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. 2024 Jul 15;31(1):2379369. doi: 10.1080/10717544.2024.2379369

Development of a novel SupraChoroidal-to-Optic-NervE (SCONE) drug delivery system

Bryce Chiang a,✉, Kathleen Heng a,b, Kyeongwoo Jang a, Roopa Dalal a, Yaping Joyce Liao a,c, David Myung a,d, Jeffrey L Goldberg a
PMCID: PMC467098  PMID: 39010743

Abstract

Purpose

Targeted drug delivery to the optic nerve head may be useful in the preclinical study and later clinical management of optic neuropathies, however, there are no FDA-approved drug delivery systems to achieve this. The purpose of this work was to develop an optic nerve head drug delivery technique.

Methods

Different strategies to approach the optic nerve head were investigated, including standard intravitreal and retroorbital injections. A novel SupraChoroidal-to-Optic-NervE (SCONE) delivery was optimized by creating a sclerotomy and introducing a catheter into the suprachoroidal space. Under direct visualization, the catheter was guided to the optic nerve head. India ink was injected. The suprachoroidal approach was performed in New Zealand White rabbit eyes in vivo (25 animals total). Parameters, including microneedle size and design, catheter design, and catheter tip angle, were optimized ex vivo and in vivo.

Results

Out of the candidate optic nerve head approaches, intravitreal, retroorbital, and suprachoroidal approaches were able to localize India ink to within 2 mm of the optic nerve. The suprachoroidal approach was further investigated, and after optimization, was able to deposit India ink directly within the optic nerve head in up to 80% of attempts. In eyes with successful SCONE delivery, latency and amplitude of visual evoked potentials was not different than the naïve untreated eye.

Conclusions

SCONE delivery can be used for targeted drug delivery to the optic nerve head of rabbits without measurable toxicity measured anatomically or functionally. Successful development of this system may yield novel opportunities to study optic nerve head-specific drug delivery in animal models, and paradigm-shifting management strategies for treating optic neuropathies.

Translational Relevance

Here we demonstrate data on a new method for targeted delivery to the optic nerve head, addressing a significant unmet need in therapeutics for optic neuropathies.

Keywords: ophthalmic drug delivery, ocular drug delivery, optic nerve head, optic neuropathy, glaucoma, suprachoroidal space

Introduction

Optic neuropathies, such as glaucomatous optic neuropathy, ischemic optic neuropathies, autoimmune disease, optic disc drusen, and toxic/metabolic optic neuropathies, are vision-threatening disorders affecting millions of patients. Though some optic neuropathies can have good visual prognosis in the near term, most cause permanent and sometimes even catastrophic vision loss. This is because as retinal ganglion cell (RGC) axons are lost, they do not spontaneously regenerate, and at some later point, the RGCs die and are not replaced (Williams et al., 2020). Thus, new clinical strategies to slow disease progression by promoting RGC survival and/or axon growth are desperately needed.

The optic nerve head (ONH) has been implicated as the site of damage and/or most aggressive change in many optic neuropathies. The optic nerve head is the site where the RGC axons coalesce to form the optic nerve. The optic nerve head is one of the most metabolically active tissues due to the highest concentration of unmyelinated axons in the body (Muench et al., 2021), which increases susceptibility to toxic/metabolic optic neuropathies. Furthermore, it is the site where the majority of vasculature enters and leaves the eye (Hayreh, 2001; Cherecheanu et al., 2013). The optic nerve head is a watershed vascular zone and is at risk for ischemic events, as in ischemic optic neuropathies (Hayreh, 2001). The lamina cribrosa, which supports the optic nerve head, is one of the mechanical weak points of the globe (Burgoyne, 2011; Downs & Girkin, 2017). There are structural changes of the optic nerve head and lamina cribrosa tied to glaucoma pathogenesis (Quigley et al., 1983; Burgoyne, 2011). Thus, treating cells and tissue within the optic nerve head may be useful in the management of optic neuropathies, however to our knowledge, no research or FDA-approved method to achieve targeted optic nerve head drug delivery exists.

Targeted drug delivery should be designed to achieve therapeutic doses at the optic nerve head and low concentrations elsewhere. Current ophthalmic delivery methods may include topical eye drops, intravitreal injections (using 30- or 32- gauge hypodermic needles), sub-Tenon’s delivery, and systemic administration – none of which specifically target the optic nerve head. Agents to be delivered could include neuroprotective agents, such as neurotrophic factors (including ciliary neurotrophic factor (Goldberg et al., 2023; Ghasemi et al., 2018; Yungher et al., 2017; Pease et al., 2009; Cen et al., 2007) and brain derived neurotrophic factor (Harper et al., 2011; Heng et al., 2024)), steroids (corticosteroids (Stunkel & Van Stavern, 2018; Saxena et al., 2014; Beck et al., 1992) and neurosteroids (Ishikawa et al., 2014; 2018; Izumi et al., 2023)), antioxidants (Martucci & Nucci, 2019; Ekicier Acar et al., 2020; Kang et al., 2021; Cáceres-Vélez et al., 2022), vasoactive drugs (Quigley et al., 2015; Mursch-Edlmayr et al., 2021; Arora et al., 2022; Eghbali et al., 2023), viruses for gene therapies (Martin & Quigley, 2004; Leaver et al., 2006; Cen et al., 2017; Yungher et al., 2017), or even cell-based therapies (Harper et al., 2011; Mead & Tomarev, 2020; Coco-Martin et al., 2021). Drugs and even gene therapies could be delivered through the smallest gauge cannulas (as small as 41 G (Irigoyen et al., 2022)) although cell therapies would require larger gauge to allow for internal diameters greater than cell diameters (Amer et al., 2015). In all cases, however, there is motivation to limit exposure of high concentrations of certain therapeutics to the retina or other ocular structures. Thus, there is a significant clinical need for targeted delivery of therapeutics directly to the optic nerve head rather than relying on diffusion from the vitreous or orbit to achieve adequate dosing. Here, we demonstrate and determine the feasibility of a new suprachoroidal approach to optic nerve head drug delivery.

Materials and methods

All materials were purchased from Sigma-Aldrich unless otherwise specified.

Localization of injection techniques to optic nerve head ex vivo

Microneedles were ground to length using previously described methods (Chiang et al., 2016). Briefly, a rotary tool was used to grind a 45° angled bevel into the needle (27 and 30 G hypodermic needles, BD, Franklin Lanes, NJ; 32 G hypodermic needles, TSK Steriject, TSK Labs, Japan). The bevel was polished with sandpaper (220 and then 400 grit, 3 M, St. Paul, MN), and the bore was cleared with a 30 G needle flushing with water and isopropyl alcohol. The needle was measured to ensure correct length, and visually inspected under microscope.

Enucleated albino New Zealand White rabbit eyes were purchased fresh (Pel Freez, Rogers, AR) and overnight shipped in DMEM solution with antibiotic and antifungal agents. Conjunctiva, Tenon’s capsule, extraocular muscles, orbital fat, and fascia were carefully dissected off the globe. An intracameral injection of 0.1–1.5 mL DMEM solution was used to raise the intraocular pressure to physiologic levels (10–20 mmHg). India ink (Blak, Allegory Ink, Orlando, FL) was diluted 1:4 in DMEM, and loaded in a 100 µL gas-tight syringe (Hamilton Company, Reno, NV) prior to injection. The injection techniques were performed as described below (Figure 1). Replicates of 6–10 were performed per injection technique, and in each case 10 µL India ink was injected unless otherwise specified.

Figure 1.

Figure 1.

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Schematic diagram of region surrounding optic nerve head highlighting trajectories of injection techniques A-G. ON = optic nerve stalk, VH = vitreous humor, re = retina, Ch = choroid, Sc = sclera, LC = lamina cribrosa, on = optic nerve.

For retroorbital optic nerve injection [Figure 1(A)], a 32-gauge ½-in needle (TSK Labs, Japan) was positioned perpendicular to the optic nerve 2 mm posterior to the globe. The needle was carefully inserted into the optic nerve, taking care not to penetrate full thickness.

For retroorbital optic nerve injection with microneedle [Figure 1(B)], a 30-gauge 750-µm microneedle was positioned perpendicular to the optic nerve 2 mm posterior to the globe. The needle was inserted until the needle hub came in contact with the nerve.

For retroorbital optic nerve injection angled toward the optic nerve head [Figure 1(C)], a 32-gauge ½-in needle was angled 45° at the junction of the optic nerve and globe. The needle was carefully inserted into the globe, taking care not to penetrate full thickness through the tissue.

For intravitreal injection [Figure 1(D)], a 32-gauge ½-in needle was positioned 2 mm posterior to the limbus at pars plana. The needle was inserted through the sclera, taking care to avoid the lens.

For intravitreal approach to optic nerve head [Figure 1(E)], a 32-gauge 1-in needle was positioned 2 mm posterior to the limbus at pars plana. The needle was inserted through the sclera, and advanced toward the optic nerve head until resistance was felt, then the needle was advanced taking care not to penetrate full thickness.

For suprachoroidal injection [Figure 1(F)], a 30-gauge 750-µm microneedle was positioned 2 mm posterior to the limbus at pars plana. The needle was inserted until the needle hub came in contact with sclera.

For suprachoroidal tunneling to optic nerve head [Figure 1(G)], a 27-G 750-µm microneedle positioned 2 mm posterior to the limbus was used to inject 50 µL viscoelastic (Provisc, Alcon, Geneva, Switzerland) into the suprachoroidal space to reduce the risk of inadvertent choroidal/retinal tear. A 32-G 1-in needle with 45° bevel (Hamilton Company, Reno, NV) was fed into the injection site tangentially and the needle was tunneled through the SCS toward the optic nerve head. Once resistance was felt, an injection of 5 µL India ink was performed. Hereafter, we refer to this technique as SupraChoroidal-to-Optic-NervE (SCONE) delivery (Figure 2).

Figure 2.

Figure 2.

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Diagram of SCONE delivery technique: (A) diagram of eye prior to procedure. Sc-sclera, Ch-choroid, re-retina, VH-vitreous humor, on-optic nerve, ONH-optic nerve head. (B) Microneedle injection of viscoelastic into suprachoroidal space (SCS). (C) Catheter introduced into SCS directed toward ONH. Can either perform: (Di) injection into peripapillary SCS, or (Dii) needle exposed and penetrated into ONH, then injection into ONH.

After injection, all eyes were snap frozen in isopropyl alcohol chilled over dry ice. A razor blade was used to make 2 sagittal sections on either side of the optic nerve while the globe was still frozen. The resulting section was photographed. Image analysis was performed offline. For photographs from each injection technique, the proportion of eyes with India ink present within a 1-mm radius circle centered on the optic nerve head was counted.

Optimization of injection parameters ex vivo

In performing SCONE delivery, further experiments were used to optimize the insertion of the catheter through the sclerotomy into the suprachoroidal space. Four parameters to facilitate insertion were explored – (A) microneedle to make the initial sclerotomy, either 27 G or 30 G; (B) performing a conjunctival peritomy or not; (C) twirling/rotating the microneedle 360° clockwise and counterclockwise within the sclerotomy or not, and (D) microneedle injection of viscoelastic within the suprachoroidal space or not. Replicates of 8–13 were performed for each experimental condition in ex vivo rabbit eyes as above. The ability to easily insert the catheter into the suprachoroidal space within 1–2 tries was tabulated.

Ethical approval statement and animal care

All procedures were reviewed and approved by the Institutional Animal Care and Use Committee at Stanford University (IACUC #33781) and performed in accordance with ARRIVE guidelines and the Association for Research in Vision and Ophthalmology Statement for the Use of Animals in Ophthalmic and Vision Research. Animals were needed to test the delivery technique in vivo to assess safety and efficacy for eventual translation human studies. A total of 25 albino New Zealand White rabbits aged 3–12 mo., of either sex, and weighing 600–1500 g were purchased from Charles River Laboratories (Wilmington, MA) or West Oregon Rabbit Company (Philomath, OR) for this study. They were singly housed on a 12:12 light cycle in an AAALAC-accredited facility, and allowed to acclimatize for at least 3 days. Rabbits were fed a standard pellet diet (Global Rabbit Diet 2030, Harlan Teklad). Food enrichment (Timothy hay or hay balls) was provided daily, and enrichment devices (hanging mirrors or rattles) were exchanged weekly. All experiments were performed at Stanford University in IACUC approved facilities from October 1, 2022 to September 1, 2023.

In vivo optic nerve head injections

Figure 2 describes the in vivo SCONE procedure. Induction of anesthesia was performed by intramuscular injection of 30 mg/kg ketamine and 5 mg/kg xylazine and then the animals maintained on 1–8% sevoflurane. The procedure was performed under direct visualization with a surgical microscope and/or RetCam fundus camera (Natus Medical, Middleton, WI), and photographs were taken during the procedure. Prior to the procedure and after, radial optical coherence tomography (OCT) images centered on the optic nerve head were acquired. Only the right eyes were used to minimize surgical variability, and the left eye was left untouched. A single surgeon (BC) performed the injections. The eye was prepped with 5% betadine solution (Alcon, Fort Worth, TX) in the usual ophthalmic fashion. Proparacaine, phenylephrine, and tropicamide drops were given. A small conjunctival peritomy was performed in the supratemporal quadrant. A 27-G 750-µm microneedle positioned 2 mm posterior to the limbus was used to inject 50 µL viscoelastic into the suprachoroidal space to reduce the risk of inadvertent choroidal/retinal tear. A 32-G stainless steel catheter with 25° or 45° bevel (Hamilton Company, Reno, NV) connected to a 25 µL gas-tight syringe (Hamilton Company, Reno, NV) was fed into the sclerotomy site tangentially and the needle was tunneled through the SCS under direct visualization toward the optic nerve head. In some experimental groups, the catheter was covered with a polyimide sheath (Zeus Industrial Products, Orangeburg, SC) that was uncovered once near the optic nerve head. In all animals, 5 µL of India ink suspension was injected. After the procedure, fundus and OCT images were taken again. After the procedure, animals were given buprenorphine 0.15 mg/kg and again in 72 hr if showing signs of pain, and/or meloxicam 0.2–0.6 mg/kg every 24 hr. At the predetermined timepoints, the animal was anesthetized for reimaging with fundus photography and OCT images, as above. Animals were sacrificed by veterinary staff at the 1 wk timepoint unless otherwise specified with a large dose of 100 mg/kg phenobarbital. If animal exhibited signs of pain not controlled by pain medication, the animal would have been prematurely euthanized (note that this occurred in no animals). Eyes were enucleated and processed for histology.

It was not possible to blind the surgeon to the exact SCONE parameters. Based on ex vivo experiments above, a power calculation were performed. A sample size of N = 3 per group was needed to detect a mean difference between 85% and 33% with a standard deviation of 20% (the best and worst performing groups in the ex vivo study) with an alpha = 0.05 and beta = 0.85. Thus, N = 3 animals per group were randomized to receive SCONE delivery with catheter bevel of 25° or 45°, and/or a protective sheath. An interim analysis was performed, and the worst performing group was not studied further. An additional N = 3 per group was performed for the remaining groups. A final N = 3 per group was performed for the best performing group. A total of 25 rabbits were tested sequentially with no exclusion or inclusion criteria. There is the possibility there was a learning curve. The primary outcome was presence of India ink at the optic nerve head. Masked graders were given pre- and post- procedure fundus photographs and asked to determine if there was India ink (i.e. black staining) at the optic nerve head. All animals were included in the analysis with no removal of any data points.

Visual evoked potential recordings

In a subset of animals that had successful SCONE injection, RETeval (LKC Technologies, Gaithersburg, MD) was used to obtain visual evoked potentials 3 wks. after injection. The animal was anesthetized with 1–8% sevoflurane. Needle electrodes were inserted subcutaneously between the shoulder blades as a grounding electrode, at the bregma for the recording electrode, and between orbits along the frontal suture for the reference electrode. A light stimulus of 8.0 cd*s/m2 at 0.99 Hz was used, and recording occurred for 300 ms for each stimulus. Each waveform averaged over 100 stimuli by the RETeval. Replicates of 3–5 were done in the treated eye and the untreated eye.

Statistical analysis

Presence or absence of India ink at the optic nerve head was graded by blinded graders. Statistical analysis was conducted using Matlab (Mathworks, Natick, Massachusetts), Microsoft Excel (Microsoft Corp., Redmond, Washington), and GraphPad Prism (GraphPad Software, San Diego, California). Mean ± standard deviation is presented where appropriate. The significance of probabilities was compared on a binomial distribution against a 50% success rate. Paired Student t-test with Holm-Sidak multiple comparison correction was used to compare means. When appropriate, Chi-squared analysis was used to compare frequencies of occurrences. A statistical significance level of 0.05 was set a priori.

Results

Enucleated rabbit eyes were used to trial different injection techniques to the optic nerve head. Rabbits were chosen as the model animal due to the size of the eye globe (on the same order of magnitude as human eyes). Representative images of sagittal sections of the frozen globes after injection of India ink are shown in Figure 3. The probability of success of enucleated eyes with India ink within 1 mm of the optic nerve head are shown in Figure 4. Retroorbital optic nerve injection angled toward optic nerve head [C], intravitreal approach to optic nerve head [E], and suprachoroidal tunneling to optic nerve head [G] were able to localize India ink to the optic nerve head better than 50% of attempts (p < 0.05 for each). Retroorbital optic nerve injection [A] and retroorbital optic nerve injection with microneedle [B] were able to localize India ink to the optic nerve but not to the optic nerve head. Intravitreal injection [D] and suprachoroidal injection [F] were not able to localize to the optic nerve or optic nerve head.

Figure 3.

Figure 3.

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Representative images of enucleated albino rabbit eye snap frozen after injection with India ink. (A) Retroorbital optic nerve injection, (B) Retroorbital optic nerve injection with microneedle, (C) Retroorbital optic nerve injection angled toward optic nerve head, (D) Intravitreal injection, (E) Intravitreal approach to optic nerve head, (F) Suprachoroidal injection, (G) Suprachoroidal tunneling to optic nerve head. (H) Schematic diagram of region surrounding optic nerve head highlighting trajectories of injection techniques A-G. Circle indicates 1 mm radius centered on optic nerve head. K = cornea, L = lens, on = optic nerve stalk, VH = vitreous humor, re = retina, Ch = choroid, Sc = sclera, LC = lamina cribrosa, on = optic nerve.

Figure 4.

Figure 4.

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Proportion of successful localization to optic nerve head (ONH) for each injection technique (probability of success ± standard error of binomial distribution). (A) Retroorbital optic nerve injection, or “RO”; (B) Retroorbital optic nerve injection with microneedle, or “RO w/MN”; (C) Retroorbital optic nerve injection angled toward optic nerve head, or “RO angled”; (D) Intravitreal injection, or “IVT”; (E) Intravitreal approach to optic nerve head, or “IVT to ONH”; (F) Suprachoroidal injection, or “SCS injection”; (G) Suprachoroidal tunneling to optic nerve head, or “SCONE”. * indicates significance (p < 0.05) based on binomial distribution compared with 50%.

Due to difficulty inserting a catheter through the sclerotomy into the suprachoroidal space, further multifactorial studies were done in ex vivo rabbit eyes (Table 1) to improve this step. Using a larger needle (27 G vs 30 G), performing a conjunctival peritomy, and twirling/rotating the microneedle all aided in passing the catheter through the sclerotomy (all over 50% and in some cases approaching 100%; p < 0.05). Injection of viscoelastic into the suprachoroidal space did not aid in passing the catheter but was still done in the in vivo studies.

Table 1.

Multifactorial experiment to optimize catheter insertion through sclerotomy.

Microneedle size Peritomy Twirl SCS injection Success Fail Success (%)
27 G yes no no 10 3 76.9*
27G yes yes no 10 0 100.0*
27G yes no yes 10 2 83.3*
27G yes yes yes 9 1 90.0*
30G yes no no 4 4 50.0
30G yes yes no 9 6 60.0
30G yes no yes 5 4 55.6
30G yes yes yes 9 1 90.0*
27G no no no 7 2 77.8
27G no yes no 8 3 72.7
27G no no yes 6 6 50.0
30G no no no 5 6 45.5
30G no yes no 5 6 45.5
30G no no yes 1 7 12.5
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*indicates p < 0.05.

SCONE delivery could be used to deliver at or near the optic nerve head (Figure 5 and Figure 6). In some animals, the optic nerve head changed concavity immediately after injection, and reverted to baseline anatomy at a later time point (Figure 5E vs F). This could be due to sudden increase in optic nerve head volume and subsequent resorption of the formulation.

Figure 5.

Figure 5.

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(A–D) Fundus photography before (a), during (B), immediately after (C), and 21 days after (D) SCONE delivery of India ink to optic nerve head in rabbit 10 A. (E-G) Optical coherence tomography of optic nerve head before (E), immediately after (F), and 21 days after (G) SCONE delivery of India ink. (H) Gross image of enucleated rabbit eye 21 days after SCONE delivery of India ink. (I and J) Histology of rabbit eye after SCONE delivery. J is magnified view of region of interest in I. ONH = optic nerve head, cath = catheter, Sc = sclera, Re = retina.

Figure 6.

Figure 6.

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SCONE delivery in rabbits 10 C (A, B, C), 10 G (D, E, F), 12 A (G, H, I), 12 F (J, K, L), 14 A (M, N, O). Fundus photographs before (A, D, G, J, M) and after (B, E, H, K, N) SCONE delivery of India ink, and gross view of the posterior globe after enucleation (C, F, I, L, O).

The effect of catheter bevel angle and presence of catheter sheath were investigated in vivo. Masked graders were shown pre- and post- procedure fundus photographs and were asked if the optic nerve head had India ink (Figure 7A). Gross examination of posterior aspect of the globe and optic nerve after enucleation was also done to look for India ink at the optic nerve (Figure 7B). Since the sclera is opaque, the India ink may be visible within the globe (with fundus photography) or outside the globe (with gross exam) so the two examinations were combined (Figure 7C). The goal of delivery to the optic nerve was achieved, although some eyes also showed delivery into the peripapillary suprachoroidal space in addition to the optic nerve head.

Figure 7.

Figure 7.

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Probability (± standard error of binomial distribution) of targeted optic nerve head (ONH) of India ink using SCONE delivery when looking at fundus images (a), when looking at gross examination of globe after enucleation (B), or when looking at the combination of the two (C). * indicates p < 0.05.

Complications were also tabulated for each of the animals (Figure 8), and these included peripheral retinal tear, retinal detachment, suprachoroidal hemorrhage, vitreous hemorrhage, and optic nerve hemorrhage. Though all these complications are considered vision threatening, optic nerve hemorrhage was attributed primarily due to the act of delivering to the optic nerve (the others may be mitigated with better technique and devices, see Discussion). The success rate was found for each of the catheter designs by taking into account the localization of India ink at the optic nerve head and no optic nerve hemorrhage (Figure 9). Two conditions did similarly well – (i) catheter bevel angle of 45° and no catheter sheath, and (ii) catheter bevel angle of 25° with catheter sheath – with a success rate of 79% and 83%, respectively. The two alternatives likely did not do as well likely because the catheter sheath may impede the actual delivery marginally and the sharper catheter without sheath was more difficult to stay within the suprachoroidal space. Further studies and development may be needed to examine this further.

Figure 8.

Figure 8.

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Complication rate (probability ± standard error of binomial distribution) with SCONE. RD = retinal detachment, ON hem = optic nerve hemorrhage, Vit hem = vitreous hemorrhage, SCS hem = suprachoroidal hemorrhage.

Figure 9.

Figure 9.

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Probability of success (± standard error of binomial distribution), defined as targeted optic nerve head (ONH) delivery of India ink using SCONE delivery and no optic nerve hemorrhage. * indicates p < 0.05.

On histology, a suprachoroidal approach to the optic nerve head usually deposited India ink in the optic nerve head sheath, i.e. in the subarachnoid space (Figure 5I and J, and Figure 10). Other eyes demonstrated delivery into the peripapillary suprachoroidal space near the optic nerve, the optic nerve parenchyma, and any combination of the above (Figure 11).

Figure 10.

Figure 10.

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Histology of rabbit 14 A after SCONE delivery of India ink. The India ink is within the sheath of the optic nerve.

Figure 11.

Figure 11.

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Histology with H&E staining at 40x zoom (A, C, and E) and 100x zoom (B, D, and F) demonstrating India ink in optic nerve sheath (a and B); in the optic nerve parenchyma (C and D); and in the sheath and parenchyma (E and F). Circle indicates region of interest that is enlarged in higher magnification. K = cornea, L = lens, Re = retina, Ch = choroid, SCS = suprachoroidal space, Sc = sclera, EOM = extraocular muscles, on = optic nerve.

In the subset of eyes with successful SCONE delivery, there was little or no difference in the latency (p = 0.88) or amplitude (p = 0.10) of visual evoked potentials performed in the treated and untreated eyes 3 wks. after injection (Figure 12). This is suggestive that the visual system, in particular the optic nerve, is not compromised with this delivery technique.

Figure 12.

Figure 12.

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(A) Visual evoked potentials (VEP) traces from rabbit 10 A in the treated eye (top) and fellow untreated eye (bottom) 3 weeks after SCONE injection. (B) Mean ± SD VEP latency for all treated and fellow eyes (N = 5). (C) Mean ± SD VEP amplitude for all treated and fellow eyes (N = 5).

Discussion

The goal of this project was to assess different approaches to ocular drug delivery methods specifically targeting the optic nerve head. Despite the well documented damage occurring at the optic nerve head in various optic neuropathies, there are no treatments that attempt to treat the diseased tissue. The need for targeted optic nerve delivery stems, in part, from the combination of known or hypothesized pathology occurring at this location in many optic neuropathies, but inadequate and/or nonspecific dosing at the optic nerve head with current approaches. Here we found a novel approach through the suprachoroidal space, SCONE delivery, which demonstrates higher specificity and efficacy for optic nerve head delivery than other approaches.

Targeted delivery to the optic nerve head may spur paradigm shifting treatments for many classes of optic neuropathies. There are known or hypothesized molecular pathways that play a role in disease, and modulation of these pathways with new or repurposed therapeutics may be possible if the diseased cells can be reached. The drug delivery techniques described herein could enable some of these new therapies. For example, targeted delivery of mitochondria may be used to achieve direct mitochondrial transfer in Leber’s hereditary optic neuropathy (Martin & Quigley, 2004; Caicedo et al., 2017; Cáceres-Vélez et al., 2022; Borcherding & Brestoff, 2023), thereby preventing the catastrophic vision loss typically experienced with this disease. There are no current treatments for ischemic optic neuropathy (Stunkel & Van Stavern, 2018) and targeted delivery of neuroprotective agents (e.g. neurotrophic factors (Goldberg et al., 2023; Ghasemi et al., 2018; Yungher et al., 2017; Pease et al., 2009; Cen et al., 2007), antioxidants, and/or steroids (Stunkel & Van Stavern, 2018; Saxena et al., 2014; Beck et al., 1992)) to the optic nerve head may ameliorate axonal loss. And finally, glaucomatous damage occurs at the optic nerve head possibly through neuro-inflammatory (Evangelho et al., 2019; Participants LIIoAaGN., 2017; Soto & Howell, 2014) or biomechanical insults (Downs & Girkin, 2017;Gerberich et al., 2021). Targeted delivery to the optic nerve head may be useful as a neuroprotective strategy separate from intraocular pressure reduction by modulating neuroinflammation and/or the biomechanics at the optic nerve head and therefore prevent vision loss from glaucoma.

Intravenous, oral, and/or intravitreal routes of administration are the current state of the art in treating optic neuropathies clinically. Molecular diffusion rates are proportional to the square of the path length (Welty et al., 2014). For example, a glucose molecule delivered in the anterior segment would take 171 min to travel 3.2 mm (the axial length of a mouse eye (Park et al., 2012)), or >50x longer at 9204 min to travel 23.5 mm (the axial length of a human (Meng et al., 2011))! An intravitreal injection may shorten the diffusion distance, but the optic nerve head occupies less than 1% of the surface area that molecules may diffuse toward. This can at least partially explain why some therapeutics that show promise in small rodents are not successful in larger animals. Other reasons for difficulty in delivering to the optic nerve head include (i) its location at the posterior aspect of the globe and deep within the orbit, (ii) presence of blood retinal barrier, (iii) its delicate structure, and (iv) its small size (Kim et al., 2014).

Looking at available research methods in the literature, intravitreal injections (Dulz et al., 2020; Cui et al., 2003; Patel et al., 2017; Kurimoto et al., 2010; Monsul et al., 2004; Junnuthula et al., 2021; Yu et al., 2001), retroorbital optic nerve injections (Mesentier-Louro et al., 2019;Tehrani et al., 2018), and systemic administration (Beck et al., 1992) are the current state of the art in dosing the optic nerve head. None of these techniques are placing the bulk of the drug at the optic nerve head to achieve therapeutic levels of drug at the optic nerve head and low concentrations elsewhere. Intravitreal injections in small animals have demonstrated uptake by the retinal nerve fiber layer (Yu et al., 2001). However, the optic nerve head represents well less than 1% of the inner surface of the eye and thus should receive less than 1% of the delivered therapeutic. Retroorbital optic nerve injections have also been used to target the optic nerve (Mesentier-Louro et al., 2019), but this is expected to localize drugs behind the lamina cribrosa and it is unclear how such signaling would reach axons or promote regeneration.

In the current study, retroorbital optic nerve injection angle toward optic nerve [C], intravitreal approach to optic nerve head [E], and suprachoroidal tunneling to optic nerve head [G] were most able to localize to the optic nerve head ex vivo. Retroorbital optic nerve injection angle toward optic nerve [C] is similar to previously described studies to utilize an orbital approach (Mesentier-Louro et al., 2019; Tehrani et al., 2018). In our study, we did not consider the difficulty of an orbital approach as these studies were done with enucleated eyes. Endoscopic view of the optic nerve within the orbit may be needed (Yu et al., 2023). Intravitreal approach to optic nerve head [E] was also able to localize to the optic nerve head, however it is unclear if manipulation within the vitreous cavity on the way to the optic nerve head would require vitrectomy in humans. Suprachoroidal tunneling to optic nerve head [G] was also able to localize to the optic nerve head, despite a smaller dose. We chose to further investigate suprachoroidal tunneling to optic nerve head [G] in vivo.

If the optic nerve head is schematized as a cylinder (Figure 3H) (Quigley et al., 1990), with a diameter of 1.5 mm and thickness 0.5 mm, it is apparent that approaching the optic nerve head roughly in line with its thickness axis (as done with retroorbital optic nerve injection angle toward optic nerve [C] and intravitreal approach to optic nerve head [E]) is more difficult because of its smaller dimension. Indeed, resident and attending ophthalmologists would have difficulty achieving the tactile precision needed to maintain a certain depth (Singh et al., 2022; Nakano et al., 2009). Approaching along the greater axis of the optic nerve head is in line with the suprachoroidal space, which is a recently FDA approved method of targeted ophthalmic drug delivery (Chiang et al., 2016;Chiang et al., 2018; Chiang et al., 2017; Chiang et al., 2017; Chiang et al., 2016; Chiang et al., 2016; Yeh et al., 2019; Khurana et al., 2021; Clearside Biomedical, 2021; Patel et al., 2012; Patel et al., 2011). Traversing the suprachoroidal space can be done with suprachoroidal tunneling, which has been used to deliver drugs under the macula for age-related macular degeneration (Tetz et al., 2012; Rizzo et al., 2012; Olsen et al., 2011), and more recently the subretinal space for inherited retinal disorders (Davis et al., 2019;de Smet et al., 2018). Since the suprachoroidal space is a potential space bound by the sclera and choroid with limited thickness (Chiang et al., 2017), tunneling within it is simplified since the radial dimension is virtually eliminated and only the 2 circumferential dimensions matter. Furthermore, the scleral canal, Bruch’s membrane, choroid, and retina are tightly bound at the border tissue of Elschnig, further eliminating degrees of freedom while performing the injection. Peripapillary suprachoroidal delivery can be done with this same technique and may also be able to achieve high concentrations within the optic nerve. We showed in this study that SCONE delivery leveraging these technical advantages can be used to access the optic nerve in vivo.

After optimization in vivo, we were able to achieve an ∼80% success rate in depositing tracer material at the optic nerve head and with no optic nerve head hemorrhage. This was possible with the 45° angled catheter needle tip without catheter sheath and with the 25° angled catheter needle tip with catheter sheath. In attempting to understand the impact of these parameters on success rate, SCONE delivery can be thought of as two sequential procedures: (i) accessing and traversing the suprachoroidal space and then (ii) delivering therapeutics into the optic nerve head. A 25° catheter tip may have made delivering therapeutics into the optic nerve head more successful, but it became much more difficult to traverse the suprachoroidal space reliably and safely, and thus the catheter sheath was necessary. Likewise, the sheath guard may have theoretically made traversing the suprachoroidal space simpler, but it became marginally more difficult to deliver the therapeutic into the optic nerve head. Thus, the base condition and the one that used both performed the best. It is reassuring to note that of the eyes with successful SCONE delivery, there was little to no difference in VEP amplitude and latency between SCONE-treated eyes and the contralateral naïve eye, suggesting a strong measure of safety in this delivery approach.

There were complications that occurred with the current iteration of SCONE delivery, including retinal tears and detachment, vitreous hemorrhage, suprachoroidal hemorrhage, and optic nerve hemorrhage. It is also possible to have retinal or suprachoroidal inflammation, thoughts were not counted in this study. The complications that occurred with SCONE delivery could be thought of in terms of procedure stage also. Most of the complications were due to accessing and traversing the suprachoroidal space, including retinal tears, retinal detachments, vitreous hemorrhage, suprachoroidal hemorrhage. Most of the retinal tears occurred in the periphery (>3mm from optic nerve head) and were likely due to the force required to pass the catheter through the sclerotomy, even with optimization. Only optic nerve hemorrhage can be attributed to delivering therapeutics into the optic nerve head, and occurred in ∼10% eyes with the optimized catheter. A device that facilitates reliable performance of the first few critical steps of SCONE delivery, namely accessing and traversing the suprachoroidal space, could likely reduce complication rates. There is likely a learning curve associated with this procedure, and similarly, such a device could reduce the learning curve. Though VEP showed no/minimal changes, there is the theoretical risk of injury to the optic nerve head with additional treatments; a one-time treatment (or example, to deliver mitochondria in a mitochondrial optic neuropathy or a gene therapy to transfect cells in the optic nerve head long term) may be best suited for this delivery technique.

Limitations of this study include not further exploring the other injection techniques. No optimization and/or iterative engineering was done to improve access to the optic nerve head, and/or improve safety. It is certainly possible that each of the tested injection techniques could be optimized to reach the optic nerve head. Thus, this series of experiments should not suggest that it is impossible to reach the optic nerve head with the other techniques, but rather, that SCONE delivery was able to achieve targeted optic nerve head more easily in this study. Furthermore, ex vivo and postmortem demonstration of injection techniques were used for most of the initial experiments. Though unlikely, it is possible that in vivo access is easier and/or more efficacious than in ex vivo and postmortem eyes. The injection techniques, including SCONE delivery, are likely not powered to comment on safety or efficacy at the level that would be desirable for translation to human use, although they suggest that use for preclinical models to study disease pathophysiology or therapeutic candidate delivery to the optic nerve head should be adequately supported by SCONE. All experiments were performed in rabbit eyes, which are quite different than human eyes, namely lack of collagenous lamina cribrosa and myelination into retina, but the current study establishes a basic technique that can be refined. Further preclinical studies in pigs, dogs, cats, and/or non-human primates will be needed to further optimize SCONE for eventual clinical studies. Further optimization of the procedure and device development will likely increase safety.

Future directions include further optimization of SCONE delivery, testing the other methods more thoroughly, determining the safety of this technique in preclinical animal models, investigating the pharmacokinetics of optic nerve head injections, and studying the efficacy of this technique in animal models of optic nerve head disease. We are actively studying the use of SCONE to deliver neuroprotective agents in optic nerve injury rabbit models. Further optimization by way of custom designed delivery devices that more readily pass the catheter into the suprachoroidal space may better avert retinal tears, which mainly occurred in the periphery and were likely due to the insertion of the catheter into the suprachoroidal space. Because retinal tears may be mitigated with such a device, retinal detachment and vitreous hemorrhage may also be avoided. Suprachoroidal hemorrhage may be avoided with use of the catheter sheath, but further studies will be needed to investigate. Nevertheless, our data suggest that once the catheter (with the 45° bevel angle at least) is within the suprachoroidal space, it stays within that space.

Conclusion

In conclusion, we investigated multiple drug delivery methods to access the optic nerve head. We found 3 methods were able to localize within the optic nerve head: (C) a retroorbital optic nerve injection angled toward the optic nerve head, (E) intravitreal approach to optic nerve head, and (G) suprachoroidal tunneling to optic nerve head. SupraChoroidal-to-Optic-NervE (SCONE) delivery was able to achieve targeted delivery to the optic nerve head with 80% success rate in vivo. Further advancements may reduce complications. Further development of targeted delivery to the optic nerve head is indicated.

Acknowledgements

We thank PL Che and NY Chiang for helping in grading images. We thank DM Moshfeghi for use of the RetCam.

Funding Statement

This work was supported by the National Eye Institute under grants K08-EY033407 and P30-EY026877, Research to Prevent Blindness, and Stanford Center for Optic Disk Drusen.

Authors’ contributions

BC - conception and design; BC, KJ, KH – performed experiments BC, KH, DM, JLG - analysis and interpretation of the data; BC, KH, DM, JLG - drafting of the paper; BC, KH, KJ, RD, YJL, MD, JLG - revising it critically for intellectual content; BC, KH, KJ, RD, YJL, DM, JLG - final approval of the version to be published. All authors agree to be accountable for all aspects of the work.

Disclosure statement

B.C. is the inventor on a patent application filed through Stanford University. An approved plan for managing any potential conflicts arising from this arrangement is in place. No potential competing interest was reported by the other authors.

Data availability statement

Authors agree to make data and materials supporting the results or analyses presented in their paper available upon reasonable request.

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