Non-viral gene therapy for GDNF production in RCS rat: the crucial role of the plasmid dose

Abstract

Glial cell line-derived neurotrophic factor (GDNF) is one of the candidate molecules among neurotrophic factors proposed for a potential treatment of retinitis pigmentosa (RP). It must be administered repeatedly or through sustained releasing systems to exert prolonged neuroprotective effects. In the dystrophic Royal College of Surgeon's (RCS) rat model of RP, we found that endogenous GDNF levels dropped during retinal degeneration time course, opening a therapeutic window for GDNF supplementation. We showed that after a single electrotransfer of 30 μg of GDNF-encoding plasmid in the rat ciliary muscle, GDNF was produced for at least 7 months. Morphometric, electroretinographic and optokinetic analyses highlighted that this continuous release of GDNF delayed photoreceptors (PRs) as well as retinal functions loss until at least 70 days of age in RCS rats. Unexpectedly, increasing the GDNF secretion level accelerated PR degeneration and the loss of electrophysiological responses. This is the first report: (i) demonstrating the efficacy of GDNF delivery through non-viral gene therapy in RP; (ii) establishing the efficacy of intravitreal administration of GDNF in RP associated with a mutation in the retinal pigment epithelium; and (iii) warning against potential toxic effects of GDNF within the eye/retina.

Introduction

Retinitis pigmentosa (RP) is a blinding disease that belongs to the heterogeneous group of inherited retinal dystrophies and results from a diversity of functional mutations.1 Most mutations concern photoreceptor (PR)-specific genes encoding proteins involved in the visual phototransduction pathway (rhodopsine, α- and β-catalytic subunits of cGMP-phosphodiesterase) or in the morphogenesis/stabilization of outer segments (OSs) (peripherin/rds). Mutations were also found in genes expressed in the retinal pigment epithelium (RPE), involved in the visual cycle (lecithin retinol acyltransferase, RPE65) and in PR OS disk phagocytosis (Mertk).

No unique therapy can treat all these conditions, even if striking advances have been made in the field of viral corrective gene therapy in numerous animal models of RP,2 leading to encouraging results in patients suffering from mutations in Rpe65 gene.3, 4 However, this approach can benefit only to a limited number of patients in whom the genetic defect has been characterized. Given the heterogeneity of genetic mutations associated with RP phenotypes, mutation-independent alternative strategies are developed in parallel to treat the symptoms instead of the causes. Numerous studies, carried out in various retinal disease models, have demonstrated the benefit of neurotrophic factors to limit or postpone the retinal neuron degeneration process, a concept called neuroprotection. Since the pioneering work of Faktorovich et al.5 demonstrating that basic fibroblast growth factor could delay PR degeneration in the Royal College of Surgeons (RCS) rat model of RP, many other neurotrophic factors have been shown to slow PR loss in several animal models of inherited and light-induced retinal degeneration including ciliary neurotrophic factor (CNTF),6, 7 brain-derived neurotrophic factor,6 glial cell line-derived neurotrophic factor (GDNF),8 pigment epithelium-derived factor9 and rod-derived cone viability factor.10 Among those, GDNF has appeared as an attractive neuroprotective candidate for the retina, apparently devoid of the toxicity observed with high doses of CNTF11 and of the undesirable proangiogenic properties of fibroblast growth factors.12, 13 An encapsulated cell therapy device (Neurotech, Lincoln, RI, USA) that delivers low and controlled dose of CNTF has been used in clinical trials to rescue PRs and is currently showing promising outcome.7

GDNF belongs to the transforming growth factor-β superfamily14 and is synthesized as a glycosylated precursor form called preproGDNF. After cleavage of its signal sequence, proGDNF is secreted by cells and is activated, by proteolytic cleavage, to generate the GDNF, biologically active as homodimer.15 GDNF was originally purified from a rat glioma cell-line supernatant as a trophic factor for embryonic midbrain dopamine neurons.14 As GDNF was later found to protect many other neuronal sub-populations of the central and peripheral nervous system from cell death,16 hopes have been raised that GDNF could be used as a therapeutic agent to treat several neurodegenerative diseases. Both GDNF and its receptors are synthesized in the retina during development and adulthood17, 18 and are increased after optic nerve transection19 or light-induced retinal degeneration20, 21 suggesting an innate neurotrophic role of GDNF in this tissue. In addition, GDNF has been shown to stimulate survival of newborn rodent PRs in vitro,22, 23, 24 to delay PR OS collapse in vitro25 and to stimulate PR8 and ganglion cell26 survival in vivo.

Many strategies have been used to administer GDNF to the eye, depending on the target cell injured. The intravitreal (IVT) route has mostly been chosen to protect ganglion cells after axotomy, glaucoma or retinal ischemia, whereas the subretinal route has been preferred to prevent PR loss in various models of retinal degeneration such as RP, retinal detachment or light-induced retinal degeneration (see Table 1). However, two studies have demonstrated that the sustained release of GDNF in the vitreous could also be beneficial in models of RP based on mutations in PR-specific genes,27, 28 but none has shown efficacy in RP caused by a mutation in the RPE.

Table 1 Summary of the use of GDNF within the eye as a function of drug-delivery systems, routes of administration and doses used

The RCS rat is an autosomal recessive RP model because of a null mutation in the RPE-specific gene encoding the receptor tyrosine kinase Mertk.29, 30 It is a relevant animal model since mutations in this gene have also been found in RP patients.31 The incapacity of RPE cells to phagocytose PR OSs leads to their accumulation in the subretinal space and subsequent PR loss,32, 33 mainly by apoptosis.34 Retinal degeneration begins at the third week of postnatal (PN) life and is completed around PN 90 days. The gradual shortening of PR OS and inner (IS) segments and the thinning of the outer nuclear layer (ONL) is accompanied by loss of visual and electroretinographic (ERG) responses.35, 36

Besides the choice of the neurotrophic factor, a crucial issue remains the way to deliver it in a sustained manner to its target inside the eye. Neurotrophic factors including GDNF37 have short half-lives within the eye and injections should be repeated to achieve any efficacy.8, 26 Various strategies using microspheres,27 cell transplantation28, 38, 39 or viral gene therapy11, 40 have been attempted to release GDNF. However, no attempt has been made to date to deliver GDNF via a non-viral gene transfer method in any inherited model of RP.

In the ciliary muscle electrotransfer (ET) technique, a focal electric field is applied to transfer plasmid DNA into ciliary muscle fibers,41, 42 which are turned into a biofactory for the secretion of any molecular weight proteins for several months, mainly into the vitreous.42 This technique was shown to be efficient on the short-term41, 43, 44 and mid-term44 to deliver anti-tumor necrosis factor molecules in two rat models of intraocular inflammation.

By electrotransferring a GDNF-encoding plasmid into the ciliary muscle of the dystrophic RCS rat, our aims were to evaluate: (1) if this method can be used as a drug-releasing system for neurotrophic factors within the eye; (2) its usefulness to slow retinal degeneration in RP; (3) the efficacy of GDNF delivery through a non-viral gene therapy in RP; and (4) the efficacy of IVT sustained release of GDNF in a model of RP suffering from a mutation in the RPE.

Results

Modulation of GDNF and CNTF expression during retinal degeneration

Intraocular fluid amounts of GDNF and CNTF were compared between dystrophic RCS rats (rdy−, p+) and congenic non-dystrophic control rats (rdy+, p−) 33 days, 78 days and 10 months after birth (Figure 1). Whereas GDNF amount was statistically unchanged among ages in healthy control eyes (46.4±5.5, 37.8±4.2 and 35.1±6.2 pg at 33 days, 78 days and 10 months PN, respectively), variations were observed in dystrophic eyes (Figure 1a) at PN78 and 10 months. Indeed, GDNF amount statistically decreased to 15.8±2.6 pg at PN78, and then increased to 68.7±2.7 pg at 10 months of age. In the neuroretina, GDNF levels parallel vitreous levels (Figures 1c and d) and show that the decrease in GDNF expression at PN78 is not due to a reduced total number of retinal cells but to a decreased production. In healthy eyes, CNTF amount remained constant among ages (943±119, 853±220 and 931±265 pg at PN33 days, PN78 days and PN10 months, respectively) (Figure 1b). In dystrophic eyes, amount of CNTF showed an early decrease at PN33 days (427±47 pg) and then gradually increased to reach 1016±212 and 2269±269 pg at PN78 days and PN10 months, respectively. At this latter time point, CNTF amount was statistically higher and about twice that measured in non-dystrophic controls.

Figure 1
figure1

Modulation of endogenous GDNF and CNTF expression in non-dystrophic and dystrophic RCS rats. Endogenous levels (in pg ml–1) of GDNF (a) and CNTF (b) in ocular media were measured by ELISA in non-dystrophic and dystrophic RCS rats at 33 days, 78 days and 10 months of age (n=4 eyes per condition). Total amounts of protein in the ocular media were calculated by multiplying the measured protein concentrations by ocular media volume (50 μl). Results are expressed as mean±s.e.m. Statistical analyses: Mann–Whitney U-test. #P<0.05 and ##P<0.01 vs the non-dystrophic age-matched group; *P<0.05. Endogenous GDNF expression in neuroretinal extracts at 33 and 78 days of age (n=2 eyes per condition) visualized on immunoblot (c) (top lane: GDNF; bottom lane: β-tubulin reporter protein) and semi-quantified by densitometric analysis (d). AU, arbitrary unit.

Production of rGDNF by RPE cells transfected in vitro with pVAX2–rGDNF

pVAX2–rGDNF plasmid was constructed by subcloning the complementary DNA of GDNF, amplified from rat brain mRNA, in a pVAX2 backbone under a cytomegalovirus-β promoter (circular plasmid map in Figure 2a, see Materials and Methods section for details). Immunoblot performed on ARPE-19 cell lysates transfected with pVAX2–rGDNF (Figure 2b) shows that rat GDNF (rGDNF) was synthesized at the expected size of the rat preproGDNF (24 kDa, white arrow). This intracellular form was of higher molecular weight than that of the 18 kDa secreted form detected in the culture supernatants by western blot (Figure 2c, white arrow). In lanes loaded with the recombinant rat GDNF protein (rrGDNF, Figure 2b), used as a positive control, the molecular weight of 18–22 kDa was that of the monomeric glycosylated GDNF (one asterisk), whereas the unglycosylated GDNF monomer had an apparent molecular weight of 15 kDa (sharp);14, 15 additional bands of higher molecular weight stood for dimeric forms of glycosylated (two asterisks) and unglycosylated GDNF. In culture supernatants, GDNF was secreted in its monomeric glycosylated mature form forming functional homodimers (Figure 2c, white and grey arrows, respectively). Whereas GDNF was synthesized and secreted (Figures 2b and c) in a plasmid dose-dependent manner, no GDNF could be detected within cells or in culture media from cells treated with the higher dose of the pVAX2 plasmid backbone. These results show that the rGDNF protein encoded by the newly generated plasmid was synthesized (preproGDNF), secreted (proGDNF) and maturated (GDNF) as in physiological conditions.

Figure 2
figure2

Functionality of rGDNF-encoding plasmid on in vitro culture cells. (a) Circular plasmid map of pVAX2–rGDNF plasmid: rGDNF complementary DNA (cDNA) was cloned in the pVAX2 backbone, downstream of the cytomegalovirus (CMV)-β promoter and upstream of the bovine growth hormone (BGH) polyA signal, using BamHI and EcoRV restriction enzymes. (b) Western blot analysis of rGDNF expression in ARPE-19 cell culture lysates 3 days after transfection with pVAX2 (4 μg) or with increasing doses of pVAX2–rGDNF (1, 2 and 4 μg) using the calcium phosphate method. rrGDNF was loaded in the three last lanes (15, 30, and 70 ng). (c) Western blot analysis of rGDNF secretion in ARPE-19 cell culture supernatants 4 days after transfection with the calcium phosphate method. The amount of plasmid used for transfection, expressed in microgram (μg), is stated below each lane. Plasmid dose-dependent rGDNF synthesis (b) and secretion (c) were visualized on immunoblots of pVAX2–rGDNF transfected samples (white and grey arrows on the left side). rGDNF was not detected in pVAX2 transfected samples. rrGDNF used as a positive control (black arrows on the right side) was detected as a monomeric glycosylated rrGDNF (*18–22 kDa) or a dimeric glycosylated rrGDNF (**). The recombinant unglycosylated rGDNF monomer (#) has an apparent molecular weight of 15 kDa.

Long-term GDNF secretion by transduced ciliary muscle cells

To determine the amount of rGDNF produced by the transduced ciliary muscle without interference with the endogenous GDNF (Figure 1a), we measured GDNF produced by anterior segment explants maintained in ex vivo culture for 24 h (Figures 3A–D, see Materials and Methods section for details), and collected from eyes enucleated 6 days and 7 months after in vivo ET of pVAX2 or pVAX2–rGDNF. As shown on Figure 3E, the relative increase in rGDNF production between pVAX2–rGDNF and pVAX2-treated samples was 66.7±8.1 pg every 24 h at 6 days and 21.3±2.6 pg every 24 h at 7 months post-ET demonstrating that production of rGDNF was still efficient at this late time point after transfection.

Figure 3
figure3

rGDNF production by the ciliary muscle in anterior segment explants cultured ex vivo. (A) Preparation of anterior segment explants. Eyes were collected 6 days (n=5 eyes per condition) or 7 months (n=3 eyes per condition) after ciliary muscle ET of 30 μg of pVAX2 or pVAX2–rGDNF in dystrophic RCS rat. (a1) Freshly enucleated eyes were first incubated for 15–30 min at 37 °C in conditioning medium (detailed in material and methods). (a2) For each eye, dissection was carried out posterior to the limbus as illustrated by the scattered line. Posterior segment containing the sclera (sc), the choroid (ch) and the retina (ret) was discarded and the lens (l) was carefully removed from the rest of anterior segment. (a3) Anterior segment explants made of the cornea (co), the iris (ir) and the ciliary body (cb), the latter containing the transfected ciliary muscle fibers, were incubated for 15 additional minutes at 37 °C in conditioning medium before being cultured ex vivo. (B) Schematic representation of the different steps of ex vivo culture of anterior segment explants: (1) explants were placed in a 96-well plate (200 μl medium per well) and (2) incubated at 37 °C for 24 h. (3) Culture medium was collected at 24 h and used for (4) rGDNF quantification by ELISA. (C, D) Photos of anterior segment explants cultured ex vivo in a 96-well plate taken at low (C) or high (D) magnification. The cornea (*), seen through the pupil, was in contact with the bottom of the culture well whereas the iris (ir) and the ciliary body (cb) faced toward the top of the well. (E) Kinetics of rGDNF production arising from pVAX2–rGDNF ET. ELISA quantification of rGDNF was performed in anterior segment explants supernatants collected from pVAX2 and pVAX2–rGDNF-treated samples. For each time point, values obtained (in concentration) were normalized by the volume of medium used for ex vivo culture to deduce the amount of rGDNF produced (in pg) per 24 h (stated on the graph). Results, are expressed as mean±s.e.m. Statistical analyses: Mann–Whitney U-test. * P<0.05 vs pVAX2. ND, not detectable.

Survival effect in pVAX2 ET eyes

In a preliminary experiment, the effect of ET in the ciliary muscle using the non-coding pVAX2 plasmid backbone (control ET) was investigated on retinal morphology. Morphological analyses carried out at PN75 showed no retinal disorganization or gross structural damage in eyes submitted to control ET 55 days before, as compared with age-matched untreated dystrophic eyes. However, the retinal structure was better preserved in control ET eyes (n=5) than in untreated dystrophic control eyes (n=3) (not shown). These observations were confirmed by measurements of retinal layers. Whereas the inner nuclear layer (INL) thickness was unchanged in the superior hemisphere (29.9±1.5 vs 25.1±1.3 μm in pVAX2 ET-treated and -untreated eyes, respectively, P>0.05), the ONL in control ET eyes was almost twofold thicker than that of untreated eyes (14.4±1.5 vs 8.1±1.2 μm, P=0.005). pVAX2 ET also significantly enhanced the thickness of PR ISs in the superior pole (P<0.0001). Data obtained from measurements in the inferior hemisphere did not show any statistical difference between both groups, whatever the layer considered.

To evaluate the functional consequence of control ET on RCS rats, electrophysiological analyses were carried out at PN53 (Figure 4) and PN75 (not shown). At PN53, control ET significantly enhanced scotopic b-wave amplitude by almost twofold (352.0±26.0 vs 186.0±23.8 μV in pVAX2 ET-treated and untreated eyes, respectively; Figure 4b) whereas a-wave amplitude was unchanged (Figure 4a). At a later time point (PN75), the b-wave amplitude was still higher in pVAX2 ET-treated eyes than in untreated dystrophic eyes (114.0±16.1 vs 24.7±13.8 μV, respectively). Note that at PN92, GDNF amount in the ocular media was statistically higher in the pVAX2 ET-treated eyes than in untreated control eyes (30.7±3.9 vs 13.1±5.2 pg; n=4 eyes per group; not shown). These results show that the ET procedure by itself, without any trophic factor genes transduced, induces significant protective effects on the retina at least 2 months after current application. The effect of GDNF-encoding plasmid ET was further evaluated as compared with control ET.

Figure 4
figure4

Electrophysiological responses of the retina induced by plasmid backbone ET in the RCS rat. Scotopic ERG a-wave (a) and b-wave (b) amplitudes (in μV) were recorded in PN53 dystrophic RCS rats treated by pVAX2 ET (30 μg) and in age-matched untreated rats (n=5 eyes per group). Results are expressed as mean±s.e.m. Statistical analyses: Mann–Whitney U-test. *P<0.05 vs untreated control eyes.

Morphological and functional rescue of PR induced by GDNF delivery

Histological analysis of retinas from RCS rats treated by pVAX2 or pVAX2–rGDNF ET at PN16-20 was performed at PN ages of 51, 70 and 87 days. As shown on Figure 5, PR ISs and ONL were better preserved in the superior hemisphere of pVAX2–rGDNF-treated eyes than in control ET at PN51 and PN70. At PN87, only focal preserved areas were still observed in the superior retina. As the rate of degeneration is higher in the inferior than in the superior hemisphere in RCS rats, thickness of retinal layers was measured separately for each pole (Figure 6). In the inferior hemisphere, no statistical difference was observed between eyes treated with pVAX2–rGDNF and pVAX2 ET, whatever the age or the retinal layer is considered (P>0.05). On the contrary, a statistically significant rescue effect was observed in the superior hemisphere at PN51: the thickness of the INL (26.3±1.5 vs 19.6±0.5 μm), the ONL (27.7±1.1 vs 22.2±0.9 μm) and the IS (8.1±0.4 vs 6.4±0.3 μm) was higher in pVAX2–rGDNF-treated eyes than in pVAX2-treated ones, with increases of 35%, 25% and 27%, respectively. At PN70, the INL thickness was still higher (27.4±0.6 vs 20.1±0.6 μm) and a trend of increase was still observed for the IS thickness (1.8±0.4 vs 0.9±0.2 μm) but the mean ONL thickness calculated on the whole superior retina was not statistically different between both groups (9.4±1.0 vs 8.5±0.7 μm). At PN87, no difference could be detected between pVAX2–rGDNF and pVAX2-treated eyes (INL: 21.4±0.7 vs 20.8±0.9 μm; ONL: 5.4±0.6 vs 4.7±0.7 μm; IS: 1.5±0.4 vs 1.1±0.5 μm).

Figure 5
figure5

Time-dependent retinal morphological rescue effect of rGDNF delivery. Histological sections of retinas from PN51 (a, d), PN70 (b, e) and PN87 (c, f) dystrophic RCS rats treated by ET of 30 μg of pVAX2 (ac) and pVAX2–rGDNF (df). Photos are taken in the superior retina, at 2 mm from the optic nerve head. GCL, ganglion cell layer.

Figure 6
figure6

Morphometric analyses of rGDNF delivery rescue effect. INL (ad), ONL (eh) and PR ISs (il) thicknesses were measured every 500 μm from the optic nerve head (ONH) in the superior (sup) and inferior (inf) retina of dystrophic RCS rats treated by pVAX2 (open circles and bars) and pVAX2–rGDNF (filled circles and bars) ET. Analyses were performed at PN51 (a, b; e, f; i, j; n=6 eyes per group), PN70 (c, g, k; n=4 eyes per group) and PN87 (d, h, l; n=6 eyes per group) and results were expressed as mean±s.e.m. Profiles (a, e, i): mean thickness of the different layers along the whole retina at PN51. Histograms (bd, fh, j–l): mean thickness of the different layers in the superior and inferior hemispheres at the three different time point. Statistical analyses: Mann–Whitney U-test. **P<0.01 and ***P<0.001 vs pVAX2-treated eyes.

As expected, at PN53, a significant increase in 60% in a-wave amplitude was observed in the group treated with GDNF (96.3±10.3 μV) as compared with control ET (60.2±8.5 μV) (Figure 7a). Moreover, the b-wave amplitude was significantly higher in GDNF-treated (475±41 μV) than in pVAX2-control eyes (352±26 μV) (increase estimated at 35%) (Figure 7b). At PN75, a slight trend of increase in a-wave and b-wave amplitudes was still detectable in pVAX2–rGDNF-treated eyes (29.7±11.3; 135±38 μV, respectively) compared with pVAX2-treated eyes (23.1±4.0; 114±16 μV, respectively) (Figure 7). At PN87, no statistical difference could be detected in the a-wave (17.1±3.2 vs 24.5±4.9 μV) and b-wave (83±7 vs 110±14 μV) amplitude between the two groups (Figure 7). Optokinetic tests (Figure 8) showed a significant increase in visual acuity in the pVAX2–rGDNF-treated group (0.56±0.02 cycles per degree (c/d) vs 0.48±0.02 c/d) at PN70 but not at PN92 (0.38±0.03 vs 0.35±0.03 c/d).

Figure 7
figure7

Increase in retinal electrophysiological functions induced by rGDNF delivery. Scotopic ERG a-wave (a) and b-wave (b) amplitudes (in μV) were recorded in PN53 (n=5 eyes per group), PN75 (n=4 eyes per group) and PN87 (n=3 eyes per group) dystrophic RCS rats treated by pVAX2 (white circles) and pVAX2–rGDNF (black circles) ET (30 μg). Results are expressed as mean±s.e.m. Statistical analyses: Mann–Whitney U-test. *P<0.05 vs pVAX2-treated eyes.

Figure 8
figure8

Longer preservation of spatial visual acuity induced by rGDNF delivery. Spatial visual acuity threshold (expressed in c/d) were recorded in PN70 (a) (n=4 eyes per group) and PN92 (b) (n=5 eyes per group) dystrophic RCS rats treated by pVAX2 and pVAX2–rGDNF ET (30 μg) using the optometry head tracking apparatus. Results are expressed as mean±s.e.m. Statistical analyses: Mann–Whitney U-test. *P<0.05 vs ET pVAX2.

Higher pVAX2–rGDNF dose results in lower neuroprotective effects

We have recently published that using ciliary muscle ET, protein production in the ocular media could be increased either by increasing the plasmid dose or by performing multi-sites treatment.42 We therefore evaluated the effect of increased dose of GDNF using two ET of 15 or 30 μg of pVAX2–rGDNF, or pVAX2 as control, and analyses were performed at PN70. For all measured parameters, no differences were observed between eyes treated with two ET of either 15 or 30 μg of the naked pVAX2 plasmid. Thus, the later condition was used as control in this experiment (Figure 9a). Unexpectedly, while no difference in the morphology of the retina could be observed in eyes treated with two ET of 15 μg pVAX2–rGDNF as compared with the control (two ET of 30 μg of pVAX2) (Figures 9a and b), a thinning of the retinal nuclear layers was observed after two ET of 30 μg of pVAX2–rGDNF (Figure 9c). As shown on Figure 10, no statistically significant variations in INL thickness were noticed between eyes treated by ET of 2 × 30 μg pVAX2, 2 × 15 μg pVAX2–rGDNF and 2 × 30 μg pVAX2–rGDNF, with values of 30.5±1.3, 32.2±1.6 and 28.3±1.3 μm, respectively. The ONL (16.8±1.9 μm) and ISs (8.3±1.7 μm) thickness of eyes treated by 2 × 15 μg pVAX2–rGDNF were neither significantly different from that of eyes treated by 2 × 30 μg pVAX2 (ONL: 17.6±1.7 μm; IS: 9.2±1.0 μm). On the contrary, ET of 2 × 30 μg pVAX2–rGDNF significantly decreased the ONL thickness by 40% (10.6±1.7 μm) and the ISs thickness by 43% (5.2±1.2 μm) as compared with the controls.

Figure 9
figure9

Retinal morphological effects of increasing doses of pVAX2–rGDNF. Histological sections of retinas from PN70 dystrophic RCS rats treated by two ET of 30 μg of pVAX2 (a) or 2 ET of 15 μg (b) and 30 μg (c) of pVAX2–rGDNF. Photos are taken in the superior retina, at 2 mm from the optic nerve head. DZ, debris zone; GCL, ganglion cell layer.

Figure 10
figure10

Morphometric analyses of retinal layer thicknesses as a function of the dose of pVAX2–rGDNF. INL (a), ONL (b) and PR ISs (c) thicknesses were measured in the superior retina of PN70 dystrophic RCS rats treated by 2 ET of 30 μg of pVAX2 and by two ET of 15 μg and 30 μg of pVAX2–rGDNF (n=3 eyes per group). Results were expressed as mean±s.e.m. Statistical analyses: Mann–Whitney U-test. *P<0.05 and **P<0.01 vs pVAX2-treated eyes. No statistical difference could be detected in the inferior retina (not shown).

Scotopic ERGs were performed at PN70 on the same animals before killing. No difference in the a-wave amplitudes was observed between the three groups: 2 × 30 μg pVAX2 (23.8±1.9 μV), 2 × 15 μg pVAX2–rGDNF (26.7±2.7 μV) and 2 × 30 μg pVAX2–rGDNF (20.9±3.7 μV) (Figure 11a). However, compared with the 2 × 30 μg pVAX2-treated group (231±5 μV), a 38% decrease in b-wave amplitude was observed after treatment with 2 × 30 μg pVAX2–rGDNF group (144±29 μV) whereas no statistical variation was detected in the 2 × 15 μg pVAX2–rGDNF group (199±17 μV) (Figure 11b), demonstrating that only the higher dose of GDNF induced a decrease in the b-wave amplitude.

Figure 11
figure11

Retinal electrophysiological effects of increasing doses of pVAX2–rGDNF used for ET. Scotopic ERG a-wave (a) and b-wave (b) amplitudes (in μV) were recorded in PN70 dystrophic RCS rats treated by 2 ETs of 30 μg of pVAX2 (white bars) or 2 ETs of 15 μg (grey bars) or 30 μg (black bars) of pVAX2–rGDNF. Results are expressed as mean±s.e.m. (n=3 eyes per group). Statistical analyses: Mann–Whitney U-test. *P<0.05 vs eyes treated by two ET of 30 μg pVAX2.

Discussion

Ciliary muscle ET allows the intraocular sustained secretion of any therapeutic proteins.42 Its efficacy was shown over the short-term41, 43, 44 and mid-term44 in two rat models of intraocular inflammation, by using secreted tumor necrosis factor-α soluble receptors. Here, we evaluated this method to produce GDNF in the RCS rat, a model of inherited retinal degeneration. Our choice for GDNF in this model relied on our preliminary experiments showing that contrarily to what was reported in other models of RP,8, 28, 45 retinal degeneration was accompanied by a drop in rGDNF levels in the RCS rat, hypothesizing that a GDNF supplementation could be beneficial. At a very late stage of retinal degeneration when no PRs could be saved anymore (10 months of age), levels of GDNF and CNTF were very high, defining a potential therapeutic window for GDNF.

We have previously shown that ciliary muscle ET was safe and did not induce inflammation or retinal structural damages in adult rats.41, 42, 43, 44 In this study, we show that it is also safe in young RCS rats (16–20 days of age) and that no deleterious effects on retinal layers thickness and electrophysiological responses could be detected in comparison with untreated animals, showing that neither the presence of plasmid DNA nor the ET procedure worsen the course of retinal degeneration in the RCS rat. Interestingly, control ET was not only safe but was also neuroprotective with a significant rescue effect observed until at least PN75 at the histological and electrophysiological levels compared with untreated control eyes.

This effect could result from the injection itself, as previously reported in the RCS rat or in light-induced degeneration models, using subretinal or IVT injections.5, 46, 47 Ocular injections could indeed trigger the release of basic fibroblast growth factor, CNTF or brain-derived neurotrophic factor, as well as GDNF that is expressed in conjunctival and limbal epitheliums48, 49 and in the retina.18 In our experiments, ET by itself induced GDNF production in the vitreous for >70 days after the procedure, which suggests an active secretion, rather than a release.

The prolonged trophic effect of control ET could also result from the electrical stimulation itself. Transcorneal electrical stimulation applied weekly for 2 to 6 weeks prolonged PR survival and delayed the decrease in retinal function in RCS rats.50 Using a subretinal implant, electrical stimulation also preserved the ERG responses51, 52 as well as the visual-evoked responses in the superior colliculus53 of RCS rats. In our experimental conditions, the electrical field in between the two electrodes did not cover the retina and was applied only for <2 s, which makes it very unlikely that such retinal electrostimulation could occur. However, the effects of ciliary region electrostimulation, on PRs survival deserves to be further investigated.

The effect of the overexpression of GDNF after pVAX2–rGDNF ET was therefore compared with control ET (pVAX2 ET). Using ex vivo culture of anterior segment explants collected at different time points after in vivo ET, we could ascertain that the transduced ciliary muscle fibers actively produced GDNF for at least 7 months. A constant production is required for GDNF that has a relatively short half-life in the vitreous, estimated around 37 h in the pig eye.37

To further clearly separate the GDNF produced by the transduction of ciliary muscle fibers from additional non-dissociable combined effects of the procedure itself that can enhance endogenous GDNF levels, we have measured the production of rGDNF after in vivo electroporation of 50 μg of plasmid in rabbit eyes and found that around 52 pg per eye (vitreous) of GDNF were produced at 80 days after the procedure (data not shown).

As compared with the pVAX2 control ET, 30 μg of pVAX2–rGDNF ET at PN16-20 in RCS rat eyes induced a neuroprotective effect at both the histological and electrophysiological (a-wave and b-wave) levels at PN53 and at the functional level (visual acuity) at PN70. No more effect was observed at PN75 and PN90.

Such neurotrophic effect of GDNF was already shown in various rodent models of inherited retinal degeneration. GDNF was administered into the subretinal space in the rd1 mouse,8, 45 by transplantation of cells genetically engineered to secrete GDNF in the RCS rat38, 39 as well as by injection of recombinant adeno-associated virus–GDNF viral vectors in TgN S334ter-4 and RCS rats.11, 40 Only two studies have demonstrated that GDNF could exert similar effects when delivered in the vitreous, one in the rd1/rd1 mouse using microspheres27 and the other one in the TgN S334ter-4 rat using transduced stem cells.28 Moreover, only one recent publication has demonstrated that non-viral gene therapy, by means of subretinal non-viral PEG-POD-GDNF injection, is a feasible approach to produce GDNF within the eye and to slow PR degeneration in a light-induced degeneration model in the mouse.54 This is therefore the first study establishing that GDNF delivery through non-viral gene therapy can be efficient in an inherited model of RP and showing that IVT delivery of GDNF can efficiently delay PR loss in a RP model caused by a mutation in the RPE cell layer.

How GDNF could exert neurotrophic effects when delivered in the vitreous? GDNF interacts with GDNF family receptor alpha-1 (GFRα1)22, 55 and mediates its biological effects through the interaction of GDNF–GFRα1 complex with the membrane-bound tyrosine kinase receptor, RET.56 Although the preferred interaction appears to be GDNF–GFRα1, GDNF signaling can also be mediated by GDNF–GFRα2 interaction with RET57 or GDNF–GFRα1 interaction with another transmembrane or linker protein than RET.58 In the retina, GDNF receptors GFR-α1,20, 59, 60 GFR-α2,20 and RET17 have been detected on PRs. Retinal Müller glial cells have also been shown to express GFR-α1,17, 60, 61 GFR-α2,59 and RET.17, 61 Direct (on PR)22, 24, 25 and indirect (through Müller cells) mechanisms have therefore been proposed to describe the neuroprotective effect of GDNF on PR cells. Exogenous GDNF activates retinal Müller glial cells8 and increases their production of basic fibroblast growth factor,59, 61 brain-derived neurotrophic factor,59 as well as GDNF itself.59 Moreover, intraocular administration of GDNF enhances the expression of the glutamate transporter GLAST within the retina,62, 63 in retinal Müller glial cells located around the degenerating PR in a RP model.45 The sustained intravitreous release of GDNF can exert neurotrophic effects through these multiple ways on PRs. In the RCS rat specifically, GDNF protective effects could also result from the activation of microglial cells that could enhance the clearance of PR debris.

In our experiments, rGDNF was continuously produced for 7 months after pVAX2–rGDNF ET, but its neurotrophic effect over the pVAX2 control treatment was only beneficial until PN70 and not maintained at PN75-PN90. In the same model, Buch et al.11 have shown that more PR could be detected up to 3 months (that is, PN100) after subretinal administration of AAV–GDNF in PN12 RCS rats. However, the effect of GDNF itself could not be ensured in this study because comparison was made with untreated control eyes instead of sham-injected or empty virus-injected eyes. In the same study, scotopic–b-wave amplitude was increased in treated eyes only until PN54 as compared with untreated eyes, with no more significant effect at PN70.11 In our experiments, GDNF-treated eyes still had better optokinetic testing results as compared with control ET until PN70, indicating that GDNF produced either by ciliary muscle ET or by subretinal viral strategies have at least comparable effects. The limited effect of GDNF could be related to the mechanisms of the disease itself but also to inappropriate amount of GDNF.

In an attempt to increase the amount of GDNF, we doubled the plasmid dose, as we previously have shown that the plasmid dose and the number of ET procedures correlate well with the dose of proteins produced.42 Comparison was thus performed between a total amount of 30 and 60 μg of plasmid using two ET of 15 and 30 μg at distinct sites.42 Increasing the dose of non-coding plasmid or using two control ET did not induce any statistical difference at each level of analysis at PN70. Surprisingly, when increasing the dose of pVAX2–rGDNF plasmid to 60 μg, a significant reduction of scotopic b-wave amplitude, ONL and PR IS thickness was observed as compared with the same dose of control plasmid ET demonstrating an inverse dose-response effect. Such potential deleterious effect of GDNF dose has not been previously reported in the eye but was described in other parts of the central nervous system such as in the inner ear64 and in the brain65, 66 using continuous GDNF-releasing systems. GDNF induced a dose-dependent protective effect on sensory cell preservation and hearing function when applied directly into the guinea pig cochlea in a noise-induced hearing loss model.64 However, at the highest dose of 1 μg ml–1 (12 ng day–1), GDNF exacerbated hair cell and hearing loss along the cochlea in noise-exposed ears but not in healthy ears, indicating that GDNF could increase hair cells sensitivity to noise damage in the cochlea.64 In the brain, a 6-month toxicologic study conducted in healthy rhesus monkeys demonstrated that chronic intraputamenal infusion of GDNF at the highest dose of 100 μg day–1 for 3 months followed by a 3-month treatment-free recovery period induced cerebellar Purkinje cell loss.65 This toxicity was neither observed in brain treated by lower doses of GDNF and submitted to the same recovery period nor in tissues continuously exposed to the highest dose of GDNF for 6 months, without any recovery period, suggesting that lesions could result from withdrawal of high GDNF dose.66

In light of our results, we cannot exclude that the endogenous decrease in GDNF levels associated with the continuous decline of ciliary muscle fibers production from the highest dose could account for the deleterious effects, as suggested in the brain.65, 66 Whether similar negative effects on retinal histology and function would be observed in other model of RP or in normal rat eyes, remains to be determined. To date, analyses performed 7 days after IVT injection of 100 ng GDNF in the pig,37 8 weeks after subretinal administration of recombinant adeno-associated virus–GDNF in the mouse11 or 1 year after IVT administration or recombinant adeno-associated virus–GDNF in the rat67 did not mention any sign of ocular toxicity or visual function disturbances in healthy eyes. GDNF was also well tolerated in several animal models of retinal pathology (see Table 1). No toxic effect has been reported after a single IVT injection of 5 μg68 or after three IVT injections of 2 mg26 in the rat, neither after two subretinal injections of 330 ng in the mouse8 but analyses were performed on the short-term (14 days after administration at the best). Long-term delivery of GDNF has been evaluated using drug-releasing systems such as microspheres, cellular therapy and gene therapy. Unfortunately, little information is available regarding the doses produced in these studies making comparison difficult (see Table 1).

The mechanism by which ET GDNF could become deleterious for retinal cells remains elusive because relatively high concentrations of GDNF had protective effects against PR OSs collapse in short-term in vitro assays.25 Direct as well as indirect effects can be hypothesized, such as a decrease in dopamine synthesis within the retina, as observed with sustained GDNF release in the brain,69 which could in turn decrease visual performances since retinal dopamine has a pivotal role in mediating visual functions.70 Another possibility is that it is the combination of ET with high dose of GDNF, which may drive the toxicity. Further studies are required in vivo to confirm such inverse dose-response effects in other models and to elucidate the mechanisms.

Materials and methods

Plasmids

Total mRNA was extracted from rat brain using Trizol reagent (Invitrogen, Cergy-Pontoise, France) according to the manufacturer's instructions. In all, 10 μg of total mRNA were subjected to reverse transcription with a 20-bp polyT DNA primer and the Superscript II enzyme (Invitrogen) in a 20 μl reaction volume. Then, 2 μl of this reaction were subjected to a PCR amplification reaction using the rGDNF-specific primers sense: 5′-IndexTermCGGGATCCGCCACCATGAAGTTATGGGATGTCGT-3′ and antisense: 5′-IndexTermGGAATTCTCAGATACATCCACACCGTT-3′ derived from the Genbank sequence N° L15305.1, with the Accuzyme enzyme (Bioline, Abcys, Paris, France). The resulting 657-bp product was digested with BamH1. This fragment was then ligated into a BamHI–EcoRV digested pVAX2 fragment, downstream of a cytomegalovirus-β promoter, to give pVAX2–rGDNF (Figure 2a). The pVAX2 backbone was described previously.71 The sequence was checked by DNA sequencing.

All plasmids were amplified in Escherichia coli bacteria and prepared endotoxin-free (EndoFree Plamid Kit; Qiagen, Courtaboeuf, France). Plasmids were diluted in endotoxin-free water containing 77 mM of NaCl (half saline) (saline, NaCl 0.9%, Versol, Laboratoire Aguettant, Lyon, France) as previously described.42 The concentration of DNA was determined by spectroscopy measurements (optical density at 260 nm).

In vitro assessment of GDNF synthesis and secretion

A spontaneously immortalized human RPE cell line displaying many differentiated properties typical of RPE cells in vivo (ARPE-19 (ATCC CRL-2302))72 was cultured in Dulbecco's modified Eagle's medium:F12 supplemented with 10% fetal calf serum and antibiotics (100 U ml–1 penicillin; 100 μg ml–1 streptomycin; Invitrogen). The cultures were kept at 37 °C in a 5% CO2 humidified atmosphere. Cells (passage 30 to 33) were seeded in 12-well plates (1 ml per well) at a density of 50 000 cells cm–2 and grown for 24 h. Medium was removed and cells were then transfected with 0.25, 0.5, 1, 2 or 4 μg of plasmid (pVAX2 or pVAX2–rGDNF) using the calcium phosphate transfection method.73, 74 Untreated cells and cells treated only with the transfection solution (CaCl2/HEBS) were used as additional controls. After 24 h, cells were washed with phosphate-buffered saline (PBS) and medium was refreshed (400 μl per well). At 72 h, adherent cells were washed and lysed in 300 μl of lysis buffer (MOPS SDS Running Buffer + LDS Sample Buffer, Invitrogen). Cell lysates were immediately heated for 5 min at 100 °C and stored at room temperature until use. At 96 h, cell culture supernatants of the remaining wells were collected, clarified by centrifugation and stored at −20 °C until use.

In vivo electroporation to rat ciliary muscle

Male and female pigmented dystrophic RCS rats (rdy−, p+) and congenic albino non-dystrophic control rats (rdy+, p−), were used and handled in accordance with the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research. Rats were anesthetized with intramuscular injection of Ketamine (88 mg kg–1) (Virbac, Carros, France) and Largactil (0.6 mg kg–1) (Sanofi-Aventis, Paris, France) before ciliary muscle ET. At the end of the experiments, rats were killed with a lethal dose of pentobarbital.

Each set of experiment was performed on rats from a given litter. Plasmid ET was performed in dystrophic RCS rats at PN days 16–20 (PN16–20) (weight of rats 30 g), an age preceding major onset of PR loss.34 Plasmid (30 μg in 10 μl half saline) was injected in the ciliary muscle using a 30-gauge disposable needle (BD Micro-Fine syringe, NM Médical, Asnière, France) transsclerally posterior to the limbus as recently described.42 For electric pulse delivery, a specially designed iridium/platinum electrode was introduced into the preformed intrasscleral tunnel. The semi-annular iridium/platinum counter anode electrode was placed around the cornea, facing the active electrode. ET was performed with eight electrical pulses (15 V voltage, 10 ms in duration, 180-ms interval), generated by the 830 BTX electropulsator (Genetronics, San Diego, CA, USA) as previously described in the adult rat eye.42 In all experiments, eyes were treated at the temporal side of the eyeball. For multiple ET, injections were performed at the temporal and nasal sides of the ciliary muscle and electrical field was applied immediately after each injection.

Quantification of GDNF endogenous levels in ocular media

Dystrophic pigmented RCS rats (rdy−, p+) and congenic non-dystrophic agouti control rats (rdy+, p−) were euthanatized 33 days, 78 days and 10 months after birth. Eyes were removed, rinsed in NaCl 0.9% and dried. Once opened under an operating microscope, aqueous humor and vitreous were collected and pooled for each rat eye. Intraocular media were immediately centrifuged and the cell-free fractions stored at −20 °C until being assayed by enzyme-linked immunosorbent assay (ELISA).

Ex vivo of GDNF production using anterior segment explants

Eyes were enucleated 6 days or 7 months after plasmid ET in the ciliary muscle. Immediately after enucleation, eyeballs were incubated at 37 °C for 15–30 min in Dulbecco's modified Eagle's medium:F12 supplemented with antibiotics (100 U ml–1 penicillin; 100 μg ml–1 streptomycin) and 1% Fungizone antimycotic (Invitrogen). Anterior segments were carefully removed from the rest of the eye under sterile conditions (see Figure 3A and legend for details). After an additional incubation of 15 min in supplemented medium, explants were placed in a 96-well plate (200 μl per well) and incubated at 37 °C in a 5% CO2 humidified atmosphere (Figures 3b–d). After 24 h, culture medium was collected, clarified by centrifugation and the supernatant was stored at −20 °C until being analyzed by ELISA.

GDNF western blot analysis

Cell lysates and cell supernatants were collected as detailed above (see ‘In vitro assessment of GDNF synthesis and secretion’). After the addition of LDS sample buffer (NuPAGE; Invitrogen) and heating for 5 min at 100 °C, equal, equal amounts of total protein extracts were electrophoresed in a NuPAGE 4–12% Bis-Tris gel using MOPS SDS Running Buffer (Invitrogen). rrGDNF (R&D Systems, Lille, France) was used as a positive control. Proteins were then ET onto nitrocellulose membranes (Schleicher and Schuell BioScience, Dassel, Germany) using NuPAGE Transfer Buffer (Invitrogen). Membranes were blocked with 5% skimmed milk in PBS/0.5‰ Tween-20/0.5‰ Triton-X100 (PBS-TT) and incubated for 2 h at room temperature with a rabbit polyclonal anti-GDNF antibody (sc-328; Santa Cruz Biotechnology; Tebu-Bio, Le Perray en Yvelines, France) diluted 1:200 in PBS-TT containing 5% skimmed milk. Horseradish peroxidase-conjugated goat anti-rabbit immunoglobulin G (H+L) secondary antibody (Vector Laboratories, Clinisciences, Montrouge, France), diluted 1:4000 in PBS-TT containing 1% skimmed milk, was used as a secondary antibody. Bands were visualized with the Amersham ECL Plus Western Blotting Detection System (GE Healthcare Europe, Orsay, France) used according to the manufacturer’s instructions.

Neuroretinas from dystrophic and non-dystrophic rats, collected at 33 days and 78 days after birth, were snap frozen and stored at −80 °C until use. Tissues were homogenized in 300 μl lysis buffer (MOPS SDS Running Buffer; Invitrogen) supplemented with protease inhibitor cocktail (Roche Diagnostics, Meylan, France) (one tablet for 50 ml). After the addition of LDS sample buffer and heating, retinal extract proteins were analyzed by western blot as mentioned above. Blots were then dehybridized and rehybridized successively with a mouse anti-β-tubulin (D-10) (sc-5274) primary antibody (dilution 1:400) and a horseradish peroxidase-conjugated goat anti-mouse immunoglobulin G secondary antibody (sc-3697) (dilution 1:5000; both purchased from Santa Cruz Biotechnology, Tebu-bio, Le Perray en Yvelines Cedex, France). The relative band intensity for GDNF was calculated in comparison with that for β-tubulin after densitometry analysis.

GDNF and CNTF quantification by ELISA

Levels of rGDNF in media (explants supernatants, intraocular fluids) were measured using a commercially available ELISA kit for human GDNF (GDNF Emax ImmunoAssay System; Promega, Charbonnières les Bains, France). Quantification was performed according to the manufacturer’s instructions, the only difference being that a rrGDNF protein (R&D Systems) was used to generate the standard curve instead of the human GDNF standard provided in the kit. Detection threshold of the rrGDNF was estimated around 20–30 pg ml–1 in preliminary experiments. Rat CNTF levels in ocular media were quantified using a commercially available ELISA kit for rat CNTF (Duoset; R&D Systems) following the manufacturer’s instructions (detection threshold around 50 pg ml–1).

Retinal histology and measurement of retinal layer thicknesses

Eyes were harvested at PN days 51, 70–75 and 87. Enucleated eyes were immediately fixed with a mixture of 4% paraformaldehyde and 0.5% glutaraldehyde in PBS for 2 h at room temperature. They were rinsed for 2 h in PBS, dehydrated at room temperature with increasing ethanol concentrations before being incubated overnight at 4 °C with infiltration solution provided in the Leica Historesin Embedding kit (Leica, Rueil-Malmaison, France). Samples were embedded in resin (Leica) and 5-μm thick histological sections passing through the optic nerve head were prepared along the superior–inferior plane of the eye using a microtome (Leica). Sections were stuck on gelatin-coated slides, stained for 2 min with 1% toluidin blue solution and photographs were made along the whole retina (Aristoplan miscroscope, Leica).

Thicknesses of PR OSs, ONL and INL were measured manually on histological sections using the National Institutes of Health (Bethesda, MD, USA) ImageJ free software. Measurements were performed every 500 μm, at a distance of 500 to 3500 μm from the optic nerve head, in both the superior and inferior hemispheres. For each eye, three different sections were analyzed and the values obtained for each distance were averaged. Thickness profiles along the retina were generated by averaging, for each distance, the values obtained for all eyes treated similarly. For statistical analyses, the values obtained from all eyes belonging to the same experimental group were averaged out to a single value corresponding to the mean thickness along the superior or inferior retina.

Electroretinography

Dark-adapted full-field ERG responses were recorded at PN53, PN70–75 and PN87. Animals were dark-adapted over a period of 24 h and anesthetized by an intraperitoneal injection of a mixture of ketamine and xylazine (120 mg kg–1 ketamine, 10 mg kg–1 xylazine). The cornea was desensitized with a drop of Novesine (Novartis Ophthalmics, Basel, Switzerland) and the pupils were dilated with a drop of Tropicamide (Novartis Ophthalmics). Animals were placed onto a heated platform (37 °C) during the measurements to keep their body temperature constant. Gold wire ring electrodes placed on the corneas of both eyes and into the mouth served as working electrodes and a reference electrode, respectively. A stainless steel needle electrode was inserted into the tail of the animals for grounding. All these manipulations were performed under dim red light, without bringing the animal into ambient light after dark adaptation. Measurements were performed using the commercial RetiPort32 device from Roland Consult Systems (Brandenburg, Germany).

Standard ERG measurements were performed simultaneously on both eyes, with scotopic flash ERG at a light intensity of 3 cd s m–2, with the amplifier set to a frequency range of 1–200 Hz. Amplitudes of a-waves were measured from the baseline to the bottom of the a-wave trough, and b-wave amplitudes were measured from the bottom of the a-wave trough to the peak of the b-wave. Results were expressed in microvolts (μV) and the data obtained from each eye belonging to the same experimental group were averaged.

Optokinetic test

Visual acuity was measured at PN70 and PN92 using an Optomotry test apparatus (Virual Optokinetic System, Cerebral Mechanics, Lethbridge, Alberta, Canada), as already described by Prusky et al.75 This device consists of a rotating cylinder covered with a vertical sine wave grating presented in virtual three-dimensional space on four computer monitors arranged in a square. Unrestrained rats were placed on a platform in the center of the square, where they tracked the grating with reflexive head movements. Rotation direction of the grating enabled to evaluate visual acuity of left (clockwise) and right (counter-clockwise) eyes. The spatial frequency of the grating was clamped at the viewing position by repeatedly recentering the ‘cylinder’ on the head of the test subject. Spatial frequency of the grating (in c/d) was progressively increased and decreased by the software until determining a maximum threshold beyond which the optokinetic reflex was lost, thereby obtaining visual acuity.

Statistical analysis

Data analyses and statistics were performed using GraphPad Prism (GraphPad Software, San Diego, CA, USA). Numerical results were expressed as mean±s.e.m. Data were compared using the non-parametric Mann–Whitney U-test. P<0.05 was considered statistically significant.

Conclusion

This study demonstrates that ciliary muscle ET is a valuable strategy for the sustained release of bioactive neurotrophic factors for the long-term application within the eye. It could become an alternative strategy to repeated intraocular injections for the management of RP and other diseases. It is an interesting tool, not only to study the biological role and therapeutic efficacy of biological proteins, but also to evaluate their potential toxicity on the long term. Thus, our study demonstrated that the continuous release of GDNF could be useful in the treatment of RP arising from mutations in the RPE but also warns against potential retinal toxic effect with continuous release of high doses of GDNF and points out that further studies are required to determine its toxicity threshold for a safe use. It would be of special importance to assess GDNF dose as well as route and duration of GDNF delivery before clinical applications for ocular diseases. Inducible and controllable GDNF expression using gene therapy strategies, like ciliary muscle ET, could be a safe option to maintain GDNF concentration within a defined therapeutic window.

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Acknowledgements

This work was funded in part by the ANR EMERGENCE project ANR-05-EMPB-001-02 and the European project EVI-GENORET LSHG-CT-2005-512036.

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Correspondence to F Behar-Cohen.

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Touchard, E., Heiduschka, P., Berdugo, M. et al. Non-viral gene therapy for GDNF production in RCS rat: the crucial role of the plasmid dose. Gene Ther 19, 886–898 (2012). https://doi.org/10.1038/gt.2011.154

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Keywords

  • retina
  • GDNF
  • electroporation
  • toxicity
  • retinal degeneration

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