheet donors, cone-rich mice (OPN1LW-EGFP;NRL-/-)14 were used as the sources of donor retinal sheets and rod-rich mice (Rho::EGFP) as the donor for cell suspensions. Cone-rich retinal sheet and rod-rich cell suspensions were successfully delivered into the subretinal space of all recipient mice using our surgical platform, as confirmed by the subretinal bleb and white sheet seen using transpupillary visualization immediately following injection. Two months post-transplantation, multimodal cSLO imaging was performed to track the states of retinal grafts in vivo. Cross-sectional OCT scanning showed that retinal grafts survived in the subretinal space and reconstituted the ONL in all recipient mice (Figure 3A). Infrared imaging detected no obvious cataracts in all transplanted eyes (Figure 3B). Other surgical complications including hemorrhage were rarely detected in transplanted retinae (n = 1/10 eyes) by multicolor reflectance imaging at two months post-transplantation (Figure 3C). We performed histological staining to further assess the survival and extent of maturation of photoreceptors in retinal grafts. Abundant cone photoreceptors expressing OPN1LW:EGFP and S-opsin in transplanted retinal sheets were observed. Likewise, transplanted retinal cell suspensions showed a large proportion of Rec+ photoreceptors in vivo, including numerous Rho::EGFP+ rods (Figure 3D). The non-transplanted control mice showed severe ONL degeneration with sparse residual cone photoreceptors (Rec+). No EGFP signal was detected in non-transplanted Rd1/NS retina (Figure 3D).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65448/65448fig01.jpg" /> 
Figure 1: Schematic of collecting donor retinal sheet and cell suspensions. Schematics showing the key steps of collecting retinal cell suspensions and sheets from donor Rho::EGFP mice. Bright-field and fluorescent imaging showed representative images of dissociated cells and a dissected sheet isolated from Rho::EGFP mice. Yellow dotted line: incisional margin. <a href="https://www.jove.com/files/ftp_upload/65448/65448fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65448/65448fig02v3.jpg" /> 
Figure 2: Procedures of subretinal transplantation of retinal sheets and cell suspension in Rd1/NS mice. (A)&#160;Schematic images of&#160; recipient mice preparation. (B) Key surgical procedures and corresponding diagrams showing subretinal transplantation of retinal cell suspensions and sheets. Surgical steps include: 1. penetrate the anterior chamber; 2. drop the sodium hyaluronate on the cornea, then mount the coverslip on top of the sodium hyaluronate to facilitate the transpupillary visualization; 3. penetrate outer layers of the eye wall (sclera-choroid-RPE complex). Penetration angles of the needle are on the top right panel of the illustration; 4. insert injection needle; 5. inject grafts. Representative images show the successful delivery of retinal cell suspensions and sheet in two individual recipients, respectively. Asterisk: optic nerve head; Red arrowheads: retinal blood vessel; White arrowhead: penetrated needle tip; White arrows: grafted retinal suspensions or sheet. <a href="https://www.jove.com/files/ftp_upload/65448/65448fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65448/65448fig03.jpg" /> 
Figure 3: Successful delivery of retinal grafts into the subretinal space and survival in vivo. (A)&#160;Representative SD-OCT images showed the subretinal distribution of transplanted retinal sheets and cell suspensions in two individual Rd1/NS mice, respectively. Non-transplanted Rd1/NS mice were collected as a control. Region of Interest (ROI) is indicated by yellow-dotted boxes on infrared (IR) fundus images. Inner retina is designated as laminae of retinal ganglion cell layer (RGC), inner plexiform layer (IPL), and inner nuclear layer (INL). (B) Representative infrared (IR) image of a transplanted Rd1/NS mouse showed no cataract through the dilated pupil. (C) Representative multicolor reflectance (MR) images of transplanted and non-transplanted Rd1/NS eyes. The transplanted eyes presented no obvious surgical complications including hemorrhage. (D) Immunohistochemistry (IHC) staining of transplanted- (sheet and suspension) and non-transplanted (control) Rd1/NS mice. The data showed numerous grafted photoreceptors expressing specific markers of photoreceptors (Rec), rods (Rho::EGFP), L/M cones (OPN1LW:EGFP), and S-cones (S-opsin) at two months post-transplantation. Recipient retinal laminae were identified by DAPI staining (blue). Retinal grafts were identified by EGFP reporter (green). The non-transplanted mice retina showed severe ONL degeneration with sparse residual cone photoreceptors (Rec+). No EGFP signal was detected in non-transplanted Rd1/NS retinae. Magnified images were presented on the right two panels. Abbreviations: RGC: retinal ganglion cell layer; INL: inner nuclear layer. ONL: outer nuclear layer. <a href="https://www.jove.com/files/ftp_upload/65448/65448fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Transpupillary-Guided Trans-Scleral Transplantation of Subretinal Grafts in a Retinal Degeneration Mouse Model at JoVE.com

Transpupillary-Guided Trans-Scleral Transplantation of Subretinal Grafts in a Retinal Degeneration Mouse Model

This protocol presents a transpupillary vision-guided trans-scleral approach to safely and precisely deliver subretinal cellular grafts, with a low rate of surgical complications, in mouse recipients with or without retinal degeneration.

Transplantation of photoreceptor cells and retinal pigment&#160;epithelial (RPE) cells provide a potential therapy for retinal degeneration diseases. ...

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1.1K Aufrufe.  Johns Hopkins University School of Medicine. Das übergeordnete Ziel dieser Studie ist es, eine transsklerale Transplantationsplattform mit direkter transpupillärer Führung zu entwickeln, um die subretinale Verabreichung von Zellen bei Mausempfängern zu erleichtern. Alle Tierversuche wurden gemäß dem Leitfaden des National Institute of Health durchgeführt. Unsere Erklärung zur Verwendung von Tieren, die vom Animal Care and Use Committee der Johns Hopkins University genehmigt wurde.Die Annahme des EGFP-Alters von Mäusen am...

Serie: Transpupillary-Guided Trans-Scleral Transplantation of Subretinal Grafts in a Retinal Degeneration Mouse Model

Patch Clamp Recordings from Mouse Retinal Neurons in a Dark-adapted Slice Preparation

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of intracellular events that lead to closure of cyclic-nucleotide-gated channels in the cell membrane. The resulting change in membrane potential leads in turn to reductions in the amount of neurotransmitter release from both rod and cone synaptic terminals. To measure how the light-evoked change in photoreceptor membrane potential leads to downstream activity in the retina, scientists have made electrophysiological recordings from retinal slice preparations for decades1,2. In the past these slices have been cut manually with a razor blade on retinal tissue that is attached to filter paper; in recent years another method of slicing has been developed whereby retinal tissue is embedded in low gelling temperature agar and sliced in cool solution with a vibrating microtome3,4. This preparation produces retinal slices with less surface damage and very robust light-evoked responses. Here we document how this procedure can be done under infrared light to avoid bleaching the visual pigment.

Here we describe a procedure for generating dark-adapted slices of the mouse retina for electrophysiological recordings.

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of ...

Dissection of a Mouse Eye for a Whole Mount of the Retinal Pigment Epithelium

The retinal pigment epithelium (RPE) lies at the back of the mammalian eye, just under the neural retina, which contains the photoreceptors (rods and cones). The RPE is a monolayer of pigmented cuboidal cells and associates closely with the neural retina just above it. This association makes the RPE of great interest to researchers studying retinal diseases. The RPE is also the site of an in vivo assay of homology-directed DNA repair, the pun assay. The mouse eye is particularly difficult to dissect due to its small size (about 3.5mm in diameter) and its spherical shape. This article demonstrates in detail a procedure for dissection of the eye resulting in a whole mount of the RPE. In this procedure, we show how to work with, rather than against, the spherical structure of the eye. Briefly, the connective tissue, muscle, and optic nerve are removed from the back of the eye. Then, the cornea and lens are removed. Next, strategic cuts are made that result in significant flattening of the remaining tissue. Finally, the neural retina is gently lifted off, revealing an intact RPE, which is still attached to the underlying choroid and sclera. This whole mount can be used to perform the pun assay or for immunohistochemistry or immunofluorescent assessment of the RPE tissue.

A formal demonstration of the dissection of a mouse eye, resulting in a whole mount of the retinal pigment epithelium.

The retinal pigment epithelium (RPE) lies at the back of the mammalian eye, just under the neural retina, which contains the photoreceptors (rods and ...

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural connectivity form during development. In mice, retinofugal projections are arranged in a topographic manner and form eye-specific layers in the Lateral Geniculate Nucleus (dLGN) of the thalamus and the Superior Colliculus (SC). The development of these precise patterns of retinofugal projections has typically been studied by labeling populations of RGCs with fluorescent dyes and tracers, such as horseradish peroxidase1-4. However, these methods are too coarse to provide insight into developmental changes in individual RGC axonal arbor morphology that are the basis of retinotopic map formation. They also do not allow for the genetic manipulation of RGCs.
Recently, electroporation has become an effective method for providing precise spatial and temporal control for delivery of charged molecules into the retina5-11. Current retinal electroporation protocols do not allow for genetic manipulation and tracing of retinofugal projections of a single or small cluster of RGCs in postnatal mice. It has been argued that postnatal in vivo electroporation is not a viable method for transfecting RGCs since the labeling efficiency is extremely low and hence requires targeting at embryonic ages when RGC progenitors are undergoing differentiation and proliferation6. 
In this video we describe an in vivo electroporation protocol for targeted delivery of genes, shRNA, and fluorescent dextrans to murine RGCs postnatally. This technique provides a cost effective, fast and relatively easy platform for efficient screening of candidate genes involved in several aspects of neural development including axon retraction, branching, lamination, regeneration and synapse formation at various stages of circuit development. In summary we describe here a valuable tool which will provide further insights into the molecular mechanisms underlying sensory map development.

Transfection of Mouse Retinal Ganglion Cells by in vivo Electroporation

We demonstrate an in vivo electroporation protocol for transfecting single or small clusters of retinal ganglion cells (RGCs) and other retinal cell types in postnatal mice over a wide range of ages. The ability to label and genetically manipulate postnatal RGCs in vivo is a powerful tool for developmental studies.

The targeting and refinement of RGC projections to the midbrain is a popular and powerful model system for studying how precise patterns of neural ...

Implementing Dynamic Clamp with Synaptic and Artificial Conductances in Mouse Retinal Ganglion Cells

Ganglion cells are the output neurons of the retina and their activity reflects the integration of multiple synaptic inputs arising from specific neural circuits. Patch clamp techniques, in voltage clamp and current clamp configurations, are commonly used to study the physiological properties of neurons and to characterize their synaptic inputs. Although the application of these techniques is highly informative, they pose various limitations. For example, it is difficult to quantify how the precise interactions of excitatory and inhibitory inputs determine response output. To address this issue, we used a modified current clamp technique, dynamic clamp, also called conductance clamp 1, 2, 3 and examined the impact of excitatory and inhibitory synaptic inputs on neuronal excitability. This technique requires the injection of current into the cell and is dependent on the real-time feedback of its membrane potential at that time. The injected current is calculated from predetermined excitatory and inhibitory synaptic conductances, their reversal potentials and the cell's instantaneous membrane potential. Details on the experimental procedures, patch clamping cells to achieve a whole-cell configuration and employment of the dynamic clamp technique are illustrated in this video article. Here, we show the responses of mouse retinal ganglion cells to various conductance waveforms obtained from physiological experiments in control conditions or in the presence of drugs. Furthermore, we show the use of artificial excitatory and inhibitory conductances generated using alpha functions to investigate the responses of the cells.

This video article illustrates the set-up, the procedures to patch cell bodies and how to implement dynamic clamp recordings from ganglion cells in whole-mount mouse retinae. This technique allows the investigation of the precise contribution of excitatory and inhibitory synaptic inputs, and their relative magnitude and timing to neuronal spiking.

Ganglion cells are the output neurons of the  retina and their activity reflects the integration of multiple synaptic inputs  arising from specific neural ...

Trypsin Digest Protocol to Analyze the Retinal Vasculature of a Mouse Model

Trypsin digest is the gold standard method to analyze the retinal vasculature 1-5. It allows visualization of the entire network of complex three-dimensional retinal blood vessels and capillaries by creating a two-dimensional flat-mount of the interconnected vascular channels after digestion of the non-vascular components of the retina. This allows one to study various pathologic vascular changes, such as microaneurysms, capillary degeneration, and abnormal endothelial to pericyte ratios. However, the method is technically challenging, especially in mice, which have become the most widely available animal model to study the retina because of the ease of genetic manipulations 6,7. In the mouse eye, it is particularly difficult to completely remove the non-vascular components while maintaining the overall architecture of the retinal blood vessels. To date, there is a dearth of literature that describes the trypsin digest technique in detail in the mouse. This manuscript provides a detailed step-by-step methodology of the trypsin digest in mouse retina, while also providing tips on troubleshooting difficult steps.

Trypsin digest is one of the most commonly used methods to analyze retinal vasculature. This manuscript describes the method in detail, including key alterations to optimize the technique and remove the non-vascular tissue while preserving the overall architecture of the vessels.

Trypsin digest is the gold standard method to analyze the retinal vasculature 1-5. It allows visualization of the entire network of complex ...

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

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

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

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

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 ...

Modeling Neuronal Death and Degeneration in Mouse Primary Cerebellar Granule Neurons

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their post-natal development, abundance, and accessibility, CGNs are an ideal model to study neuronal processes, including neuronal development, neuronal migration, and physiological neuronal activity stimulation. In addition, CGN cultures provide an excellent model for studying different modes of cell death including excitotoxicity and apoptosis. Within a week in culture, CGNs express N-methyl-D-aspartate (NMDA) receptors, a specific ionotropic glutamate receptor with many critical functions in neuronal health and disease. The addition of low concentrations of NMDA in conjunction with membrane depolarization to rodent primary CGN cultures has been used to model physiological neuronal activity stimulation while the addition of high concentrations of NMDA can be employed to model excitotoxic neuronal injury. Here, a method of isolation and culturing of CGNs from 6 day old pups as well as genetic manipulation of CGNs by adenoviruses and lentiviruses are described. We also present optimized protocols on how to stimulate NMDA-induced excitotoxicity, low-potassium-induced apoptosis, oxidative stress and DNA damage following transduction of these neurons.

This protocol describes a simple method for isolating and culturing primary mouse cerebral granule neurons (CGNs) from 6-7 day old pups, efficient transduction of CGNs for loss and gain of function studies, and modelling NMDA-induced neuronal excitotoxicity, low-potassium-induced cell death, DNA-damage, and oxidative stress using the same culture model.

Cerebellar granule neurons (CGNs) are a commonly used neuronal model, forming an abundant homogeneous population in the cerebellum. In light of their ...

M&#252;ller Glia Cell Activation in a Laser-induced Retinal Degeneration and Regeneration Model in Zebrafish

A fascinating difference between teleost and mammals is the lifelong potential of the teleost retina for retinal neurogenesis and regeneration after severe damage. Investigating the regeneration pathways in zebrafish might bring new insights to develop innovative strategies for the treatment of retinal degenerative diseases in mammals. Herein, we focused on the induction of a focal lesion to the outer retina in adult zebrafish by means of a 532 nm diode laser. A localized injury allows investigating biological processes that take place during retinal degeneration and regeneration directly at the area of damage. Using non-invasive optical coherence tomography (OCT), we were able to define the location of the damaged area and monitor subsequent regeneration in vivo. Indeed, OCT imaging produces high-resolution, cross-sectional images of the zebrafish retina, providing information which was previously only available with histological analyses. In order to confirm the data from real-time OCT, histological sections were performed and regenerative response after the induction of the retinal injury was investigated by immunohistochemistry.

M&amp;#252;ller Glia Cell Activation in a Laser-induced Retinal Degeneration and Regeneration Model in Zebrafish

The zebrafish is a popular animal model to study mechanisms of retinal degeneration/regeneration in vertebrates. This protocol describes a method to induce localized injury disrupting the outer retina with minimal damage to the inner retina. Subsequently, we monitor in vivo the retinal morphology and the M&#252;ller glia response throughout retinal regeneration.

A fascinating difference between teleost and mammals is the lifelong potential of the teleost retina for retinal neurogenesis and regeneration after ...

Partial Optic Nerve Transection in Rats: A Model Established with a New Operative Approach to Assess Secondary Degeneration of Retinal Ganglion Cells

Previous studies have shown that the secondary degeneration of retinal ganglion cells (RGCs) occurs commonly in glaucoma. Partial optic nerve transection is considered a useful and reproducible model. Compared with other optic nerve injury models used commonly for assessing secondary degeneration, e.g. complete optic nerve transection and optic nerve crush models, the partial optic nerve transection model is superior as it distinguishes primary from secondary degeneration in situ. Therefore, it serves as an excellent tool for evaluating secondary degeneration. This study describes a novel operative approach of partial optic nerve transection by directly accessing the area of the retrobulbar optic nerve through the orbital lateral wall of the eyeball. Moreover, we present a newly designed, low cost surgical instrument to assist with transection. As demonstrated by the representative results in distinguishing the boundary of primary and secondary injury areas, the new approach and instrument ensures high efficiency and stability of the model by providing adequate space for surgical operation. This in turn makes it easy to separate the meningeal sheath and ophthalmic vessels from the optic nerve before transection. An additional benefit is that this space-saving operative approach improves the investigators&#39; ability to administer drugs, carriers, or selective RGC tracers to the stump of the partially transected optic nerve, allowing the exploration of mechanisms behind secondary injury in RGCs, in a new way.

Secondary degeneration of retinal ganglion cells (RGCs) occurs commonly in glaucoma. This study describes an innovative operative approach for partial optic nerve transection. The use of this space-saving operative approach extends the model's application range, and allows exploration of secondary injury mechanisms in RGCs in a new way.

Previous studies have shown that the secondary degeneration of retinal ganglion cells (RGCs) occurs commonly in glaucoma. Partial optic nerve transection ...