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

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

1. 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.
2. Use OPN1LW-EGFP/NRL^-/- mice¹⁴ (aged P3) as donors of retinal sheets.
3. Use adult retinal degeneration Rd1/NS mice with immune deficiency (either sex) as recipients.
4. House all mice in cages under a 12:12 h light-dark cycle with access to water and food as needed.

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.

1. Donor mice preparation: Euthanize donor mice with an overdose of carbon dioxide and confirm euthanasia with cervical dislocation.
2. Isolate mice eyeballs. To do so follow the steps below.
   1. Carefully open pups' eyelids using micro scissors to expose the eyeball.
   2. Use smooth forceps to grab the optic nerve and pull out the eyeball with a pair of forceps.
3. Isolate neural retina
   NOTE: This step is performed under a dissection microscope.
   1. Incise a hole in the center of the cornea using a 25 G sterile needle.
   2. Cut the cornea, through the hole, in half and enlarge the incision to the sclera and RPE, then remove the sclera and RPE.
   3. Use micro-toothed forceps to gently remove the lens and the vitreous to isolate the neural retina.
   4. Proceed with section 3 to prepare donor retinal suspension or section 4 to prepare donor retinal sheet, as required.

3. Prepare donor retinal suspension (Figure 1)

1. Incubate the neural retina in Papain solution at 37 °C for 20-30 min until no cell clumps are detected.
   NOTE: Gently pipette the cell mixture 15x every 10 min.
2. Follow manufacturer's instructions of the Papain Dissociation Kit to collect single cells.

4. Prepare donor retinal sheets (Figure 1)

NOTE: This step follows section 2 and is performed under a dissection microscope.

1. Put the isolated neural retina cup into a 35 mm Petri dish with prechilled sterile PBS.
2. Use micro scissors to cut the neural retina cup into multiple retinal sheets (around 1 mm x 2 mm).

5. Prepare recipient mice

1. Anesthetize recipient mice with intraperitoneal injection of ketamine (100 mg/kg body weight) and xylazine hydrochloride (20 mg/kg body weight).
2. Confirm the surgical plane of anesthesia by ensuring the loss of blink and pain reflexes, regular breathing, and respiration.
3. Assess the anesthetic depth by the lack of response to the tail pinch or pedal reflexes withdrawal.
4. 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.
5. 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.
6. Disinfect the operating mouse eye and surrounding ocular tissues with povidone-iodine, wash with sterilized PBS.
7. Apply 0.5% Proparacaine hydrochloride on the mouse eye for analgesia.

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.

1. 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.
2. 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.
3. 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.
4. Use a microinjection needle to perpendicularly penetrate the sclera.
5. Carefully rotate the needle direction tangentially to allow complete penetration of the sclera-choroid-RPE complex into the subretinal space.
6. Insert the needle tangentially to access the terminal injection locus.
   NOTE: Verify the complete positioning of the needle's bevel by transpupillary vision guidance. Use the superficial retinal vessels as an anatomical reference for accurate needle positioning within the subretinal space.
7. 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.
   1. If the cornea becomes cloudy^24,25, 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.
8. 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.
9. Clean the surgical area of the eye with artificial tears and apply ointment if necessary.
10. Post-surgery administration
    1. Place transplanted mice in a pre-warmed (37 °C) clean recovery cage and carefully monitor for distress.
    2. Apply a drop of topical 0.5% Proparacaine Hydrochloride on the surgical eye 2-3 times if distress occurs.
    3. 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.

7. Multimodal confocal scanning laser ophthalmoscopy (cSLO) imaging

1. 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.
2. Dilate pupils with 1% (wt/vol) tropicamide eye drops.
3. Perform standard 55° multimodal imaging using the confocal scanning laser ophthalmoscope cSLO system²².
4. Obtain MR and SD-OCT images using 30 frames of ART averaging.

8. Histological staining

1. Prepare frozen slides of transplanted Rd1/NS mice eyes as previously reported ¹⁴.
2. Block and penetrate the frozen slides of transplanted retinae using a mixture of 5% goat serum and 1% Triton X100 in PBS.
3. Incubate samples with primary antibodies over night at 4 °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).
4. 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.