Subretinal Transplantation of Human Embryonic Stem Cell-Derived Retinal Tissue in a Feline Large Animal Model

Podsumowanie

Presented here is a surgical technique for transplanting human pluripotent stem cell (hPSC)-derived retinal tissue into the subretinal space of a large animal model.

Streszczenie

Retinal degenerative (RD) conditions associated with photoreceptor loss such as age-related macular degeneration (AMD), retinitis pigmentosa (RP) and Leber Congenital Amaurosis (LCA) cause progressive and debilitating vision loss. There is an unmet need for therapies that can restore vision once photoreceptors have been lost. Transplantation of human pluripotent stem cell (hPSC)-derived retinal tissue (organoids) into the subretinal space of an eye with advanced RD brings retinal tissue sheets with thousands of healthy mutation-free photoreceptors and has a potential to treat most/all blinding diseases associated with photoreceptor degeneration with one approved protocol. Transplantation of fetal retinal tissue into the subretinal space of animal models and people with advanced RD has been developed successfully but cannot be used as a routine therapy due to ethical concerns and limited tissue supply. Large eye inherited retinal degeneration (IRD) animal models are valuable for developing vision restoration therapies utilizing advanced surgical approaches to transplant retinal cells/tissue into the subretinal space. The similarities in globe size, and photoreceptor distribution (e.g., presence of macula-like region area centralis) and availability of IRD models closely recapitulating human IRD would facilitate rapid translation of a promising therapy to the clinic. Presented here is a surgical technique of transplanting hPSC-derived retinal tissue into the subretinal space of a large animal model allowing assessment of this promising approach in animal models.

Wprowadzenie

Millions of people around the world are impacted by retinal degeneration (RD) with resulting visual impairment or blindness associated with loss of the light-sensing photoreceptors (PRs). Age-related macular degeneration (AMD) is a major cause of blindness resulting from a combination of genetic risk factors and environmental/lifestyle factors. In addition, over 200 genes and loci have been found to cause inherited RD (IRD)¹. Retinitis pigmentosa (RP), the commonest IRD, is genetically heterogenous with more than 3,000 genetic mutations in approximately 70 genes being reported^2,3,4. Leber Congenital Amaurosis (LCA), which causes blindness in childhood is also genetically heterogenous^5,6. Gene augmentation therapy has been developed and is in clinical trials for treating a small number of IRDs^3,7. However, a separate therapy must be developed for the treatment of each distinct genetic form of IRD and thereby only treating a small subset of patients. Furthermore, gene augmentation relies on the presence of a population of rescuable photoreceptors and is, therefore, not applicable for advanced degeneration.

There is, therefore, an urgent and yet unmet clinical need for the development of therapies addressing and treating advanced RDs and profound to terminal blindness. Over the last 2 decades neuroprosthetic implants have been developed and tested in large animal models, such as the cat, prior to human use^{8,9,10,11,12,13,14}. Likewise, in the past 20 years retinal replacement therapies utilizing sheets of embryonic or even mature mammalian retina grafted subretinally have been developed^{15,16,17,18,19,20,21,22} and even tested successfully in RD patients^23,24,25. Both approaches utilize the idea of introducing new sensors (photovoltaic silicon photodiodes in the case of neuroprosthetic devices^26,27, and healthy mutation-free photoreceptors organized in sheets, in the case of retinal sheet implantation) into retina with degenerated PRs. Recent studies have investigated the use of stem cells-based approaches such as transplantation of human pluripotent stem cell (hPSC)-derived retinal progenitors^28,29, hPSC-photoreceptors³⁰, and hPSC-retinal organoids^31,32,33. Retinal organoids enable the formation of retinal tissue in a dish and derivation of photoreceptor sheets with thousands of mutation-free PRs, which resemble the photoreceptor layer in the developing human fetal retina^{34,35,36,37,38,39,40}. Transplanting hPSC-derived retinal tissue (organoids) into the subretinal space of patients with RD conditions is one of the new and promising investigational cell therapy approaches, being pursued by a number of teams^31,32,41,42. Compared to transplantation of the cell suspension (of young photoreceptors or retinal progenitors), transplanted sheets of fetal photoreceptors were demonstrated to result in vision improvements in clinical trials^23,24.

The protocol presented here describes, in detail, a transplantation procedure for subretinal delivery of the whole retinal organoids (rather than organoid rims^33,41) as a potentially better way to introduce intact retinal sheets with PRs, to increase graft survival and improve the sheet preservation. Though procedures for introducing a flat piece of human retina and also RPE patches have been developed^43,44,45, transplantation of larger 3D grafts has not been investigated. Stem cell-derived retinal organoids provide an inexhaustible source of photoreceptor sheets for developing vision restoration technologies, are free of ethical restriction, and are considered an excellent source of human retinal tissue for therapies focused on treating advanced RD and terminal blindness⁴⁶. Development of surgical methods for precise subretinal implantation of retinal organoids with minimal injury to the host retinal niche (neural retina, retinal pigment epithelium and retinal and choroidal vasculature) is one of the critical steps for advancing such therapy toward clinical applications^31,32. Large animal models such as cats, dogs, pigs, and monkeys have proven to be good models for investigating surgical delivery methods as well as to demonstrate the safety of implanted sheets of tissue (retinal pigment epithelium (RPE) cells) and investigate the use of organoids^{41,44,45,47,48,49,50}. The large animal eye has a similar globe size to human as well as similar anatomy including the presence of a region of high photoreceptor density, including cones (the area centralis), resembling the human macula^6,51,52.

In this manuscript, a technique for the implantation of hPSC-derived retinal tissue (organoids) into the subretinal space of feline large animal models (both wild-type and Crx^Rdy/+ cats) is described, which, together with promising efficacy results^32,53 builds a foundation for further development of such investigational therapy toward clinical applications to treat RD conditions.

Protokół

Procedures were conducted in compliance with the Association for Research in Vision and Ophthalmology (ARVO) statement for Use of Animals in Ophthalmic and Vision Research. They were also approved by the Michigan State University Institutional Animal Care and Use Committee. Wild-type and Crx^Rdy/+ cats from a colony of cats maintained at Michigan State University were used in this study. Animals were housed under 12 h : 12 h light-dark cycles and fed a commercial complete cat diet.

1. Pre-implantation procedures and surgical set-up

1. Select wild-type or Crx^Rdy/+ cats depending on the study design. Perform a pre-surgical ophthalmic examination, including slit lamp biomicroscopy and indirect ophthalmoscopy. Exclude any animals with fundus abnormalities not related to their genotypes.
2. One week prior to implantation, start the animals on an immunosuppressive protocol of oral cyclosporine 2 mg/kg and prednisolone 1 mg/kg both twice daily to help prevent transplant rejection.
3. Fast the animals over 4 months of age overnight (at least 8 h). Provide a limited amount of wet food overnight to the animals less than 4 months of age, and then, fast them for 2 h prior to the surgery.
4. Perform a general physical examination, including chest auscultation (using a stethoscope). Record the heart and respiratory rate, temperature, mucous membrane color, and capillary refill time.
5. Pre-medicate the animals less than 4 months of age with buprenorphine (0.02 mg/kg), and those over 4 months of age with buprenorphine (0.02 mg/kg) combined with acepromazine (0.02 mg/kg) subcutaneously or intramuscularly 30–45 min prior to general anesthesia induction.
6. Apply topical 1% tropicamide ophthalmic solution and 10% phenylephrine ophthalmic solution to the ocular surface at least twice to dilate the pupils. Repeat if pupils are not well-dilated.
7. Place a 22 G intravenous catheter in the cephalic vein in all the animals: first clip hair from a 2 x 3 cm area over the cephalic vein; prepare the skin by first scrubbing it with 70% ethanol and then with a chlorhexidine scrub; place the catheter and secure with medical tape and flush with heparinized saline. In animals under 4 months, this can be done following induction of anesthesia.
8. Induce general anesthesia 30–45 min after premedication: animals under 4 months of age are induced with isoflurane delivered by mask, those over 4 months of age are induced using intravenous propofol (4–6 mg/kg).
9. Intubate with an appropriate size of endotracheal tube. Visualize the larynx with an examination light or laryngoscope. Spray 0.1 mL of lidocaine 2% on to the larynx, wait for a few seconds and then intubate.
10. Maintain anesthesia with isoflurane (between 2%–3.5%) in oxygen (300–600 mL/kg/min) via a Bain system.
11. Place the animal in dorsal recumbency on a positioning pillow on which a heated water blanket covered by towels is positioned. Attach a patient monitor and monitor the heart rate and electrocardiogram (ECG), respiratory rate, blood pressure, oxygen saturation, and end-tidal carbon dioxide. Regularly monitor the body temperature during the procedure. A suitably trained person is responsible for the maintenance and monitoring of anesthesia.
12. Position the animal with the help of the positioning pillow such that the corneal surface is horizontal when the eye is rotated into a primary gaze position. Secure the head in place using medical tape.
13. Cover the animal with blankets to help maintain the body temperature.
14. Flush the intravenous catheter with heparinized saline and start an intravenous infusion of Ringer lactate supplied at 2–5 mL/kg/h for the duration of the procedure.
15. Prepare the ocular surface, conjunctival sac, and eyelids for aseptic surgery using 0.2% povidone-iodine, sterile cotton swab applicators, and cotton balls.
16. Position the operating microscope with eyepieces also adjusted appropriately. Place the foot control for the microscope (focus, zoom, and XY-axis control) and vitrectomy machine such that the surgeon can operate them.
17. Prepare for aseptic surgery: all personnel must wear surgical scrubs, a surgical hat, and a mask. Ensure that the surgeon and the assistant(s) scrub, gown, and glove as for routine aseptic surgery. All personnel should be familiar with aseptic techniques.
18. Once the personnel are scrubbed, gowned, and gloved, open the surgical packs, and lay out the instruments. Place sterile manipulation knobs on the microscope adjustment knobs to allow the surgeon or the assistant to adjust without breaking asepsis. Drape the animal in a routine fashion for ocular surgery.
19. Prepare the vitrectomy machine following the manufacturer’s instructions for a two-port vitrectomy.
    1. Place a vitrectomy contact lens and two port 23 G vitrectomy on the assistant’s table. Attach the wet field cautery. Attach and prime the infusion tubing and the 23 G vitrectomy handpiece.
    2. Place the vitrectomy instruments and fluid lines over the sterile drape and maintain them in position using towel clamps. Set the vitrectomy machine in Proportional Vacuum mode between 1,500 and 2,500 cuts per minute (cpm) and maximum vacuum of 500 mmHg. This is performed by pressing on the arrow buttons (upward or downward) present on the front panel of the vitrectomy machine.

2. Preparation of the organoids for subretinal implantation (Figure 1)

1. Derive retinal organoids as outlined previously³¹, and, if needed, ship overnight at 37 °C as previously reported⁵⁴.
2. Wipe down the surfaces in the tissue culture room in the receiving laboratory with a disinfectant and maintain the same high level of aseptic technique as in a surgical suite for ocular surgeries, to avoid bacterial or fungal contamination, and carry over into the surgical room to avoid surgical site infection.
3. Wear surgical shoe covers, disposable lab coats, bonnets, surgical masks, sleeves, and surgical scrubs inside the tissue culture room assigned for organoid preparation for ocular surgeries.
4. Pre-equilibrate neural media with 20 ng/mL of basic fibroblast growth factor (bFGF) and 20 ng/mL of brain-derived neurotrophic factor (BDNF) for 1 h in a tissue culture incubator (37 °C, 5% CO₂). Place organoids in media in ultra-low adhesion plates using approximately one-fourth volume of conditioned medium (from the overnight shipment) and three-fourth volume of fresh medium^31,54. Maintain for 24–48 h.
5. Prepare and equilibrate several fresh 60 mm plates with neural medium (as prepared in step 2.4) for at least 1 h in the tissue culture incubator (37 °C, 5% CO₂) before anesthetizing the first animal. Note, these plates are used for transferring the organoids for each individual surgery, with an assumption that pH conditions and CO₂ saturation will be maintained in the media for at least 20 min, sufficient for moving the plate to the surgical room and loading organoids into the cannula.
6. Use a fresh 60 mm plate with pre-saturated neural media for each transplantation case, while keeping the remaining plates in the tissue culture incubator at 37 °C.
7. Transfer 6–9 organoids into one 60 mm dish (with re-saturated media) by pipetting them out with a manual single-channel pipette (20–200 µL) with 200 µL sterile filter tip having a wide (~0.7 mm) opening. These tips can be prepared in advance or during the procedure by snipping off ~3-4 mm of the tip with a pair of sterile scissors (done in the tissue culture hood to avoid contamination). Place the 60 mm dish inside a 100 mm tissue culture dish (to avoid contamination during room-to-room transport) and quickly transport to the surgical suite.
8. Place the plate with organoids on a 37 °C electric heating pad covered by a sterile drape.
9. Change into a fresh pair of sterile gloves before prepping the organoids.
10. Rinse the organoids in sterile balanced salt solution (BSS) (optional), and then load into the injector, which consists of a thin-walled borosilicate glass cannula (outer diameter, OD 1.52 mm; inner diameter, ID 1.12 mm) with a polished blunt end attached by sterile plastic tubing to a sterile 500 µL Hamilton syringe pre-filled with sterile BSS.
11. Pass the cannula to the surgical team in a sterile fashion when they are ready to inject the organoids.
12. Keep the length of the entire procedure (from removing the organoids from tissue culture media to loading the cannula) to 20 min or less.
13. If there is a delay in implanting the organoids, place them back into the tissue culture incubator, to avoid changes in pH/CO₂ saturation inside the dish.
14. Retain unused organoids for 1 week in the same tissue culture incubator and monitor for any evidence of contamination.

3. Subretinal organoids implantation

1. Perform a 0.5–1 cm lateral canthotomy with Stevens tenotomy scissors. Place an appropriately sized Barraquer eyelid speculum to keep the eyelids open. Ensure that the surgical assistant irrigates the cornea regularly with BSS for the duration of the procedure.
2. Place 2 stay sutures of 6–0 silk suture in the conjunctiva immediately adjacent to the limbus at the 4 and 8 o’clock positions to hold the eye in primary gaze and retract the third eyelid. Use 0.5 Castroviejo corneal tying forceps and a small mosquito hemostat to gently grasp the bulbar conjunctiva next to the limbus. Leave the sutures long and clamp the ends with small mosquito hemostats to help with their manipulation.
   NOTE: Placing the suture under the third eyelid is more difficult; be sure to place it immediately adjacent to the limbus.
3. Place another suture at the 12 o’clock position at the limbus partially through the thickness of the limbus taking care not to penetrate the eye. Knot this suture loosely and cut the ends short. This provides a robust anchor suture, which is used during the surgery as a “handle” to rotate the eye to allow the surgeon access to different parts of the globe during different stages of the procedure.
   NOTE: Steps 3.2 and 3.3 can be inverted. The sequence of stay sutures placement is the surgeon's preference.
4. Reflect the bulbar conjunctiva between 10–2 o’clock (a perimetry). Using tenotomy scissors, incise the conjunctiva 2–3 mm from the limbus. Undermine it and clear the tenon’s capsule to expose the sclera at the sites for the 2 and 10 o’clock vitrectomy ports, which will be placed about 3–5 mm from the limbus depending on the age of the animal. Use surgical cellulose spears for hemostasis and to clear any blood.
5. Identify sites for sclerotomy following the reflection of the conjunctiva and tenon’s capsule using calipers. Choose sclerotomy sites (at 2 and 10 o’clock aiming to pass through pars plana), to avoid the major scleral vessels that can be prominent in the cat. Use wet-field cautery on the sclera at the planned sclerotomy sites to reduce scleral bleeding, planning for an ~3 mm region at the instrument port (10 o’clock port for a right-handed surgeon) as this will be enlarged following the vitrectomy to allow the implantation cannula to be introduced.
6. Pre-place cruciate pattern sutures (6-0 or 7-0 polyglactin suture) without tying the knot at the site of the proposed sclerotomies before the vitrectomy ports are placed.
   NOTE: This facilitates rapid closure of the sclerotomies at the end of the procedure. The suture for the planned instrument port needs to have a longer bite of sclera as this will be a long incision.
7. Once the sutures are placed, ask the assistant to help position the globe by holding the 12 o’clock stay sutures by tying or using Bishop-Harmon forceps. Let the surgeon stabilize the globe as well, using a 0.12 mm Castroviejo forceps to hold the tissue next to the sclerotomy site.
   1. Introduce a 23 G vitrectomy port using a trocar through the sclera at both the 2 o’clock and 10 o’clock positions directed at an angle toward the optic nerve to avoid contacting the lens.
   2. Check that the irrigation port is in the vitreous by gently pushing on the port using tying forceps so that the tip can be visualized in the vitreous. Once correct positioning is confirmed, attach the irrigation line to the port and position the line using thin adhesive bandages to tape it in place. Set the vitrectomy infusion of the BSS buffer initially at 30–35 mmHg by pressing on the arrow buttons (upward or downward) present on the front panel of the vitrectomy machine.
8. Let the assistant hold a Machemer magnifying irrigating vitrectomy lens onto the cornea to allow visualization of the posterior segment of the eye during the next stages. Attach the irrigating vitrectomy lens to a drip set providing constant fluid supply to couple the lens to the cornea.
   NOTE: Other forms of contact vitrectomy lens could also be used. Dim or switch off the room lights to help visualization through the operating microscope.
9. Insert the 23 G vitrectomy probe/cutter (2,500 cpm) through the instrument port (adjacent to the surgeon’s dominant hand, i.e., the 10 o’clock port for a right-handed surgeon) and perform a partial core vitrectomy.
   1. Then, completely remove the vitreous from the retinal surface in the region that will receive the transplant (this is important for the success of the procedure) by detaching the vitreal face from the retina.
   2. Place the vitrectomy probe over the optic nerve head with the port facing away from the retinal surface and apply higher vacuum to start the vitreal face detachment.
10. Prepare triamcinolone crystals for intraocular use. If the triamcinolone suspension is not specifically for intravitreal use, wash the crystals and then re-suspend in BSS.
    1. Initially, filter the suspension with the aid of a sterile 0.22 µm pore syringe filter with PES membrane (attached to a 1 mL syringe) to trap the crystals.
    2. Then wash the trapped triamcinolone crystals by aspirating BSS in the 1mL syringe and flushing through the filter (the crystals remain trapped in the filter). This removes the preservatives in the solution. Repeat the washing 3 times after which the crystals are re-suspended in 1 mL BSS.
11. Introduce the needle of the syringe holding the triamcinolone through the instrument port. Be careful not to touch the lens by watching the tip of the needle through the pupil while introducing the needle through the instrument port. Inject 0.25 to 0.5 mL of crystal suspension. Then insert the vitrectomy probe through the instrument port and advance it close to the optic nerve head with the port away from the retinal surface and use high vacuum to help detach the vitreal face from the retina.
    NOTE: The triamcinolone crystals stick to and thus highlight the remaining vitreous. Carefully remove any vitreous above and next to the site of planned organoid implantation (e.g., the central tapetal fundus close to the area centralis region).
12. Create a small focal retinal detachment bleb at the desired implantation site using a subretinal injector, e.g., 23 G subretinal injector with an extendable 41 G cannula attached to a sterile 250 µL gas tight Luer lock syringe filled with sterile BSS.
    1. Prior to creating the subretinal bleb, reduce the infusion pressure to 10 mmHg to facilitate bleb formation.
    2. Insert the injector through the instrument port and advance it toward the retinal surface. Extrude the cannula tip and gently press it on to the retinal surface. Ask the assistant to give a slight quick push on the syringe plunger to start the retinal detachment that reduces the injection pressure to permit a slow increase of the retinal detachment until the desired size is achieved (approximately 100 to 200 µL of BSS is used).
13. If the retinotomy created is at a suitable position, which avoids cutting the retina between the implantation site and the optic nerve head to prevent the sectioning of the nerve fiber layer derived from the implantation site and to avoid major retinal blood vessels (this is also determined by the study design, in our case the central retina was chosen), then, slightly enlarge it using the 41 G cannula of the injector, aiming to facilitate the introduction of retinal scissors.
14. Remove the injector from the eye and remove the 10 o’clock scleral port. Enlarge the sclerotomy at this site using a straight 2.85 mm slit knife/keratome oriented toward the optic nerve to avoid touching the lens. Maintain the infusion pressure at 10–15 mmHg.
15. Extend the retinotomy using vitreoretinal vertical 80° scissors with squeeze handle, avoiding cutting retinal vessels to prevent retinal hemorrhage and be sure to cut the retina at the edge of the bleb away from the optic nerve head to avoid cutting nerve fibers leading from the retina in the transplant area to the optic nerve. The retinotomy should be of sufficient width to receive the organoids.
16. Insert the pre-loaded glass capillary containing the organoids (see step 2.10) through the enlarged sclerotomy and advance it toward the retinotomy site under direct visualization.
    1. Use the tip of the glass capillary to slightly open the retinotomy to access the opening into the subretinal bleb.
    2. Ask the assistant to slowly press the plunger of the injector, while watching through the microscope, injecting the organoids into the subretinal bleb. BSS should precede the organoids and flush the retinotomy open.
    3. Gently push the organoids within the bleb with the tip of the glass capillary if they are at the edge of the retinotomy.
17. Hold the glass capillary at the retinotomy for a few seconds to try and close the retinotomy and prevent the organoids from escaping into the vitreous.
18. Very slowly remove the glass capillary from the eye avoiding any sudden release of fluid from the eye to avoid fluid movement within the eye that might expel the organoids from the subretinal bleb.
19. Slowly increase the infusion pressure to 20–30 mmHg to help prevent intraocular hemorrhage taking care that the infusion fluid does not wash directly onto the bleb. This is performed by pressing on the upward arrow buttons present on the front panel of the vitrectomy machine.
20. Close the sclerotomy using the pre-placed suture in a cruciate pattern (6–0 or 7–0 polyglactin). Add additional simple interrupted sutures as needed to seal the sclerotomy.
21. Ask the assistant to remove the infusion port and let the surgeon quickly tie the pre-placed suture to seal the sclerotomy and prevent loss of intraocular pressure.
22. Close the conjunctival incision (peritomy) using 6–0 or 7–0 polyglactin in a simple continuous pattern.
23. Image the fundus (e.g., using a RetCam II video fundus camera, Clarity, or similar) and record the immediate position of the organoids within the subretinal space.
24. Re-prep the ocular surface after imaging using 0.2% povidone iodine solution with the aid of cotton tip applicators and cotton balls. Remove the three stay sutures and the lid speculum.
25. Close the lateral canthotomy with 6-0 polyglactin suture. Place a single buried suture followed by a figure of 8 skin sutures to reform the lateral canthus and then use simple interrupted skin sutures to close the rest of the wound.
26. After the surgical procedure is completed, give a subconjunctival injection of a steroid and antibiotic combination (2 mg methylprednisolone acetate, 0.1 mg dexamethasone, and 1 mg gentamicin). Place an ophthalmic lubricant (artificial tears) on the cornea.
27. Recover the animal from anesthesia and monitor closely during recovery for any signs of pain that would require additional treatment, such as marked blepharospasm, severe reluctance to be manipulated, lethargy or decreased appetite, and increased respiratory and heart rate. Provide post-operative analgesia as needed and monitor closely for any discomfort.
    NOTE: Only properly trained personnel should be in charge of anesthetizing and monitoring the feline patients before, during, and post-surgery. Follow local animal ethics committee recommendations for post-operative care.

4. Post-implantation procedures, post-operative treatment, and assessment

1. Continue to treat with oral immunosuppressive medications (1 mg/kg prednisolone and 2 mg/kg cyclosporine orally twice a day) to help control inflammation and rejection of the organoids. Provide systemic antibiotic coverage (e.g., oral doxycycline 5 mg/kg twice daily – this antibiotic was selected because of the risk of opportunist mycoplasmal infection in immunosuppressed animals).
2. Perform regular ophthalmic examinations to monitor for inflammation or the development of adverse events. Monitor the retinal bleb for flattening, which occurs over the first few days following surgery despite the large retinotomy required for organoids to be injected into the subretinal space. Record fundus images at each examination to record the position and appearance of the transplanted organoids.
3. Image the animals under general anesthesia by confocal scanning laser ophthalmoscopy (cSLO) and spectral domain – optical coherence tomography (SD-OCT)^5,31 during the monitoring period and prior to the termination of the experiment.
4. Humanely euthanize the animals at the termination of the experiment using an AVMA approved method, e.g., intravenous administration of 85 mg/kg of pentobarbital. After sedation, place an intravenous catheter for administration of the pentobarbital to minimize stress. Confirm the death by cessation of heartbeat and make an incision into the thorax to create a pneumothorax.
5. Remove the eyes via a standard transconjunctival approach and fix as required. For immunohistochemistry (IHC), immediately immerse the eyes in 4% paraformaldehyde solution after 0.3 mL of the fixative solution is injected into the posterior segment through 3 slits made through pars plana with a 11 Parker blade (~3-4 mm from the limbus).
6. Dissect the eyecups after 3.5 h of fixation at 4 °C. Remove residual vitreous making sure to preserve the retina and organoids intact. Place back in fixative for an additional 30 min at 4 °C then rinse the eyecups 3 times each for 10 min in PBS. Transfer the eyecup to 15% sucrose for 2 h, then to 30% sucrose for a further 2 h.
7. Rinse the eyecup twice with PBS and embed in OCT medium (optimal cutting temperature compound) and flash freeze then store at -80 °C until sectioning for histology and immunochemistry.
8. Select antibodies for IHC based on the study protocol and aims.

Wyniki

This procedure enables the successful and reproducible implantation of hPSC-derived retinal organoids in the subretinal space of a large eye animal model (demonstrated here using 2 examples: wild-type cats with healthy photoreceptors (PRs) and Crx^Rdy/+ cats with degenerating PRs and retina). Using the steps indicated in Figure 1 prepare and load the hPSC-derived retinal organoids into the borosilicate glass cannula of the injection device so that the organoids ar...

Dyskusje

Implantation of hPSC-derived retinal tissue (retinal organoids) into the subretinal space is a promising experimental approach for restoring vision for late-stage retinal degenerative diseases caused by PR cell death (profound or terminal blindness). The presented approach builds on an earlier developed and successfully tested experimental therapy based on subretinal grafting of a piece of human fetal retinal tissue^23,24,25. It ...

Materiały

Name	Company	Catalog Number	Comments
0.22 µm pore syringe filter with PES membrane	Cameo	NA	can be found through various suppliers
23G subretinal injector with extendable 41 G cannula	DORC	1270.EXT
250 µL hamilton gas tight luer lock syringe	Hamilton	NA	can be found through various suppliers
6-0 Silk suture	Ethicon	707G
6-0/7-0 polyglactin suture	Ethicon	J570G
Acepromazine maleate 500mg/5mL (Aceproject)	Henry Schein Animal Health	NA	can be found through various suppliers
Buprenorphine 0.3 mg/mL	Par Pharmaceutical	NA	can be found through various suppliers
cSLO + SD-OCT	Heidelberg Engineering	Spectralis HRA+ OCT
Cyclosporine	Novartis	NA	can be found through various suppliers
Dexamethasone 2mg/mL (Azium)	Vetone	NA	can be found through various suppliers
Doxycyline 25mg/5mL	Cipla	NA	can be found through various suppliers
Fatal Plus solution (pentobarbital solution)	Vortech	NA	can be found through various suppliers
Gentamicin 20mg/2mL	Hospira	NA	can be found through various suppliers
Glass capillary (Thin-Wall Single-Barrel Standard Borosilicate (Schott Duran) Glass Tubing	World Precision Instruments	TW150-4
Methylprednisolone actetate 40 mg/mL	Pfizer	NA	can be found through various suppliers
Microscope	Zeiss	NA
OCT medium (Tissue-Tek O.C.T. Compound)	Sakura	4583
Olympic Vac-Pac Size 23	Natus	NA	can be found through various suppliers
Paraformaldehyde 16% solution	EMS	15719
Phenylephrine Hydrochloride 10% Ophthalmic Solution	Akorn	NA	can be found through various suppliers
Prednisolone 15mg/5mL	Akorn	NA	can be found through various suppliers
Propofol 500mg/50mL (10 mg/mL) (PropoFlo28)	Zoetis	NA	can be found through various suppliers
RetCam II video fundus camera	Clarity Medical Systems	NA	can be found through various suppliers
Triamcinolone 400mg/10 mL (Kenalog-40)	Bristol -Myers Squibb Company	NA	can be found through various suppliers
Tropicamide 1% ophthalmic solution	Akorn	NA	can be found through various suppliers
Vitrectomy 23G port	Alcon	Accurus systems
Vitrectomy machine	Alcon	Accurus systems
Vitreo-retinal vertical 80° scissors with squeeze handle	Frimen	FT170206T