Brain Advance Access originally published online on January 17, 2006
Brain 2006 129(4):1014-1026; doi:10.1093/brain/awl015
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Synergistic action of brain-derived neurotrophic factor and lens injury promotes retinal ganglion cell survival, but leads to optic nerve dystrophy in vivo
1 Department of Pathology and Cell Biology and 2 Department of Ophthalmology, Université de Montréal, Quebec, Canada
Correspondence to: Adriana Di Polo, Department of Pathology and Cell Biology Université de Montréal 2900, Boul. Edouard-Montpetit Pavillon Roger Gaudry, Room N-535 Montreal, Quebec H3T 1J4, Canada E-mail: adriana.di.polo{at}umontreal.ca
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Summary |
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Trauma or disease in the CNS often leads to neuronal death and consequent loss of functional connections. The idea has been put forward that strategies aimed at repairing the injured CNS involve stimulation of both neuronal survival and axon regeneration. We tested this hypothesis in the adult rat retinocollicular system by combining two strategies: (i) exogenous administration of brain-derived neurotrophic factor (BDNF), a potent survival factor for damaged retinal ganglion cells (RGCs) and (ii) lens injury, which promotes robust growth of transected RGC axons. Our results demonstrate that BDNF and lens injury interact synergistically to promote neuronal survival: 71% of RGCs were alive at 2 weeks after optic nerve injury, a time when only
10% of these neurons remain without treatment. Intravitreal injection of BDNF, however, led to regeneration failure following lens injury. The effect of BDNF could not be generalized to other growth factors, as ciliary neurotrophic factor did not cause a significant reduction of lens injury-induced regeneration. Growth arrest in optic nerves treated with BDNF and lens injury correlated with the formation of hypertrophic axonal swellings in the proximal optic nerve. These swellings were filled with numerous vesicular bodies, disorganized neurofilaments and degenerating organelles. Our results demonstrate that: (i) increased neuronal survival does not necessarily lead to enhanced axon regeneration and (ii) activation of survival and growth pathways may produce axonal dystrophy similar to that found in neurodegenerative disorders including glaucoma, Alzheimer's disease and multiple sclerosis. We propose that loss of axonal integrity may limit neuronal recovery in the injured, adult CNS.
Key Words: brain-derived neurotrophic factor; lens injury; retinal ganglion cells; neuroprotection; axonal regeneration
Abbreviations: AAV = adeno-associated virus; BDNF = brain-derived neurotrophic factor; CNTF = ciliary neurotrophic factor; GFAP = glial fibrillary acidic protein; GFP = green fluorescent protein; RGCs = retinal ganglion cells
Received July 12, 2005. Revised December 14, 2005. Accepted December 22, 2005.
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Introduction |
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Injury to the brain or spinal cord of adult mammals leads to neuronal death, loss of cell–cell interactions and persistent functional deficits. Attempts to repair the injured CNS have identified two fundamental conditions that need to be fulfilled: (i) to prevent or delay the loss of neurons following injury and (ii) to overcome the failure of axons to extend towards appropriate targets (Bray et al., 1991
; Doster et al., 1991). Therefore, therapies that stimulate both neuronal viability and axon growth may prove beneficial after CNS lesion.
The rat retinocollicular system is ideal to test strategies that promote survival and regeneration of adult retinal ganglion cells (RGCs). RGCs are a well-characterized CNS neuronal population, with cell bodies located in the inner retina and axonal processes along the optic nerve that reach specific targets in the brain. This highly polarized cytoarchitecture allows the use of retrograde and anterograde neurotracers to unequivocally identify RGC soma or axons, respectively (Sapieha et al., 2003; Pernet et al., 2005). Furthermore, the accessibility and anatomy of the eye is suitable for intravitreal injection of survival factors and other reagents that can readily diffuse to reach RGCs.
Neurotrophins play important roles in the survival response of adult RGCs to injury. Among neurotrophins, brain-derived neurotrophic factor (BDNF) is the most potent survival factor for damaged RGCs (Mey and Thanos, 1993; Mansour-Robaey et al., 1994; Peinado-Ramon et al., 1996; Di Polo et al., 1998; Klocker et al., 2000; Chen and Weber, 2001). BDNF, however, lacks the ability to promote RGC axon regeneration into peripheral nerve grafts or within the optic nerve (Yip and So, 2000; Isenmann et al., 2003). In contrast, robust RGC axon regeneration has been achieved by causing a small wound puncture to the lens of adult rats (Fischer et al., 2000; Leon et al., 2000). In this paradigm, if the lens is damaged at the time of optic nerve crush, many RGCs regenerate their axons into the distal nerve. The effect of lens injury on axon growth has been proposed to be mediated by macrophage-released factors, but the precise mode of action remains unknown (Yin et al., 2003). Importantly, the molecular mechanisms that mediate the regenerative effect of lens injury are independent of BDNF, as neutralizing antibodies against this neurotrophin failed to diminish the number of RGC axons that extended after lens injury both in vivo (Leon et al., 2000) and in vitro (Lorber et al., 2002).
Here we tested the hypothesis that combination of a potent survival factor (BDNF) with a strategy that promotes axon growth (lens injury) may increase RGC viability and, consequently, enhance long-distance axon regeneration in the injured CNS. Our results demonstrate that combined BDNF and lens injury had a remarkable, synergistic effect on the survival of injured RGCs. However, intravitreal administration of BDNF at the time of lens injury completely blocked the ability of RGC axons to regrow past the lesion site. Moreover, lack of axon regeneration correlated with the formation of hypertrophic axonal swellings in the proximal optic nerve. Together, our data demonstrate an unexpected, adverse effect of combining two potent neuroprotective and regenerative strategies in the injured mammalian visual system.
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Experimental procedures |
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Intravitreal injections
Animal procedures were in accordance with the guidelines of the Canadian Council on Animal Care for the use of experimental animals. All surgeries were performed in adult female Sprague–Dawley rats (180–200 g) under general anaesthesia (2% isoflurane, 0.8 l/min). Five microlitres of BDNF recombinant protein (1 µg/µl, Regeneron Pharmaceuticals, Tarrytown, NY), ciliary neurotrophic factor (CNTF) recombinant protein (1 µg/µl, PeproTech Inc, Rocky Hill, NJ) or phosphate buffered-saline (PBS) were injected into the vitreous chamber of the left eye using a 10 µl Hamilton syringe adapted with a 32 gauge needle. Contralateral eyes or unoperated eyes served as intact controls. The needle tip was inserted into the superior hemisphere of the eye, at the level of the pars plana, at a 45° angle through the sclera into the vitreous body. This route of administration avoided retinal detachment or injury to eye structures, including the lens and the iris, which release factors that induce RGC regeneration and survival (Mansour-Robaey et al., 1994
; Leon et al., 2000). The injection was performed within 1 min and the needle was kept in place for an additional period of 2 min, after which it was gently removed. Surgical glue (Indermill, Tyco Health Care, Mansfield, MA) was used to seal the site of injection. Intraocular injections were performed immediately after optic nerve lesion as described below.
Optic nerve axotomy and analysis of neuronal survival
RGCs were backlabelled by application of FluoroGold (2%, Fluorochrome, Englewood, CO) onto both superior colliculi. One week later, the left optic nerve was transected at 0.5–1 mm from the optic nerve head avoiding damage to the ophthalmic artery. At 2 weeks post-lesion, rats were killed by intracardial perfusion with 4% paraformaldehyde. Fundus examination was performed to check the integrity of the retinal circulation after surgery, and animals showing signs of compromised blood supply were excluded from the study. The left (injured) retinas and the right (intact control) retinas were removed, fixed for an additional 30 min and flat-mounted vitreal side up on a glass slide for examination of the ganglion cell layer. RGCs backfilled with FluoroGold were counted in 12 standard retinal areas as previously described (Cheng et al., 2002; Sapieha et al., 2003). Microglia and macrophages, which may have incorporated FluoroGold after phagocytosis of dying RGCs, were excluded from our analysis of neuronal survival based on their morphology (Raibon et al., 2002) that can be easily identified in retinal whole mounts. In addition, the identity of these cells has been confirmed by immunostaining with antibodies against the microglia and macrophage markers isolectin-B4 and ED1 (Cheng et al., 2002). Data analysis and statistics were performed using the GraphPad Instat software (GraphPad Software Inc., San Diego, CA) by a one-way analysis of variance (ANOVA) test.
Optic nerve micro-crush, lens injury and analysis of axonal regeneration
The optic nerve sheath, comprising meninges and dura, was cut to expose the optic nerve. Then, a complete knot was tied around the nerve using a fine silk suture (10–0), the nerve was constricted for exactly 60 s, then the suture was carefully released as described (Sapieha et al., 2003). Thirteen days after optic nerve injury, 5 µl of 1% cholera toxin ß subunit (CTß, List Biological Laboratories, Campbell, CA) was injected into the vitreous chamber. Twenty-four hours later (2 weeks post-micro-crush lesion), animals were killed and the eyes were processed for optic nerve immunostaining as described below. Some animals received a wound puncture to the lens at the time of micro-crush lesion as described (Leon et al., 2000). Briefly, a 30 gauge needle was used to puncture the posterior plane of the lens and the wound was immediately monitored through the cornea. This procedure led to complete opacification of the lens within the first week after injury.
Axonal growth was quantified by counting CTß-positive axons that crossed a virtual line parallel to the lesion site at 250 µm, 500 µm and 1 mm past the lesion site in three to four longitudinal sections of optic nerve per animal as described (Sapieha et al., 2003). Sections were chosen in both central and peripheral regions of the optic nerve to quantify axon growth in the entire nerve. During quantification, the injury site was identified in the same optic nerve section by dark field microscopy. In addition, the location of the lesion site was routinely confirmed in the same section or in an immediately adjacent section using haematoxylin/eosin staining. The thickness of the optic nerve was measured at each point where axons were counted and this value was used to normalize the number of axons per millimetre of nerve width. Statistics were performed using the GraphPad Instat program by ANOVA.
Retina and optic nerve immunohistochemistry
Animals were perfused with 4% paraformaldehyde and the eyes and optic nerves were rapidly dissected, the cornea and the lens were removed. Tissue was embedded in optimal cutting temperature (OCT) compound (Tissue-Tek, Miles Laboratories, Elkhart, IN). Optic nerve (14 µm) or retinal (16 µm) cryosections were collected onto gelatin-coated slides. Tissue sections were pre-incubated in 5% BSA, 3% normal serum, 0.2% Triton X-100 (Sigma) in PBS for 30 min at room temperature to block non-specific binding. Primary antibody incubation was performed in 5% BSA, 3% normal serum, 0.3% Triton X-100 in PBS overnight at 4°C. Slices were then incubated with the appropriate secondary antibody for 1 h at room temperature and mounted with an anti-fade reagent (SlowFade, Molecular Probes, Eugene, OR). Primary antibodies were: goat CTß antibody (1 : 4000 dilution, List Biological Labs), monoclonal anti-rat monocytes/macrophages (2.5 µg/ml, Clone ED1, Chemicon International, Temecula, CA), monoclonal anti-glial fibrillary acidic protein (GFAP, 2 µg/ml Chemicon International), chicken anti-human BDNF (2.5 µg/ml, Promega, Woods Hollow Road Madison, WI), monoclonal anti-synaptophysin (1 : 200 dilution, NCL-SYNAP-299, Novocastra, UK), polyclonal anti-vesicle glutamate transporter 2 (VGlut2, 1 : 1000 dilution, Synaptic Systems Gmbh, Göttingen, Germany), monoclonal anti-synaptic vesicle 2 (SV2, 1 : 4000 dilution, Developmental Studies Hybridoma Bank, Department of Biological Sciences, University of Iowa, USA) or mouse RT97 antibody (1 : 200 dilution, gift from Dr John N. Wood, University College London, London, UK). Fluorescent staining was examined using a Zeiss Axioskop 2 Plus microscope (Carl Zeiss Canada, Kirkland, QC) and pictures were captured with a CCD video camera (Retiga, Qimaging, Burnaby, British Columbia) and analysed with Northern Eclipse software (Empix Imaging, Mississauga, ON).
Preparation of recombinant AAV.GFP for visualization of RGC axons
An adeno-associated virus (AAV) serotype 2 containing the green fluorescent protein (GFP) gene (AAV.GFP) was used to trace RGC axons. AAV.GFP was generated by inserting a GFP cDNA downstream of the hybrid cytomegalovirus enhancer/ chicken ß-actin promoter in the plasmid pXX-UF12, a derivative of pTR-UF5 (Zolotukhin et al., 1996). AAV.GFP vector was packaged, concentrated and titred as previously described (Hauswirth et al., 2000). The number of infectious particles/ml (i.p./ml) was determined by infectious centre assay as described (McLaughlin et al., 1988) and was 1010 ip/ml for AAV.GFP. No helper adenovirus or wild-type AAV contamination was detected in these preparations. AAV-mediated transgene expression reaches a plateau between 3 and 4 weeks after administration of the vector into the rodent eye (Bennett et al., 1997; Ali et al., 1998; Bennett et al., 2000; Cheng et al., 2002) and persists thereafter (Guy et al., 1999). The reason for delayed onset of AAV-mediated gene expression in vivo is unclear, but may arise from the need to convert single-stranded viral DNA to a double-strand prior to active transcription (Ferrari et al., 1996). Therefore, to ensure high levels of GFP protein expression in RGC axons at the time of analysis, AAV.GFP (5 µl = 5 x 107 i.p.) was injected into the vitreous chamber and subsequent surgical procedures were performed 3 weeks after virus administration (Fig. 2A). GFP labelling was directly examined on retinal flat-mounts or optic nerve sections at 2 weeks after optic nerve injury (5 weeks following virus injection).
Electron microscopy
Animals were perfused intracardially with 3.5% glutaraldehyde in Sorensen phosphate buffer (0.067 M) and optic nerves were collected. Tissue was then incubated in 2% osmium tetroxide, successively dehydrated in graded solutions of ethanol and embedded in epoxy resin (TAAB 812 resin, Marivac, Montreal, QC, Canada). Semi-thin longitudinal sections (0.5 µm-thick) were cut and stained with 1% toluidine blue mixed with 1% borax. For electron microscopy, 70-nm-thick sections were cut in the central region of the proximal optic nerve, 1 mm before the lesion site. Ultrathin sections were counterstained with 3% uranyl acetate and Reynolds's lead citrate and visualized with a Zeiss CEM902 electron microscope.
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Results |
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BDNF and lens injury interact synergistically to promote robust RGC survival in vivo
We compared the independent and combined effects of BDNF and lens injury on the survival of axotomized RGCs. For this purpose, RGCs were retrogradely labelled with FluoroGold 1 week prior to optic nerve transection. BDNF injection and/or lens injury were performed immediately after axotomy, and the density of surviving neurons was evaluated at 2 weeks post-lesion as shown in Fig. 1A.
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Experimental groups included animals with optic nerve axotomy that received: (i) a single intraocular injection of BDNF (n = 4); (ii) a single puncture to the lens (n = 4); (iii) combined BDNF injection and lens injury (n = 5) or (iv) no treatment (uninjected, n = 4). As expected, a single intraocular injection of BDNF (Fig. 1B) or lens injury (Fig. 1C) had marked, independent effects on RGC survival. Our quantitative analysis showed that a similar number of RGCs survived with BDNF (971 ± 87 RGCs/mm2, mean ± SEM) or lens injury (958 ± 66 RGCs/mm2) at 2 weeks after axotomy (Fig. 1F). These results are consistent with previous reports on the neuroprotective effect of BDNF (Mansour-Robaey et al., 1994
; Cheng et al., 2002) or lens injury (Leon et al., 2000; Fischer et al., 2001).
We then examined if lens injury combined with a single intravitreal injection of BDNF protein at the time of axotomy could potentiate RGC survival. This approach led to a dramatic increase in the density of RGCs after optic nerve injury (Fig. 1D, F). The combination of BDNF with lens injury protected 71% of the total number of RGCs (1542 ± 109 RGCs/mm2) compared with independent administration of BDNF or lens injury, which promoted only 44% neuronal survival (ANOVA, ***P < 0.001). These data indicate that BDNF and lens injury interact synergistically to enhance the survival of axotomized RGCs.
Intravitreal injection of BDNF leads to regeneration failure following lens injury
To determine if increased neuronal survival correlated with greater axon regeneration, we used the optic nerve micro-crush model. In this paradigm, although all RGC axons are axotomized, there is formation of a focal glial scar that facilitates accurate identification of the injury site and quantification of axon growth (Sapieha et al., 2003). Importantly, we previously demonstrated that axotomy and micro-crush lesion produce an identical time-course of RGC death (Sapieha et al., 2003). The experimental protocol used for these experiments is outlined in Fig. 2A.
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As expected, lens injury alone stimulated robust RGC axon growth within the distal optic nerve at 2 weeks after nerve micro-crush (Fig. 2B), consistent with previous reports (Leon et al., 2000
; Fischer et al., 2004a). Surprisingly, a single intravitreal injection of BDNF (1 µg/µl) provided at the time of lens injury completely blocked axon regeneration (Fig. 2C). Intraocular injection of BDNF by itself did not stimulate axon extension into the distal nerve (Fig. 2D), in agreement with other studies showing lack of regeneration in the presence of this neurotrophin (Yip and So, 2000; Isenmann et al., 2003). Quantitative analysis of the number of axons that extended past the lesion site confirmed that RGC axon growth was arrested by the combination of BDNF with lens injury (Fig. 2D).
To establish if the effect of BDNF could be generalized to other growth factors, we examined the effect of intravitreal injection of CNTF on lens injury-induced regeneration (Fig. 2E). In contrast to BDNF, CNTF did not cause a significant reduction of axon growth following lens injury (n = 3, Student's t-test, P = 0.6). Like BDNF, injection of CNTF alone did not promote axon regeneration within the optic nerve (Fig. 2E), which is consistent with a previous study showing lack of CNTF-induced regeneration within the optic nerve (Leon et al., 2000). We used the same source and concentration of recombinant human CNTF (rhCNTF, PreproTech, 1µg/µl) shown to effectively promote RGC survival in similar models of optic nerve injury (Mey and Thanos, 1993; Weise et al., 2000; van Adel et al., 2003; Ji et al., 2004; Park et al., 2004; van Adel et al., 2005; Zhang et al., 2005). Taken together, our data demonstrate that the effect of BDNF on lens injury-induced regeneration appears to be specific and may not be generalized to other neurotrophic factors.
BDNF does not alter the macrophage or Müller cell response induced by lens injury
To investigate the mechanisms by which BDNF may arrest lens injury-induced axon growth, we examined the effect of this neurotrophin on cellular responses associated with damage to the lens. For example, lens injury actively stimulates macrophage infiltration in the inner retina after optic nerve injury (Leon et al., 2000; Yin et al., 2003). Furthermore, macrophage-derived proteins have been shown to enhance the regenerative effect of axon growth promoting factors (Li et al., 2003). We investigated the effect of BDNF on macrophage infiltration and activation following lens injury by immunohistochemistry using ED1, an antibody that selectively labels activated rat macrophages (Fig. 3A–F). Only a small number of ED1-positive cells were visualized in the ganglion cell layer at 3 and 6 days after optic nerve lesion and lens injury (Fig. 3A, B). In contrast, substantial infiltration of active macrophages was seen in the inner retina, including the fibre layer, ganglion cell layer and inner plexiform layer, at 2 weeks post-injury (Fig. 3C). In contrast, Leon et al. (2000) reported a relatively high number of macrophages at 7 days after injury. This discrepancy may be due to differences between animal strains: we used adult female Sprague–Dawley rats, whereas Leon et al. used adult male Fisher rats. It is also possible that the 1 day difference between our observations (6 days post-axotomy) and those of our colleagues (7 days post-axotomy) contribute to this discrepancy. Importantly, a single intravitreal injection of BDNF at the time of lesion did not affect macrophage infiltration or activation at 3, 6 or 14 days after lens injury (Fig. 3D–F). We also examined the optic nerve, including the optic nerve head, for macrophage infiltration after nerve lesion (Fig. 4). At 3 and 14 days after lens injury, there was massive infiltration of activated macrophages, visualized by ED1 immunostaining, which accumulated around the site of nerve injury. At 14 days after lesion, ED1-positive cells had also infiltrated the distal optic nerve (Fig. 4C, D). The combination of lens injury with a single injection of BDNF did not alter the distribution of macrophages in injured nerves.
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Müller cells, the main glial type in the mammalian retina, play critical roles in the maintenance and function of retinal neurons. Lens injury has been shown to stimulate Müller cell activation after optic nerve injury (Leon et al., 2000
), suggesting that Müller cells may participate in the regenerative process. To investigate the effect of BDNF on Müller cell activation following lens injury, we performed retinal immunostaining using an antibody against GFAP (Fig. 3G–L). At 3 days after optic nerve and lens injuries, GFAP labelling was confined to resident astrocytes and Müller cell end-feet in the fibre layer (Fig. 3G). At 6 and 14 days post-injury, robust GFAP labelling was visualized in a large number of Müller cell radial processes spanning the retina (Fig. 3H, I). The Müller cell response to lens injury was not changed by exogenous BDNF (Fig. 3J–L). Taken together, these results indicate that BDNF does not alter macrophage or Müller cell activation elicited by optic nerve and lens injuries.
Growth arrest correlates with hypertrophic axonal swellings in the optic nerve
To determine whether the combination of BDNF with lens injury induced structural changes in RGC axons that may prevent their ability to regenerate, we used adeno-associated virus (AAV) serotype 2 carrying the GFP gene to visualize fine axon morphology in the optic nerve. We previously demonstrated that 70% of adult RGCs can be effectively transduced with AAV serotype 2 (Cheng et al., 2002; Pernet et al., 2005). Here we show that AAV-mediated GFP is effectively transported, in anterograde fashion, by RGC axons and can be used to visualize fine axon morphology in retinal flat mounts (Fig. 5A) and in longitudinal sections of the optic nerve (Fig. 5B). Using this approach, we examined the structure of RGC axon fibres in micro-crushed optic nerves after lens injury alone or combined with BDNF. Optic nerves from eyes that received only lens injury showed GFP-positive RGC axons that coursed along the dorsal–ventral plane of the nerve and had normal morphology (Fig. 6A), similar to that observed in non-treated, control nerves (Fig. 5B). In contrast, optic nerves from eyes that received combined lens injury and BDNF injection displayed axons with large, hypertrophic swellings (Fig. 6B). These structures, which reached up to 30 µm in diameter, were not confined to the terminal stump of the transected axons but appeared anywhere along the axon's trajectory between the optic nerve head and the lesion site.
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To further characterize axonal swellings within the optic nerve, we performed immunohistochemistry in longitudinal optic nerve sections using an anti-synaptophysin antibody. We chose synaptophysin, a synaptic vesicle protein (Sudhof and Jahn, 1991
), as marker because it has been previously used successfully to visualize axon enlargements (Wang et al., 1995). Synaptophysin immunoreactivity was found primarily in axon stumps near the lesion site following lens injury alone (Fig. 6C), BDNF injection alone or in untreated, injured nerves (Supplementary Fig. 1). Combined treatment with lens injury and BDNF, however, led to robust synaptophysin staining in a large number of bulbous structures or swellings scattered throughout the proximal optic nerve (Fig. 6D). These dystrophic structures corresponded to axonal swellings as shown by superimposition of synaptophysin and GFP staining (Fig. 6F). As expected, all synaptophysin-positive structures were not co-labelled with GFP because AAV.GFP infected only a subpopulation of RGCs. Visualization of synaptophysin-positive swellings at higher magnification demonstrated that these structures were an integral part of the axon shaft (Fig. 6G–I).
To establish if axonal swellings were also present within the retina, we examined intraretinal axons between the optic nerve head and RGC bodies. The morphology of intraretinal axons was visualized on flat-mounted retinas using two methods: neurotracing with AAV.GFP and immunostaining with an antibody against the 200-kDa-neurofilament subunit (RT-97), which is abundantly present in RGC axons (Mansour-Robaey et al., 1994). Our data demonstrate that the combination of BDNF and lens injury did not cause axonal swelling within the retina (Fig. 7A, B).
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Light microscopy analysis of longitudinal sections from nerves treated with lens injury and BDNF showed that axonal swellings were invariably scattered throughout the entire proximal segment of the optic nerve (Fig. 8A, B). Electron microscopy analysis revealed that, in contrast to the normal ultrastructural organization in intact axons, axonal swellings in dystrophic nerves had massive accumulation of vesicular bodies in their axolemma (Fig 8C, D). Dystrophic axons conserved an apparently intact myelin sheath (Fig. 8C) that contained a very high density of vesicles (Fig. 8D). Taken together, our data indicate that growth arrest with combined BDNF and lens injury correlates with the formation of hypertrophic axonal swellings and nerve dystrophy.
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Discussion |
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TopSummaryIntroductionExperimental proceduresResultsDiscussionSupplementary materialReferences |
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Strategies aimed at repairing the mature, injured CNS must fulfil the fundamental requirements of promoting neuronal survival and axon regeneration. We used the adult rat retinocollicular system to test the effect of combined stimulation of RGC survival and axon growth by exogenous BDNF and lens injury, respectively. Our data convincingly show that combination of BDNF and lens injury had synergistic effects on cell survival: 71% of RGCs remained alive at two weeks after optic nerve injury, a time when
90% of these neurons are lost irreversibly in the absence of treatment. The synergistic neuroprotection of BDNF and lens injury shown here confirmed that these two strategies use distinct pathways to stimulate RGC survival. The mechanisms involved in BDNF-induced neuroprotection are relatively well known. For example, BDNF has been shown to activate the extracellular signal-regulated kinases 1/2 (Erk1/2) and the phosphatidylinositol 3 kinase (Klocker et al., 2000; Cheng et al., 2002; Nakazawa et al., 2002). Furthermore, we recently demonstrated that activation of the Erk1/2 pathway by itself was sufficient to mediate robust RGC protection (Pernet et al., 2005). In contrast, the molecular mechanisms underlying lens injury-mediated survival or growth are not well understood. It has been proposed that macrophage-derived (Yin et al., 2003) and lens-derived (Lorber et al., 2002) factors may participate in increased neuronal survival and regeneration after lens injury. A comparison of gene expression profile of RGCs subjected to axotomy alone or axotomy plus lens injury revealed a list of candidate transcription and growth factors that may play a role in neuronal survival or regeneration induced by lens injury (Fischer et al., 2004b). Proteins such as SOCS-3, sphingosine kinase 1 and tissue inhibitor of metalloproteinase-1 (TIMP-1), which are selectively upregulated following lens injury, are likely to enhance cell viability (La Fleur et al., 1996; Edsall et al., 1997; Raghavendra Rao et al., 2002). Although the precise contribution of these molecules to RGC survival remains to be elucidated, it is possible that they interact synergistically with survival molecules downstream of BDNF, such as Erk1/2 (Pernet et al., 2005), to promote the robust neuronal survival produced by lens injury and exogenous BDNF.
We also report the unexpected finding that combination of BDNF and lens injury failed to enhance RGC axon regeneration. Our results show that BDNF did not alter the macrophage or glial cell response in the retina and in the optic nerve after lens injury, supporting the conclusion that these cells do not play a role in regenerative failure. The effect of BDNF on axon growth arrest appeared to be specific to this neurotrophin, as CNTF did not cause a significant reduction of lens injury-induced regeneration. CNTF has been previously shown to enhance RGC survival and axon regeneration within a peripheral nerve graft (Cui and Harvey, 2000), but uses signalling pathways different from those activated by BDNF (Peterson et al., 2000; Park et al., 2004). For example, whereas BDNF promotes RGC survival mainly via the Erk1/2 pathway (Cheng et al., 2002; Pernet et al., 2005), CNTF promotes RGC survival and regeneration via protein kinase A and Janus kinase/signal transducer and activator of transcription 3 (JAK/STAT3) (Ji et al., 2004; Park et al., 2004). Thus, it is possible that signalling components downstream of BDNF overcome those activated by lens injury leading to growth arrest. Another possibility is that BDNF sends a stop signal to RGC growth cones in the injured optic nerve. For example, BDNF has been shown to increase nitric oxide (NO) production in RGC axons in vivo (Klocker et al., 1999; Zhang et al., 2005). Moreover, combined exposure to NO and BDNF, but not either factor alone, has been shown to stabilize developing RGC growth cones inhibiting axon extension in vitro (Ernst et al., 2000; Gallo et al., 2002). Thus, rapid transport of BDNF and NO to RGC axon stumps (Caleo et al., 2000) may overly stabilize growth cones at the lesion site leading to regenerative failure following lens injury.
Importantly, we demonstrate that combination of BDNF and lens injury led to the formation of large, hypertrophic axonal swellings in the proximal injured optic nerve. Given that the structural integrity of the axon is an indispensable requisite for regeneration, it is also possible that axonal swellings caused by combined BDNF and lens injury hinder RGC axon growth. Using AAV vectors containing GFP, we demonstrate that axonal swellings were not exclusively located at the distal ends of the transected RGC axons, but they were found anywhere along the axon's trajectory between the optic nerve head and the lesion site. Swellings in dystrophic RGC axons were not only immunopositive for synaptophysin but also for other markers of synaptic vesicle proteins, such as vesicle glutamate transporter 2 (VGlut2) and synaptic vesicle 2 (SV2) (Supplementary Fig. 2),indicative of abnormal accumulation of synaptic vesicles in certain domains along the axon. Interestingly, swellings were not found in intraretinal axons between the optic nerve head and RGC cell bodies, indicating that axonal dystrophy does not occur close to RGC bodies, but only in the myelinated segment of the axon within the optic nerve.
Electron microscopy analysis confirmed that swellings were filled with a large number of vesicular bodies, disorganized neurofilaments and degenerating organelles. Thus, it is likely that regeneration failure in this situation results from disruption of axonal transport. The occlusion of the axolemma by vesicular material hinders the normal transport of cytoskeletal proteins, such as tubulin and neurofilaments, which are required for axon extension. In the optic nerve, interruption of slow axonal transport after lesion is resumed during RGC axon regeneration into peripheral nerve grafts (McKerracher et al., 1990a, b). Exogenous BDNF injected into the optic tectum of chick embryos greatly increased the density of vesicles in retinotectal synapses (Liu et al., 2003). This finding suggests that BDNF may contribute to axon occlusion by stimulating vesicle production in RGCs. Of interest is the formation of large axonal swellings in an experimental model of amyotrophic lateral sclerosis that is caused by the selective impairment of slow axonal transport, whereas fast axonal transport is preserved (Griffin et al., 1978).
Axonal swellings are characteristic of axon pathology in many neurodegenerative diseases including multiple sclerosis (Trapp et al., 1998), amyotrophic lateral sclerosis (Delisle and Carpenter, 1984), Alzheimer's disease (Brendza et al., 2003), Parkinson's disease (Galvin et al., 1999), stroke (Dewar et al., 1999) and traumatic brain injury (Cheng and Povlishock, 1988). Importantly, significant swelling of RGC axons, filled with vesicles and mitochondria, has been observed in human glaucomatous optic nerves (Quigley, 1981) and in experimental models of glaucoma (Morrison et al., 1997), a disease caused by the selective death of RGCs. Although the origin of axonal swellings in CNS disease is poorly understood, a recent study suggests that common mechanisms may underlie the development of axonal swellings in seemingly disparate CNS disorders (Mi et al., 2005). Combined BDNF with lens injury may provide a useful model to address questions related to the origin of RGC axonal swellings in degenerative diseases of the optic nerve.
In summary, our data demonstrate that combined BDNF and lens injury promote robust survival of axotomized RGCs but lead to marked optic nerve dystrophy. A better understanding of the mechanisms that lead to axonal dystrophy appears to be essential to prevent degeneration and promote nerve repair.
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Supplementary material |
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For a copy of the MSSS-88 scale, see Supplementary material at Brain online.
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Acknowledgements |
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This work was supported by grants to A.D.P. from the Canadian Institutes of Health Research, the Canadian Institute for the Blind (E.A. Baker Foundation), the Glaucoma Research Foundation and the Glaucoma Foundation. We thank Dr. Timothy Kennedy (McGill University) for comments on the manuscript, Dr. Laurent Descarries and Michel Lauzon (Université de Montréal) for assistance with electron microscopy. A.D.P. is a scholar of Fonds de recherche en santé du Québec (FRSQ).
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References |
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TopSummaryIntroductionExperimental proceduresResultsDiscussionSupplementary materialReferences |
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