An Alternative and Validated Injection Method for Accessing the Subretinal Space via a Transcleral Posterior Approach

Summary

Subretinal injections are the most common technique for delivering large therapeutic agents such as proteins and viral vectors to photoreceptors and the retinal pigment epithelium. An alternative method in mice that successfully targets the subretinal space with minimal collateral damage and fast recovery times is described here.

Abstract

Subretinal injections have been successfully used in both humans and rodents to deliver therapeutic interventions of proteins, viral agents, and cells to the interphotoreceptor/subretinal compartment that has direct exposure to photoreceptors and the retinal pigment epithelium (RPE). Subretinal injections of plasminogen as well as recent preclinical and clinical trials have demonstrated safety and/or efficacy of delivering viral vectors and stem cells to individuals with advanced retinal disease. Mouse models of retinal disease, particularly hereditary retinal dystrophies, are essential for testing these therapies. The most common injection procedure in rodents is to use small transcorneal or transcleral incisions with an anterior approach to the retina. With this approach, the injection needle penetrates the neurosensory retina disrupting the underlying RPE and on insertion can easily nick the lens, causing lens opacification and impairment of noninvasive imaging. Accessing the subretinal space via a transcleral, posterior approach avoids these problems: the needle crosses the sclera approximately 0.5 mm from the optic nerve, without retinal penetration and avoids disrupting the vitreous. Collateral damage is limited to that associated with the focal sclerotomy and the effects of a transient, serous retinal detachment. The simplicity of the method minimizes ocular injury, ensures rapid retinal reattachment and recovery, and has a low failure rate. The minimal damage to the retina and RPE allows for clear assessment of the efficacy and direct effects of the therapeutic agents themselves. This manuscript describes a novel subretinal injection technique that can be used to target viral vectors, pharmacological agents, stem cells or induced pluripotent stem (iPS) cells to the subretinal space in mice with high efficacy, minimal damage, and fast recovery.

Introduction

Subretinal injections are the primary means of delivering cellular and viral agents to the retinas of mice to study their effects on photoreceptors and the underlying RPE^1,2. Most subretinal injection protocols in mice use a transcorneal or a transcleral injection site anterior to the equator (Figure 1). This approach can result in inherent collateral damage that includes nicking and resultant clouding of the lens, disruption of the integrity of the vitreous, penetration of the neurosensory retina and iris, retinal hemorrhage, substantial retinal detachments and lasting subretinal edema ^3-9. Experimental manipulations must overcome these effects in order to evaluate the effects of therapeutic interventions^3,7,10,11. This study provides a detailed description and validation of a posterior transcleral injection method that avoids these complications, minimizes trauma and has a high success rate of targeting the subretinal space.

Injections targeting the subretinal space in mice are often very difficult to perform and most investigators encounter a high frequency of failed attempts in which the vector is delivered to an incorrect location or there is significant retinal damage, for example in a complete retinal detachment⁶. The number of eyes excluded from analysis because of injection complications is typically not reported in mouse studies, but in our own experience and in discussion with other investigators, the number of failed injections can be as high as 50% and vary dependent on the experience and capabilities of the investigator who is performing the injections. The success of the injection is typically assessed by direct fundus imaging and/or optical coherence tomography (OCT)^7,9. An easily mastered method with high success rates for subretinal injections in mice can hasten experimentation and reduce the cost of preclinical studies of treatments for retinal diseases that are major causes of blindness in the United States.

The posterior, transcleral subretinal injection technique described here is an adaptation from clinical and preclinical protocols^9,12. The noninvasive diagnostic assessments performed in injected mice demonstrate mild and highly localized damage and lack additional collateral lens, retinal and RPE injury. Furthermore, with relatively little practice, an experimenter can achieve these results with a high success rate (80 - 90% or better), thereby reducing the costs associated with such studies. This procedure can be used to deliver cellular, viral, or pharmacological therapeutic interventions to photoreceptors and/or RPE in preclinical studies and to easily evaluate experimental interventions.

Protocol

Animals: Wild type C57Bl/6J mice bred at the University of California at Los Angeles (UCLA). All animals were between 11 - 17 weeks old, and included male and female mice. All mice were group-housed, maintained in a 12:12 light/dark cycle with food and water ad libitum. All experiments were performed in accordance with the institutional guidelines of UCLA and the Association for Research in Vision and Ophthalmology Statement for the Use of Animals in Ophthalmic and Vision Research.

NOTE: All drugs and injectable agents are United States Pharmacopeia (USP) grade.

1. Surgical Preparation

1. Anesthetize the mouse with an intraperitoneal injection of 100 mg/kg ketamine and 8 mg/kg xylazine in a saline mix. Administer anesthesia to a depth such that the mouse has no toe pinch or corneal touch reflexes.
2. Maintain body temperature at 37.0 °C with a circulating water pad.
3. Dilate pupils with 2.5% phenylephrine eye drops and trim whiskers to facilitate visualization. Whiskers provide significant sensory input to the mouse, therefore, whisker trimming should remove only the portion that blocks clear access to the eye, and not to the base of the whisker. In our experience, mice show normal recovery after this procedure. Apply methylcellulose eye drops to prevent dryness and minimize anesthetic induced transient cataracts¹³.
4. Sterilize instruments prior to surgery (i.e., betadine and ethanol or hot beads).
5. Prepare diluted fluorescein (0.01% using 0.9% saline) in a sterile environment (i.e., biosafety cabinet) if visualization will be performed (see section 3 below).

2. Injection Site Preparation

1. Prepare a syringe (e.g., 5 µl syringe) with the appropriate injection volume (e.g., 0.3 to 1.0 µl).
2. Position the mouse so the eye is facing up and clearly visible in the dissecting microscope.
3. Gently pinch the temporal conjunctiva with fine tipped forceps. Make a circumferential incision of approximately 90 degrees using curved Vannas scissors.
4. Repeat step 2.3 with the underlying Tenon's capsule.
5. Resect the surrounding connective tissue with fine tipped forceps while rotating the globe nasally. Work towards the injection site approximately 0.5 mm temporal to the optic nerve. Use great care to avoid disrupting the retro-orbital sinus.

3. Sclerotomy and Subretinal Injection

NOTE: It is recommended that the injection of 0.01% fluorescein in 0.9% Saline be used to assist with visualization while learning this procedure. The topographic distribution of fluorescein can be effectively documented with fundus imaging (see section 4 below).

1. Make a small scleral incision at the injection site by gently scratching the eyecup with a 22.5-degree ophthalmic blade. This incision should only be large enough to allow the tip of the needle to pass through the sclera.
2. Insert the beveled 33 G needle (angled 5 - 10° into the sclerotomy with the bevel facing and angled parallel to the retina. Inject desired volume (e.g., 0.3 to 1.0 µl of 0.01% Fluorescein for learning purposes).
   NOTE: Maintain sterility of the syringe by thoroughly cleaning with successive washes of a suitable solvent and DI water before each injection.
3. Depress the plunger slowly (over ~ 3 sec) without moving the needle and with even pressure.
   NOTE: When the needle is in the subretinal space, a slight resistance will be felt while depressing the plunger. There will be no to minimal resistance if the needle punctures the retina, and high resistance if the needle does not penetrate the sclera or RPE.
4. Wait several seconds before withdrawing the needle to minimize backflow.
5. Rinse the eye with sterile buffered saline and ensure the eye has rotated back to its normal position.

4. Assessment of Retinal Detachment by OCT and Fundus Imaging

1. Perform OCT imaging immediately after injection to evaluate the quality of the injection and at appropriate time points post-injection as needed to evaluate retinal structure.
   NOTE: Examples of the use of OCT in similar studies has been previously described^7,14.
   1. Adjust and align the OCT image to target the site of injection. The injection site should be midline and 0.5 mm temporal to the optic nerve head. Repeat as needed if the detachment is out of frame or not optimally centered.
2. Visualize retinal detachment and dye injection area with en-face fundus imaging^7,14.
   NOTE: If an OCT imaging system is not available, injection of a small amount of fluorescein with a vector for practice will allow visualization with any fundus camera that performs fluorescein angiography using the same excitation wavelengths and blocking filters. Localized areas of hyper-fluorescence will appear underneath the vasculature and the vasculature will have sharp and distinct boundaries if the subretinal space is targeted correctly. The edge of the bleb from the injection will be demarcated by the transition from hyper- to hypo- fluorescence. Several instruments provide this capability for the mouse; the instrumentation used here is described elsewhere¹⁴.

5. Post-operative Care

1. Apply a thick coat of triple antibiotic ophthalmic cream to the corneal surface of the injected eye.
2. Place mice in clean solitary cages for recovery. Do not combine mice that have undergone surgery until they are fully recovered.
3. Monitor respiration and temperature during anesthesia recovery. Monitor animals until they can maintain sternal recumbency.
4. Perform additional appropriate post-operative monitoring and treatment, including a sub-cutaneous injection of carprofen (5 mg/kg) for post-surgical pain management.

6. Assessment of Retinal Function by Electroretinography (ERG)

1. Perform ERG analysis pre-injection and at appropriate times post-injection as needed to evaluate retinal function. If the injection was made into the subretinal space, the retinal detachment should resolve within 72 hr.
   1. Use standard ERG techniques to evaluate retinal function before and after injection as previously described^14,15.

7. 3D Reconstruction and Bleb Volume Quantification

NOTE: OCT scans with high contrast encompassing the entire detachment within the frame of view are optimal for use. ImageJ/Fiji^17,18 and Imaris were used, but other software can be used.

1. Export the b-scan of interest, import to ImageJ/Fiji and crop (Image > Crop) the portion of the OCT scan to be modeled using the rectangular selection tool.
   1. Adjust contrast (Image > Adjust > Brightness/Contrast) and delineate any missing boundaries by connecting two sections with a line.
   2. Draw a straight line with the line tool (holding shift) that spans the RPE to the photoreceptor layer. Measure (Analyze > Measure) the length of the line to obtain size of maximum detachment for step 7.8.
2. Import cropped frames to the 3D-reconstruction software (See Table of Materials) using the "RGB to Gray" plugin and MATLAB Compiler Runtime.
3. Set the voxel size (under Image Properties) using the calibration parameters from the OCT scan (x,y,z).
4. Execute the "RGB to Gray" plugin (under Image Properties), with equal weighting to each channel, to create a fourth channel. Delete original Red-Green-Blue channels.
5. Invert the gray channel using contrast change. Store Image.
6. Click the "add a new surface" button in the 3D-View tab, and begin the guided 4 step process of creating the surface.
   1. Set the surface level detail (step 1 of 4).
      NOTE: In our experience 8.0 to 12.0 was the most effective range.
   2. Set the maximum sphere size (under Background Selection) to slightly less than the maximum detachment size measured in 7.1.2. Create the surface and undo gray channel inversion (step 2 of 4).
   3. Set the threshold to the maximum value so the surface of the negative spaces outside of the retina and the detachment do not come in contact (step 3 of 4).
   4. Set the filter type to the number of voxels and isolate the negative space in the detachment site by size. Finish the surface (step 4 of 4).
      NOTE: The volume of the detachment surface is located under volume in the statistics tab.

Results

Posterior approach transcleral subretinal injections were performed on 31 healthy eyes from 16 wild type mice with injections of 0.3 µl (n = 18), 0.5 µl (n = 8) and 1.0 µl (n = 5) of 0.01% fluorescein. One eye was excluded from injection due to a pre-existing corneal opacity that prevented structural and functional analysis. Every injected eye is included in this report. No unintended retinal detachments, punctures of the neurosensory retina, or leakage into the vitreous were detected nor...

Discussion

Subretinal injections are the method of choice for the delivery of viral vectors and stem cell-derived therapy for manipulating photoreceptors and the RPE in both basic research and clinical treatment. In patients, subretinal injections are typically done with an anterior sclerotomy at the pars plana, a posterior core vitrectomy and penetration of the retina by the needle with direct visualization. As with most vitrectomy procedures, it is common for cataract formation to occur prematurely unless the eye is already pseud...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Hamilton Model 62 RN SYR	Hamilton	87942	Syringe x 1
Hamilton Needle 33 G, 1.0", 20 DEG, point 3 (304 stainless steel)	Hamilton	7803-05	Needles x 6
Vannas Curved Scissors	Ted Pella, INC.	1347	5 mm Blade
22.5 Degree Microsurgery Knife	Wilson Ophthalmic Corp.	91204
Ketaject	Phoenix	NDC 57319-609-02	Ketamine
Anased	Lloyd Laboratories	NDC 61311-482-10	Xylazine
Fluorescein 10% AK-Fluor	Akorn	NDC 17478-253-10	100 mg/ml
0.9% Saline USP	Hospira	NDC 0409-4888-50	0.9% NaCl
Antibiotic Ointment	Akorn	NDC 17478-235-35	Ophthalmic
Water Circulating Pump	Gaymar	TP-500 T/Pump	P/N 07999-000
sd-OCT	Bioptigen	R-Series	Commercial
Fundus Camera	Phoenix Research Laboratories	MICRON III
Tweezers Type 3	Ted Pella, INC.	5385-3SU
2.5% Phenylephrine	Paragon BioTeck	NDC 42702-102-15	Ophthalmic
IMARIS8	Bitplane		Version 8.1.2
ImageJ	NIH		V1.8.0_77
Hypromellose 2.5%	Goniovisc	AX0401	Methylcellulose
Eye Drops (Rinse)	Bausch & Lomb		Saline Solution
Microscope	Zeiss	Stemi 2000	Microscope
Light source	Fostec	P/N 20520	Light source