Optimized Minimally Invasive Transscleral Subretinal Injection Technique in Mouse

Summary

Subretinal injections are an indispensable technique for preclinical development of therapeutics for photoreceptor and RPE diseases. We describe an optimized transscleral subretinal injection technique in mice that improves intraoperative success rates and minimizes retinal damage.

Abstract

The conventional method of material delivery to the subretinal space in the mouse involves dual perforation of the neural retina, which causes extensive surgical damage. This leads to variability in the subsequent outcome measures of the visual function, such as electroretinogram (ERG) recordings or behavioral vision assays, which confound efficacy assessments of experimental therapeutics. To overcome these barriers, we optimized a transscleral minimally invasive subretinal injection technique in mice. In this technique, the superior fornix is accessed using a custom-made tungsten wire eyelid speculum to perform conjunctival peritomy and tenotomy. A pinpoint sclerotomy is made using a diamond knife without penetrating the retina, through which a fine glass needle is carefully inserted at a shallow angle. The payload is delivered to the subretinal space using a microinjection pump. Optical coherence tomography (OCT) is used to assess the size and position of the subretinal bleb immediately following injection. We compared the outcomes of this technique to the conventional transretinal method performed using a spring-loaded syringe with a 33G needle sclerotomy. ERG recordings indicated excellent preservation of retinal function in transsclerally-injected mice, comparable to that of uninjected control mice. In contrast, an appreciable ERG signal reduction was observed in the conventionally injected cohorts. In summary, we have optimized a minimally invasive technique for subretinal injection in mice. We demonstrate that this technique is a robust and efficient method for gene therapy administration that minimizes anatomic damage to the retina and has minimal impact on retinal function. This method lowers the threshold for the development of therapies targeting both the retinal pigmented epithelium and photoreceptors to treat a wide range of retinal and macular diseases.

Introduction

Subretinal injection is an ocular surgical procedure in which material is delivered to the potential space between the retina and the retinal pigmented epithelium (RPE). Compared to other methods of drug delivery to the posterior segment, such as intravitreal or suprachoroidal injections, subretinal injections achieve the most immediate access to both the photoreceptors and the RPE. For delivery of viral gene therapy particles, subretinal injection achieves much higher rates of photoreceptor and RPE transduction, which allows for lower viral titers and reduced risk of viral toxicity¹. Additionally, the subretinal space is anatomically confined, limiting the systemic spread of viral particles and lowering the risk of immune responses². In humans, the safety profile of subretinal injections is very favorable, with retinal structure and function recovering to baseline levels within 1 month post-injection^3,4. Subretinal injections are used in a number of applications, such as stem cell transplantation and small molecule drug delivery², with their most common use being the delivery of viral vectors for gene therapy treatments of retinal diseases. The relatively low viral titers, minimal systemic exposure, and high rate and specificity of transduction associated with subretinal injections have made them an indispensable technique in the retina field. Subretinal injections are successfully used in patients for the treatment of inherited retinal disorders, including retinitis pigmentosa and Leber congenital amaurosis^5,6.

Given its importance to the field, the ability to perform subretinal injections in mice is crucial for preclinical studies investigating therapies for inherited retinal diseases. However, current methods for mouse subretinal injections have high failure rates and often cause surgical damage to the retina, which may limit the reliability and consistency of functional outcome measures. Conventionally, subretinal injections in rodents are performed transretinally, such that the needle penetrates the anterior retina, passes by the lens, and enters the subretinal space as a result of a second penetration of the retina^7,8 (Figure 1A). While relatively easy to perform, this procedure introduces marked inconsistency in experimental outcomes due to the variable extent of surgical damage. This is particularly limiting for studies utilizing behavioral and functional vision outcome measures. In addition, the crystalline lens is disproportionately large in mice compared to humans; this significantly increases the risk of damage to the lens during transretinal injection, rendering the animal unusable for functional vision assessments⁴.

An alternative option is the transscleral injection route, which avoids these issues by accessing the subretinal space using a posterior approach, where the needle penetrates the sclera, choroid, Bruch's membrane, and the RPE^4,9 (Figure 1B). This approach minimizes retinal damage by completely avoiding retinal penetration. It is also associated with reduced reflux of injected material from the subretinal space¹⁰, which allows for more consistent results and smaller experimental cohorts. Although the technique for transscleral subretinal injection in mice has been previously described in the literature, its widespread adoption remains limited by its technical difficulty, which can impede the success rate⁴. Here, we describe several improvements to the transscleral injection procedure, which make it easier and faster to perform, reduce the experimental variability, and improve injection success rates. These improvements include the development of a mouse eyelid speculum, preoperative administration of atropine, creation of a pinpoint sclerotomy using a diamond knife, and optimization of needle size and injection approach. This technique achieves excellent viral transduction coverage of photoreceptors and the RPE, without causing any surgical impact detectable by ERG.

Protocol

All experiments were performed in accordance with the Duke University Institutional Animal Care and Use Committee and the Association for Research in Vision and Ophthalmology Statement for the Use of Animals in Ophthalmic and Vision Research.

1. Animal preparation

1. Administer an intraperitoneal injection of 1 mg/kg of atropine (in a total volume of 50 μL) 30 min prior to performing the subretinal injection. This step greatly increases success rates by preventing potentially lethal oculocardiac and oculorespiratory reflexes triggered by transient intraocular pressure elevation during the injection^11,12.
   NOTE: Our atropine dosage is optimized for C57BL/6 mice and may need to be adjusted for other mouse strains. We have not observed any appreciable sex or age differences in atropine dosing.
2. During the entire procedure, keep the mouse on a warm surface to maximize animal comfort and reduce cold cataract development.
3. Anesthetize mouse using 2% inhaled isoflurane in oxygen (2 L/min) until recumbency is observed and animal is no longer responsive to toe-pinch. Apply a drop of 0.5% proparacaine to the eye for topical anesthesia.
4. Apply a drop of the mixture of 0.2% cyclopentolate and 1% phenylephrine to the eye to dilate the pupil for postoperative imaging of the subretinal bleb by OCT. Phenylephrine also serves to reduce intraoperative bleeding by constricting superficial blood vessels¹³.

2. Sclerotomy

1. Prior to starting the procedure, load a glass microneedle with the desired volume of solution to be injected. Attach the needle to the microinjector, with compensation pressure set at 5 hectopascals (hPa) to counteract capillary action. We typically use 1 μL volumes for subretinal injections.
2. Place the mouse under a dissecting stereomicroscope. Use 20x magnification for steps 2.3-2.6 and 50x magnification for steps 2.7-3.5.
3. For young mice, perform lateral canthotomy prior to the procedure, in order to provide ample access and visualization of the surgical site.
4. Expose the superior fornix with the help of a custom-made miniature wire-speculum (Figure 1C). Immobilize the speculum to allow hands-free upper eyelid retraction.
5. Using angled Vannas scissors, perform a paralimbal incision (~1 mm posterior to the limbus) in the superior conjunctiva and Tenon's capsule to expose the superior 3-4 clock hours of the sclera. Take care to avoid puncturing the retro-orbital venous sinus. Remove any remnants of Tenon's capsule overlying the intended injection site carefully but thoroughly.
6. Grasp the anterior edge of the conjunctival peritomy and tenotomy with locking toothed forceps and inferoduct the globe gently to expose the superior sclera. Immobilize the locked forceps to allow hands-free inferoduction.
7. Apply a drop of balanced buffered saline (BSS) immediately to the eye to prevent the bare sclera from drying. We also find the optical magnification produced by the BSS droplet to be helpful.
8. Using a diamond knife, make a pinpoint sclerotomy at ~12 o'clock and ~1-2 mm from the limbus. Position the blade in a tangential plane with respect to the sclera to ensure the sclerotomy is as small and shallow as possible.
   NOTE: This is done for two reasons: (1) a small sclerotomy allows for a tight fit of the glass needle, thereby reducing reflux during the injection step, and (2) a shallow sclerotomy is critical to avoid penetrating the retina. Note that a small amount of choroidal blood may appear at the sclerotomy site, which does not connote any negative implications for the outcome of the procedure.

3. Transscleral subretinal injection

1. In preparation for injection, use a surgical sponge to gently remove all BSS from the eye surface.
2. Hold the needle bevel-down at a shallow angle to the sclera and introduce the tip slowly and carefully into the sclerotomy to access the subretinal space. In order to avoid penetrating the retina, do this at a relatively shallow angle and just deep enough to clear the sclerotomy opening by 0.5-1 mm.
   NOTE: The bevel of the needle is positioned downward in order to minimize the possibility of choroidal delivery and to reduce the probability of puncturing the retina with the advancing needle tip.
3. While holding the needle in a steady position, initiate injection at 500 hPa using a foot-pedal and maintain uninterrupted pressure for 15 s.
4. To finish the injection, maintain injection pressure until the needle is completely removed from the sclerotomy. If the pressure is prematurely discontinued while the needle is still in the subretinal space, a large portion of the injected payload can quickly exit the eye back into the needle by capillary action.
5. Once the needle is withdrawn, notice a partial reflux of the injected solution through the sclerotomy site. This is expected, given that 1 μL volume exceeds the capacity of the subretinal space even in adult mice. Absence of reflux may indicate a failed injection due to inadvertent choroidal or intravitreal delivery.
6. Remove the locking forceps and the eyelid speculum from the eye. Take care to avoid applying pressure to the globe, as it may cause further reflux of the injected solution.
7. Immediately following the injection, apply ample lubricant eye ointment to the eye to allow for visualization of the subretinal bleb by OCT¹⁴.
8. Apply topical erythromycin ointment to the eye and provide general post-operative care to the animal.

Results

A successful injection using this transscleral technique generates a subretinal bleb observable on OCT immediately following the injection procedure (Figure 2A). Given the technical challenge of this technique, the imaging step is indispensable for identifying successfully injected animals. For researchers who plan to use this technique for gene therapy experiments, we recommend performing practice injections with adeno-associated virus (AAV) particles carrying a fluorescent reporter gene. A...

Discussion

The technique described here introduces several improvements to the previously described transscleral injection approach⁴, which enhance the reliability of the procedure and further reduce surgical damage to the eye. One such improvement is the usage of a speculum to retract the upper eyelid, allowing for stable, unencumbered, and hands-free access to the eye. Commercial specula are often poorly optimized, even for adult mice, and outright unavailable for juveniles. To overcome this obstacle, we d...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.9% Sodium Chloride Injection, USP	Hospira	PAA130066
1% Atropine Sulfate Ophthalmic Solution, USP	Akorn	NDC 17478-215-05
30° diamond knife	Katena	K2-6560
AmScope 6W LED dual gooseneck illuminator	AmScope	LED-11CR
Angled Vannas scissors	Roboz	RS-5618
BSS Sterile Irrigating Solution	Alcon	289010-0902
Cyclomydril dilating eyedrops	Alcon	0065-0359-02	0.2% cyclopentolate, 1% phenylephrine
epT.I.P.S. MicroloaderTM pipette tips	Eppendorf	5242 956.003
FemtoJet 4i Microinjector	Eppendorf	E5252000021	Discontinued product
GenTeal Tears lubricant eye ointment	Alcon	16027802
Glass microinjection needles	Clunbury Scientific	B100-58-50	30° bevel. Diameter: 50 µm outer, 29 µm inner
Isoflurane solution	Covetrus	29405
MICRON IV retinal imaging microscope with image-guided OCT	Phoenix	N/A
Stereomicroscope SteREO Discovery.V8	Zeiss	495015-0001-000
Stern-Castroviejo locking toothed forceps	Katena	K5-2552
Toothed forceps	Roboz	RS-5172
Tungsten cleaning wires for 23s gauge needles	Hamilton	18306
Ultracell PVA Eye Spears	BVI Medical	40400-8
Ultra-Fine2015 insulin syringes	BD	328438	3/10 mL, 8 mm (5/16”), 31G