Oxidative damage to mitochondria at the nodes of Ranvier precedes axon degeneration in ex vivo transected axons

Experimental Neurology 261 (2014) 127–135

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Regular Article

Oxidative damage to mitochondria at the nodes of Ranvier precedes axon degeneration in ex vivo transected axons Helena Bros a,b,c, Jason M. Millward a,b, Friedemann Paul b,c,d, Raluca Niesner e, Carmen Infante-Duarte a,b,⁎ a

Institute for Medical Immunology, Charité-Universitätsmedizin Berlin, Augustenburger Platz 1, 13353 Berlin, Germany Experimental and Clinical Research Center, a joint cooperation between the Charité-Universitätsmedizin Berlin and the Max-Delbrück Center for Molecular Medicine, Lindenberger Weg 80, 13125 Berlin, Germany c NeuroCure Clinical Research Center, Charité-Universitätsmedizin Berlin, Charitéplatz 1, 10117 Berlin, Germany d Clinical and Experimental Multiple Sclerosis Research Center, Department of Neurology, Charité-Universitätsmedizin Berlin, Charitéplatz 1, 10117 Berlin, Germany e Deutsches Rheuma-Forschungszentrum, Charitéplatz 1, 10117 Berlin, Germany b

a r t i c l e

i n f o

Article history: Received 7 January 2014 Revised 23 May 2014 Accepted 18 June 2014 Available online 25 June 2014 Keywords: Mitochondria Oxidative stress Axonal degeneration Node of Ranvier NAD+ Neurodegenerative disease

a b s t r a c t Oxidative stress and mitochondrial dysfunction appear to contribute to axon degeneration in numerous neurological disorders. However, how these two processes interact to cause axonal damage—and how this damage is initiated—remains unclear. In this study we used transected motor axons from murine peripheral roots to investigate whether oxidative stress alters mitochondrial dynamics in myelinated axons. We show that the nodes of Ranvier are the initial sites of mitochondrial damage induced by oxidative stress. There, mitochondria became depolarized, followed by alterations of the external morphology and disruption of the cristae, along with reduced mitochondrial transport. These mitochondrial changes expanded from the nodes of Ranvier bidirectionally towards both internodes and eventually affected the entire mitochondrial population in the axon. Supplementing axonal bioenergetics by applying nicotinamide adenine dinucleotide and methyl pyruvate, rendered the mitochondria at the nodes of Ranvier resistant to these oxidative stress-induced changes. Importantly, this inhibition of mitochondrial damage protected the axons from degeneration. In conclusion, we present a novel ex vivo approach for monitoring mitochondrial dynamics within axons, which proved suitable for detecting mitochondrial changes upon exogenous application of oxidative stress. Our results indicate that the nodes of Ranvier are the site of initial mitochondrial damage in peripheral axons, and suggest that dysregulation of axonal bioenergetics plays a critical role in oxidative stress-triggered mitochondrial alterations and subsequent axonal injury. These novel insights into the mechanisms underlying axon degeneration may have implications for neurological disorders with a degenerative component. © 2014 Elsevier Inc. All rights reserved.

Introduction Axon degeneration is a common feature of many neurologic disorders, affecting both the central (CNS) and the peripheral (PNS) nervous system (Coleman, 2005). In some cases, it precedes neuronal cell death (Ferri et al., 2003; Fischer et al., 2004; Gould et al., 2006; Li et al., 2001; Stokin et al., 2005), and contributes to the progression of clinical disability (Medana and Esiri, 2003; Trapp et al., 1998). Thus, understanding the mechanisms initiating axonal injury may be crucial to prevent and treat neurodegeneration. Due to their roles in producing ATP and regulating calcium homeostasis, mitochondria are particularly important to neurons. Within the

⁎ Corresponding author at: Institute for Medical Immunology, Charité-Universitätsmedizin Berlin, Augustenburger Platz 1, 13353 Berlin, Germany. Fax: +49 30 450 539906. E-mail addresses: [email protected] (H. Bros), [email protected] (J.M. Millward), [email protected] (F. Paul), [email protected] (R. Niesner), [email protected] (C. Infante-Duarte).

http://dx.doi.org/10.1016/j.expneurol.2014.06.018 0014-4886/© 2014 Elsevier Inc. All rights reserved.

axons, they are transported to the areas of highest energetic demands, such as synapses and nodes of Ranvier, where they accumulate (Hollenbeck and Saxton, 2005). A disruption of mitochondrial transport might result in energy failure and trigger axon degeneration (Perlson et al., 2010). Indeed, previous studies describe alterations of mitochondrial transport in a number of neurodegenerative diseases (Bilsland et al., 2010; Calkins et al., 2011; De Vos et al., 2007; Misko et al., 2010; Orr et al., 2008; Stokin et al., 2005). Apart from being distributed along the axon, mitochondria change their morphology according to the energetic status of the cell. Specific proteins regulate mitochondrial shape through fusion and fission of mitochondrial membranes (Chen and Chan, 2009). Under normal conditions, axonal mitochondria adopt a tubular conformation (Court and Coleman, 2012), and mitochondrial swelling seems to affect both locomotion (Kaasik et al., 2007) and function (Court and Coleman, 2012). Although the cause of mitochondrial dysfunction in axon degeneration is not clear, high levels of oxidative stress—an imbalance between production and scavenging of reactive oxygen species (ROS)—are

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present in the course of neurodegenerative disorders (Lin and Beal, 2006). In chronic CNS inflammation, activated immune cells persistently release ROS into the infiltrated tissue, which might contribute to axonal damage (Glass et al., 2010; Nikic et al., 2011). While small amounts of ROS are necessary for a number of physiological processes (Finkel, 1998; Rhee, 1999), uncontrolled generation of oxygen radicals rapidly oxidizes macromolecules and becomes toxic to organelles (Lin and Beal, 2006). However, the effects of oxidative stress in axonal mitochondria and how this might lead to axon degeneration are unresolved issues. Here, we developed a model for monitoring mitochondrial dynamics within explanted myelinated axons, to dissect the involvement of mitochondria in axonal degeneration induced by oxidative stress. Materials and methods Ethics statement Experimental procedures were approved by the regional animal study committee of Berlin (Landesamtes für Gesundheit und Soziales Berlin). Animal work was conducted in accordance with the European Communities Council Directive of 24 November 1986 (86/609/EEC). Preparation of spinal roots Ventral spinal roots were obtained from adult C57BL/6 female mice. Animals were deeply anesthetized with isoflurane prior to cervical dislocation. The dorsal side of the spine was exposed, and the vertebrae were cut laterally from rostral to caudal. The spinal cord was sectioned at the thoracic level, and gently lifted up to expose the spinal roots. The roots were cut distal to the spinal cord, before the formation of peripheral nerves. The explanted spinal cord, together with the attached spinal roots, was placed into an artificial cerebrospinal fluid solution, rigorously oxygenated with carbogen (95% O2 and 5% CO2). Under a dissecting microscope, the lumbar ventral roots were selected and separated from the spinal cord. Labeling of mitochondria and mitochondrial membrane potential To label mitochondria ex vivo, explanted ventral roots were transferred into fresh artificial cerebrospinal fluid solution, containing 100 nM of MitoTracker® orange (Life Technologies, Darmstadt, Germany) dissolved in dimethyl sulfoxide. The mixture was continuously gassed with carbogen to ensure physiologic pH and oxygenation. Ventral roots were incubated for 30 min at room temperature. To measure membrane potential, ventral roots were incubated with 10 μg/ml of JC-1 (5,5′,6,6′-tetrachloro-1,1′,3,3′tetraethylbenzimidazolylcarbocyanine iodide; Life Technologies, Darmstadt, Germany) in artificial cerebrospinal fluid at room temperature for 1 h, and washed thoroughly before imaging to minimize background. Confocal microscopy Explanted ventral roots were placed onto a glass coverslip and mounted on a microscope chamber. We used transected ventral roots that were approximately 0.8 cm long, and imaged axonal segments located in the middle of the root. All analyzed segments were at least 2 mm away from the transected ends of the root. For all imaging experiments, we used an inverted laser-scanning confocal microscope adapted for live cell imaging (LSM 710, Carl Zeiss, Jena, Germany). MitoTracker orange was excited with a DPSS laser at 561 nm. JC-1 was excited with dual illumination with argon (514 nm) and DPSS (561 nm) lasers. Mitochondria were visualized through a 100 ×/1.46 oil immersion objective (Plan-Apochromat, Carl Zeiss, Jena, Germany). Exposure time and laser power were minimized to avoid

photobleaching and phototoxicity. Experiments were conducted at room temperature. For analyzing mitochondrial morphology, serial z stacks were acquired every 0.45 μm over a total thickness of 3.6 μm, with a resolution of 1024 × 1024 pixels. Serial stacks were then merged into one single 2D file. Mitochondrial shape was analyzed with Volocity 6.0.1 (Perkin Elmer, Rodgau, Germany) and Image J 1.44 (NIH, open source). To characterize mitochondrial morphology, we calculated the length and shape factor of individual mitochondria. Shape factor (4π × [Area] / [Perimeter]2) is a measure of circularity that indicates how closely a 2D structure resembles a circle. It ranges from 0 to 1, with a value of 1 indicating a perfect circle. As the value approaches 0, it indicates an increasingly irregular or elongated shape. For mitochondrial transport, images were collected every 2 s over 1 min with a resolution of 512 × 512 pixels. To quantify mitochondrial transport, an axonal segment was selected and the number of mitochondria that changed position during the imaging time was quantified for each experimental condition. Only the mitochondria with a displacement of at least 1 μm were considered as motile. Transmission electron microscopy Transected spinal roots were fixed with 2% glutaraldehyde in 0.1 M cacodylate buffer for 16 h, at 4 °C. Samples were washed and postfixed with 1% OsO4 and 0.8% K4[Fe(CN)6] in 0.1 M cacodylate buffer for 1.5 h. Tissue was then dehydrated with graded ethanol solutions and embedded in Epon (glycid ether 100). Next, 1 μm semithin sections were cut with a microtome (RM2065, Leica, Wetzlar, Germany) and stained with Richardson methylene blue. After observing the sections under a light microscope, areas of interest were further trimmed. 70 nm ultrathin sections were stained with uranyl acetate and lead citrate, and examined by transmission electron microscopy (EM 906, Carl Zeiss, Jena, Germany). Solutions and drugs Explanted ventral roots were bathed in artificial cerebrospinal fluid containing the following (in mM): 124 NaCl, 1.25 NaH2PO4 × H2O, 10 Glucose × H2O, 1.8 MgSO4, 1.6 CaCl2 × 2H2O, 3 KCl and 26 NaHCO3. pH was adjusted to 7.4 with carbogen (95% O2 and 5% CO2). Hydrogen peroxide (H2O2), nicotinamide adenine dinucleotide (NAD+) and methyl pyruvate were purchased from Sigma-Aldrich (Schnelldorf, Germany). Statistical analysis Statistical analysis was performed using SPSS (IBM Deutschland, Ehningen, Germany) and Prism (Graph Pad, CA, USA). Comparisons between two groups were done by the Mann–Whitney U test for nonparametric data. Comparisons between more than two groups were analyzed either with one-way ANOVA followed by Tukey post hoc test, or with Kruskal–Wallis test followed by Dunn's post hoc test. Correlation analysis was done with Spearman's rank-order correlation method. p values below 0.05 were considered significant. Results Mitochondria are motile and elongated in acutely transected axons To investigate mitochondrial behavior within axons, we explanted ventral roots from the murine spinal cord and analyzed mitochondrial transport and morphology with confocal microscopy. Imaging experiments were performed within the first 3 h after axotomy, when transected axons were still electrically excitable (Tsao et al., 1999) and ATP levels had not significantly decreased (Wang et al., 2005). In our axonal explants, about 15% of mitochondria were mobile and

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mitochondrial transport did not decrease for at least 2 h after transection. Moreover, most mitochondria appeared elongated and oriented parallel to the long axis of the axon. We quantified mitochondrial morphology by means of length and shape factor—a measure of circularity that ranges from 0 to 1 (a perfect circle). On average, mitochondria length was 1.56 ± 1.02 μm, and shape factor 0.49 ± 0.20 (mean ± SD). Oxidative stress alters mitochondrial morphology in ex vivo transected axons To explore whether oxidative stress can cause mitochondrial alterations within myelinated axons, we incubated transected ventral roots with 300 μM of hydrogen peroxide and analyzed the morphology of mitochondria 2 h later. In untreated transected axons most mitochondria were elongated; however, hydrogen peroxide clearly altered the geometry of mitochondria, which became significantly shorter (length: 1.18 ± 0.73 μm) and rounder (shape factor: 0.55 ± 0.20; mean ± SD; p b 0.001) (Fig. 1A, B). We confirmed that these observations were not due to changes of mitochondrial orientation along the axon by using confocal 3D reconstructions of z stacks (Fig. 1C). Mitochondrial alterations following an oxidative insult begin at the nodes of Ranvier We observed that mitochondrial alterations induced by hydrogen peroxide did not begin uniformly along the axon, but rather they occurred initially near the nodes of Ranvier. Therefore, we sought to determine the progression of mitochondrial changes. We treated transected axons with 300 μM hydrogen peroxide, and imaged the same axonal segments before and at different time points after exposure to H2O2

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Fig. 1. Changes in mitochondrial morphology in axons exposed to oxidative stress. (A) Laser scanning confocal images of mitochondria within a myelinated motor axon. Top, mitochondria within transected axons are rod-shaped. Bottom, treatment with hydrogen peroxide alters the morphology of mitochondria, which adopt a round conformation. Scale bar: 5 μm. (B) Length and shape factor of axonal mitochondria, with and without hydrogen peroxide. Mitochondria exposed to hydrogen peroxide are significantly shorter compared with unexposed controls. 624 (control) and 465 (H2O2) mitochondria were analyzed, from 12 different axons in 4 independent experiments. (C) Representative 3D reconstruction of the morphology of mitochondria unexposed (left) and exposed (right) to H2O2. The corresponding length and shape factor of these mitochondria are represented in (B) as larger dots. 1 unit = 0.4 μm. ***p b 0.001.

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(0, 30, 60 and 120 min after adding H2O2) (Fig. 2A). At each time point, we compared the morphology of mitochondria immediately adjacent to the nodes of Ranvier (0–10 μm away) with those located distally (10–30 μm and 30–40 μm away from the node). In the mitochondria proximal to the nodes (0–10 μm), a significant reduction of length was already detected after 30 min of exposure to H2O2 (before H2O2: 1.63 ± 1.02 μm; at 30 min: 1.12 ± 0.50 μm; p = 0.047), whereas distal mitochondria were not yet affected. Significant alterations of mitochondria located distal to the nodes of Ranvier (30–40 μm) occurred only after 120 min of incubation with hydrogen peroxide (length: 0.85 ± 0.27 μm; p = 0.002; shape factor 0.61 ± 0.17; p = 0.006) (Fig. 2B). This confirmed that mitochondrial alterations began near the nodes of Ranvier and progressively extended to adjacent mitochondria. As shown in Fig. 3A, following exposure to H2O2 mitochondria were spherical at two consecutive nodes of Ranvier, but elongated in the center. Alterations of mitochondrial morphology progressed bilaterally towards both internodes (Fig. 3B). Examination of the ultrastructure with electron microscope revealed that abnormal mitochondria at the nodes of Ranvier had a complete disruption of the cristae. In contrast, the internal structure of the elongated mitochondria in the internodes was preserved (Fig. 3C). Altogether, these data demonstrate that oxidative stress-induced mitochondrial changes in transected peripheral nerves initiate locally at the nodes of Ranvier. From this point, mitochondrial alterations expand bidirectionally to adjacent mitochondria, ultimately leading to generalized mitochondrial alterations. Oxidative stress depolarizes mitochondria at the nodes of Ranvier After exposing axons to oxidative stress, we frequently observed diffusion of MitoTracker orange into the axonal cytoplasm. This diffusion typically began near the nodes of Ranvier and progressively expanded into the internodes (Fig. 4A). The diffusion of MitoTracker orange had been previously shown to occur during the depolarization of mitochondria in cultured cells (Buckman et al., 2001). Therefore, we set out to determine whether the diffusion of MitoTracker orange in our axotomy model corresponded to mitochondrial depolarization induced by oxidative stress. We measured the mitochondrial membrane potential of H2O2-treated axons with the ratiometric indicator JC-1 (5,5′,6,6′tetrachloro-1,1′,3,3′-tetraethylbenzimidazolylcarbocyanine iodide). JC-1 specifically accumulates within mitochondria and changes conformation depending on the membrane potential. As a consequence, the emitted fluorescence shifts from red (590 nm) to green (529 nm) upon depolarization, and the green/red fluorescence ratio can be used to estimate the mitochondrial polarization state. We found that mitochondria adjacent to the nodes of Ranvier (0–10 μm away from the nodes) had a significant dissipation of the membrane potential compared with distal mitochondria (located 40–70 μm away from the nodes) (node: 0.84; internode: 8.28; p b 0.001) (Fig. 4B). Similarly, MitoTracker orange diffusion was significantly most intense in areas proximal to the nodes (node: 20.09) than in the internodes (internode: 11.44; p = 0.006) (Fig. 4C). Importantly, we found a positive correlation between JC-1 fluorescence and MitoTracker orange diffusion along the axon after H2O2 exposure. Both dyes indicated the highest degree of depolarization in the areas proximal to the nodes of Ranvier, when analyzed within 10-μm intervals at various distances from the nodes (Spearman r = 0.9466; p b 0.001) (Fig. 4D). MitoTracker diffusion at the nodes of Ranvier could already be seen in areas with elongated, normal-appearing mitochondria, suggesting that mitochondrial depolarization preceded shape changes. To investigate the temporal relationship between mitochondrial depolarization and morphological alterations, we used MitoTracker orange, which in our model was shown to be more accurate than JC-1 for evaluating mitochondrial morphology. This was due to the poor penetration and very low fluorescence intensity of JC-1 in the transected axons (data not shown). We monitored the MitoTracker orange diffusion and mitochondrial shape factor of the same axonal regions at different time

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minutes with H2O2 Fig. 2. Mitochondrial alterations begin proximal to the nodes of Ranvier and progress to the internodes. (A) Top, Mitochondria within a single myelinated axon, imaged before applying hydrogen peroxide (300 μM). The rest of the images below show the same mitochondria 30, 60 and 120 min after applying H2O2. Nodes of Ranvier are situated at the left side (*). Scale bar: 5 μm. (B) Length (upper row) and shape factor (lower row) of mitochondria exposed to oxidative stress. Mitochondria are grouped into 3 axonal segments, according to their proximity to the node of Ranvier: 0–10 μm (left column), 10–30 μm (middle column) and 30–40 μm (right column) away from the node. Earliest mitochondrial changes occur within 30 min near the nodes of Ranvier. Distal mitochondria do not undergo significant shape changes until 120 min with hydrogen peroxide. 27–72 mitochondria from 4 different axons were analyzed per group (3 independent experiments). Error bars show 95% confidence intervals. *p b 0.05, **p b 0.01, ***p b 0.001.

points after exposure to H2O2 (Fig. 4E). This analysis was done in the internodal areas (10–40 μm away from the nodes), because both morphological changes of mitochondria and MitoTracker diffusion at the nodes of Ranvier occur within the first few minutes after H2O2 exposure. We found a significant diffusion of MitoTracker into the cytosol 30 min after adding H2O2 (before H2O2: 12.31; at 30 min: 22.96; p b 0.001), but mitochondria were still elongated (before H2O2: 0.46; at 30 min: 0.51; p = 0.656). Remarkably, significant changes of mitochondrial geometry were first detected only after 60 min incubation with H2O2 (at 30 min: 0.51, at 60 min: 0.70; p = 0.007). Thus, mitochondrial depolarization and therefore dysregulation of mitochondrial bioenergetics appear to be an early event, which precedes alterations of external morphology.

Oxidative stress reduces mitochondrial transport In our nerve axotomy model, mitochondria are mobile after transecting the axons, and motility remains stable at least for 2 h (data not shown). Considering that oxidative stress dramatically altered mitochondrial morphology, we examined whether it would also interfere with axonal transport. We incubated explanted peripheral roots with hydrogen peroxide for 1 h and analyzed mitochondrial transport at the nodes of Ranvier and at the internodes with time-lapse confocal microscopy. To quantify mitochondrial transport, we selected 10-μmlong axonal segments at the nodes of Ranvier (0–10 μm away from the nodes) and at the internodes (30–40 μm away from the nodes) and counted the number of mitochondria that moved over 1 min.

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treated: 1.1 mitochondria moving; p = 0.063) (Fig. 5A). In untreated axons, mitochondrial transport at the nodes of Ranvier was not significantly different from the transport at the internodes (nodes: 2.3 mitochondria moving; internodes: 1.6 mitochondria moving; p = 0.366) (data not shown). To investigate whether altered mitochondrial motility induced by H2O2 treatment was associated with altered mitochondrial morphology, we then quantified the length and shape factor of moving vs. stationary mitochondria at the nodes of Ranvier. We found that stationary mitochondria were significantly more circular than motile mitochondria (shape factor of motile: 0.66 ± 0.17; stationary: 0.79 ± 0.19; p b 0.001). However, we did not find significant differences in mitochondrial length between the two groups (length of motile: 1.62 ± 0.83 μm, stationary: 1.91 ± 1.37 μm; p = 0.746) (Fig. 5B). These data indicate that oxidative stress interferes with mitochondrial transport at the nodes of Ranvier, which is particularly decreased in rounded mitochondria. Yet, from these data we cannot conclude that mitochondrial swellings were the exclusive cause of the decreased mobility, since other alterations of the axonal transport machinery (e.g. microtubule disruption) may also contribute to this effect.

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Fig. 3. Earliest mitochondrial changes affect exclusively the nodes of Ranvier and progress on both directions. (A) The schematic drawing represents a myelinated axon with two adjacent nodes of Ranvier. Dashed squares delineate the axonal areas that are imaged below (a, b, c and d). Fluorescence pictures show mitochondria within an axon exposed to hydrogen peroxide. Mitochondria from both nodes of Ranvier (a and d) are globular, whereas mitochondria located distal to the nodes are elongated (b and c). Nodes of Ranvier are signaled with either one or two asterisks. Scale bar: 10 μm. (B) Mitochondria within a node of Ranvier (*), without (upper panel) and with (lower panel) hydrogen peroxide. After 120 min of incubation with hydrogen peroxide, rounded mitochondria are seen all along the axon on both sides of the node of Ranvier, showing that mitochondrial changes progress on a bidirectional manner. Scale bar: 10 μm. (C) Electron micrographs of nodal (left) and internodal (right) mitochondria exposed to oxidative stress. Internodal mitochondria are elongated, have intact cristae and intact double membranes. Nodal mitochondria are globular and characterized by dilatation and complete disruption of the cristae. The arrows point at the myelin sheath. Arrowheads delineate the node of Ranvier, without myelin. 12930 X. Scale bar: 0.5 μm.

Mitochondria were classified as motile when their displacement was ≥1 μm and stationary when it was b 1 μm. Analysis of the overall transport (both anterograde and retrograde) at the nodes of Ranvier revealed that mitochondrial transport was significantly reduced in H2O2treated axons compared with untreated controls (untreated: 2.3 mitochondria moving; treated: 1.1 mitochondria moving; p = 0.040). However, at the internodes the transport between the two groups did not reach statistical significance (untreated: 1.6 mitochondria moving;

Mitochondria need both nicotinamide adenine dinucleotide (NAD+) and pyruvate to generate ATP via the oxidative phosphorylation pathway. When applied exogenously, NAD+ and methyl pyruvate—a membrane-permeant form of pyruvate—were shown to delay axon degeneration in both transected sciatic nerves (Park et al., 2013) and cultured dorsal root ganglia explants (Wang et al., 2005). Given their protective effects, we examined whether NAD+ and methyl pyruvate would also prevent mitochondrial alterations induced by oxidative stress. We applied a mixture of 1 mM NAD+ and 20 mM methyl pyruvate, together with 300 μM hydrogen peroxide, in the axonal explants. After 2 h of incubation, mitochondria exposed only to hydrogen peroxide were clearly altered, particularly at the nodes of Ranvier. However, addition of NAD+ and methyl pyruvate strikingly preserved mitochondria morphology (Fig. 6A). Mitochondria exposed to hydrogen peroxide were significantly shorter (length: 0.83 ± 0.58 μm) and rounder (shape factor 0.70 ± 0.15) than mitochondria exposed also to NAD+ and methyl pyruvate (length: 1.21 ± 0.90 μm; p = 0.001; shape factor: 0.62 ± 0.17; p b 0.001) (Fig. 6B). NAD+ and methyl pyruvate protected mitochondria equally within the two heminodes (i.e. left side and right side of the node of Ranvier), since at both locations mitochondria presented a similar morphology (Fig. 6B). 72% of axons contained spherical mitochondria when incubated only with H2O2, whereas only 17% of axons contained abnormal mitochondria when co-incubated with NAD+ and methyl pyruvate (p = 0.029) (Fig. 6C). In agreement with this, NAD+ and methyl pyruvate prevented mitochondrial depolarization following H2O2 exposure (MitoTracker diffusion after H2O2: 10.75 ±6.85; diffusion after H2O2 and NAD-pyruvate: 4.48 ± 2.82, arbitrary units; p b 0.001) (Fig. 6D). Next, we investigated whether preventing oxidative damage in mitochondria would also protect axons from degeneration. For this purpose, we extended the incubation time with hydrogen peroxide to 8 h. By this time point, several axons had developed swellings and spheroids (Fig. 7A), which are common elements of axon degeneration resulting from accumulation of organelles due to focal blockages of axonal transport (Coleman, 2005). Additionally, most mitochondria were not visible anymore because the specific staining had diffused out of the organelles due to dissipation of their membrane potential (Fig. 7A). In contrast, when NAD+ and methyl pyruvate were included in the extracellular solution together with hydrogen peroxide, mitochondria retained fluorescence after 8 h in culture, suggesting that even at this late time point they were actively respiring and energized. Most of them were elongated, even proximal to the nodes of Ranvier. Importantly, we did not

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minutes with H2O2 Fig. 4. Oxidative stress depolarizes mitochondria at the nodes of Ranvier. (A) Mitochondrial depolarization within axons treated with hydrogen peroxide. Upper panel: JC-1 staining. Upon depolarization, emitted fluorescence shifts from red (590 nm) to green (529 nm). Mitochondria near the node of Ranvier are depolarized compared with distal mitochondria. Lower panel: axonal segment labeled with MitoTracker orange, which diffuses out of the mitochondria upon depolarization. MitoTracker diffusion is most intense near the node of Ranvier. In both panels, the nodes of Ranvier are marked with an asterisk. Scale bar: 5 μm. (B) JC-1 fluorescence ratio in H2O2-treated axons. Each dot represents the ratio of green to red fluorescence in one individual mitochondrion. Nodal mitochondria (0–10 μm away from the nodes of Ranvier) are significantly depolarized compared with internodal mitochondria (40–70 μm away from the nodes). 160 (nodal) and 84 (internodal) mitochondria were analyzed, from 3 independent experiments. (C) MitoTracker orange diffusion into the cytoplasm in H2O2treated axons. Each dot represents the mean fluorescence intensity of MitoTracker orange in the cytosol of one axonal segment. The fluorescence contributed by the mitochondria themselves was excluded from the analysis to obtain values exclusively for the cytosol. The cytosolic diffusion of MitoTracker orange is significantly higher near the nodes of Ranvier (0–10 μm away from the nodes) than in the internodes (40–70 μm away from the nodes). 12 axons were analyzed per group, from 4 independent experiments. (D) Correlation between JC-1 (green/red depolarization ratio) and MitoTracker orange diffusion in H2O2-treated axons. For quantification, 220-μm-long axonal segments labeled with either JC-1 or MitoTracker orange were divided into 10-μm sections according to their distance from the node of Ranvier. For each 10-μm section, JC-1 and MitoTracker fluorescences from 9 different axons were averaged and plotted as one single dot. The color scale shows the position of the dots relative to the nodes of Ranvier. Data are pooled from 3 independent experiments. (E) Mitochondrial depolarization (left; measured with MitoTracker orange diffusion) and shape factor (right) of internodal mitochondria (10–40 μm away from the nodes) at different time points after applying H2O2. Each line represents the changes in the diffusion and shape factor of one single mitochondrion over time. The black line depicts the mean values. To calculate MitoTracker orange diffusion associated with each mitochondrion, the immediate cytoplasmic area around the mitochondrion was selected and the mean fluorescence intensity was calculated (excluding the contribution of the mitochondrion itself). MitoTracker orange diffusion initiates before any detectable change in mitochondrial morphology. 18 mitochondria were analyzed, from 8 axons pooled from 3 independent experiments. a.u.: arbitrary units. *p b 0.05, **p b 0.01, ***p b 0.001.

detect axonal spheroids or any morphological signs indicative of axon degeneration (Fig. 7B). Thus, NAD+ and pyruvate not only protected mitochondria from oxidative damage, but also prevented subsequent degeneration of axons. Discussion Neurons are polarized cells with a unique morphology, in which dendrites and the cell body are separated from the axon and the synaptic terminal (Rasband, 2010). Due to this large distance between cellular compartments, axonal and neuronal integrity are highly dependent on

effective transport and function of mitochondria all along the axons (Perlson et al., 2010). There is evidence that oxidative stress plays roles in neuronal and axonal death, and it has been linked to mitochondrial damage (Lin and Beal, 2006; Mattson and Magnus, 2006; Nikic et al., 2011). However, whether oxidative stress is a primary cause or a downstream consequence of mitochondrial malfunctioning is still a matter of debate. In this study, we investigated the effects of oxidative stress in axonal mitochondria using an ex vivo model of peripheral axon degeneration. This is important because in neuroinflammatory diseases of the nervous system, such as multiple sclerosis, activated microglia produce high

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amounts of reactive oxygen and nitrogen species (Glass et al., 2010), which might directly trigger axon degeneration (Nikic et al., 2011; Tegenge et al., 2013). We showed here that this model is suitable and accurate for real time visualization and quantification of mitochondrial dynamics under experimental conditions. Using this model, we showed that oxidative stress alters mitochondrial shape, motility and polarization within myelinated axons. Importantly, we demonstrated for the first time that oxidative stress-induced mitochondrial changes initiate at the nodes of Ranvier, and from there extend progressively throughout the whole axon. Finally, we showed that energy supplementation with NAD+ and pyruvate protects mitochondria from oxidative damage and prevents axon degeneration. First, we investigated the suitability of explanted murine ventral roots to monitor mitochondrial transport and morphology ex vivo. It has been reported that during the first 3 h after axon transection, axons maintain their electrical excitability (Tsao et al., 1999) and ATP levels do not significantly decrease (Wang et al., 2005). Accordingly, the percentage of motile mitochondria in our explants within the first hours after axotomy is consistent with previous data on mitochondrial transport in entire neurons (Misgeld et al., 2007). Moreover, during this time period mitochondria maintain an elongated morphology, which is similar to that observed in living mice (Misgeld et al., 2007). Mitochondrial dynamics in explanted ventral roots resemble those of the in vivo situation, and can therefore be used as a simplified model system for monitoring mitochondrial behavior.

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left side of the node of Ranvier

C

nodes with spherical mitochondria

D

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100 80 60 40 20

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Fig. 5. Mitochondrial transport is reduced at the nodes of Ranvier. (A) Overall motility (anterograde and retrograde) of mitochondria at the nodes of Ranvier (left) and at the internodes (right) in untreated vs. H2O2-treated axons. The scheme of an axonal segment illustrates the nodal and internodal areas from where mitochondria were analyzed. The node of Ranvier is marked with an asterisk. The graphs show the number of mitochondria that were moving within a 10-μm axonal segment during 1 min. After 1 h of treatment, mitochondrial transport was significantly reduced at the nodes of Ranvier, but not at the internodes. Horizontal lines represent the mean values. 24 axons were analyzed per group, from 4 independent experiments. (B) Shape factor (left) and length (right) of moving and stationary mitochondria at the nodes of Ranvier, after 1 h of treatment with H2O2. Stationary mitochondria were significantly more circular than motile mitochondria (38 moving mitochondria and 138 stationary mitochondria were analyzed). Mitochondrial length did not differ significantly between the two groups (36 moving mitochondria and 125 stationary mitochondria were analyzed). 29 axons were analyzed, from 4 independent experiments. *p b 0.05; ***p b 0.001; n.s.: not significant.

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10 8 6 4 2

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no. of moving mitochondria

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mitochondrial membrane potential

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25 20 15 10 5 H2O2

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Fig. 6. Local application of NAD+ and methyl pyruvate prevents oxidative damage in mitochondria. (A) Top, axon incubated with hydrogen peroxide. It contains spherical mitochondria and diffuse MitoTracker staining. Bottom, axon incubated with NAD+ and methyl pyruvate (pyr), in addition to hydrogen peroxide. Mitochondria are elongated and retain their specific staining. The nodes of Ranvier are situated in the middle (*). Scale bar: 10 μm. (B) Length and shape factor of mitochondria located near the nodes of Ranvier of axons incubated with H2O2, with and without NAD+ and methyl pyruvate. NAD+ and methyl pyruvate-treated mitochondria had significantly longer length and lower shape factor. In black, mitochondria located at the left side of the nodes of Ranvier; in gray, mitochondria located at the right side. There were no significant differences between the two heminodes. 59–113 mitochondria were analyzed per group, from 9 to 12 axons, pooled from 3 independent experiments. (C) Percentage of nodes of Ranvier that have globular mitochondria in spinal roots incubated with H2O2. When treated with NAD+ and methyl pyruvate, significantly less nodes of Ranvier showed globular mitochondria. (D) Diffusion of MitoTracker orange into the cytoplasm in H2O2-treated axons, with and without NAD+ and methyl pyruvate. For the analyses, an axonal area of 30– 50 μm2 was selected, excluding the mitochondria, and the mean fluorescence intensity of that area was calculated for each axon. Diffusion is lower in NAD+ and methyl pyruvate-treated axons, indicating that mitochondria within these axons retained their membrane potential. The diffusion of MitoTracker was analyzed in 21–22 axons, from 3 independent experiments. a.u.: arbitrary units. *p b 0.05; **p b 0.01; ***p b 0.001; n.s.: not significant.

Using explanted ventral roots, we demonstrated that the application of an oxidative insult led to changes in mitochondrial shape. Mitochondria lost their elongated, rod-like form and became significantly shorter and rounder (Fig. 1). Previous studies had reported similar alterations of

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A

H2O2

B

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Fig. 7. NAD+ and methyl pyruvate protect oxidative stress-treated axons from degeneration. (A) Transected axons incubated with hydrogen peroxide for 8 h. Axonal mitochondria are not visible due to diffusion of the staining out of the organelles. Some small caliber axons present axonal spheroids (arrowheads). (B) Axons incubated with hydrogen peroxide, NAD+ and methyl pyruvate (pyr), for 8 h. Most mitochondria are still labeled and are elongated. Morphological signs of axonal degeneration are not visible. Scale bar: 20 μm.

mitochondria in experimental models of multiple sclerosis and amyotrophic lateral sclerosis (Jaarsma et al., 2001; Nikic et al., 2011; Vande Velde et al., 2011), disorders characterized by the presence of axonal degeneration. As in our data, these studies showed that mitochondrial changes preceded axon degeneration, suggesting that mitochondrial dysfunction is an early event that could be targeted for preventing axonal damage. In addition, we estimated changes of mitochondrial membrane potential after H2O2 treatment based on the diffusion of MitoTracker orange into the cytosol. Intensity of MitoTracker orange staining has long been used to assess mitochondrial transmembrane potential in various cellular models (Bajic et al., 2013; Czarna et al., 2010; Ikegami and Koike, 2003; Kessel, 2014; Macho et al., 1996; Matylevitch et al., 1998). However, the results obtained with this dye used as a potentiometric probe must be interpreted with caution (Buckman et al., 2001; Ferlini et al., 1998; Scorrano et al., 1999). Here, we showed that MitoTracker orange diffused into the cytosol after axons were exposed to H2O2. MitoTracker diffusion was positively correlated with mitochondrial depolarization assessed by the potentiometric dye JC-1. These data are in agreement with previous reports of MitoTracker orange redistribution into the cytosol upon exposure to a depolarizing stimulus (Buckman et al., 2001), and suggest that MitoTracker orange diffusion can be used in explanted roots to estimate changes in mitochondrial potential. Importantly, we showed that both mitochondrial swelling and depolarization induced by H2O2 began proximal to the nodes of Ranvier. Remarkably, MitoTracker diffusion at the nodes preceded any detectable change in mitochondrial morphology (Fig. 4E). The enhanced susceptibility of nodal mitochondria to oxidative stress might be due in part to the lack of myelin, which may act as a barrier limiting the penetration of ROS into the axon. However, we cannot exclude an active contribution of nodal ion channels to this increased sensitivity, as blocking sodium channels has been recently shown to protect axons from degeneration after inducing mitochondrial dysfunction (Persson et al., 2013). Previous in vitro investigations showed that, following an oxidative insult, Na+ and Ca2+ increase locally within the axon, and this constitutes a preceding step towards the formation of axonal spheroids and axon fragmentation (Barsukova et al., 2012). Since the nodes of Ranvier

have the highest density of voltage-gated sodium channels in the axon (Poliak and Peles, 2003), oxidative stress-induced elevations of Na+ might initially occur there. Excess Na+ might then be locally exchanged with Ca2 + by the Na+/Ca2 + exchanger. Accordingly, Zhang and colleagues showed focal increases of Ca2+ at the nodes of Ranvier in response to nerve activity (Zhang et al., 2010). Together with oxidative stress, focal Ca2+ elevations at the nodes of Ranvier might create a situation highly favorable for the activation of the mitochondrial permeability transition pore (mPTP). mPTP activation promotes dissipation of mitochondrial membrane potential (Bernardi et al., 2006), mitochondrial swellings and unfolding of the cristae (Rasola and Bernardi, 2007). Alterations following mPTP activation closely resemble the changes of mitochondrial polarization, internal and external morphology that we observed at the nodes of Ranvier. Thus, our data point to an early involvement of mPTP at the nodes of Ranvier following an oxidative insult, which could directly promote axon degeneration (Barrientos et al., 2011). Furthermore, with time-lapse confocal imaging we found that 60 min of treatment with H2O2 significantly reduced mitochondrial transport at the nodes of Ranvier. Swollen mitochondria were mainly stationary (Fig. 5), suggesting that mitochondrial swellings might be associated with transport deficits. Depolarized mitochondria fail to produce ATP (Perry et al., 2011), which is used by the motor proteins to translocate mitochondria along microtubules. Therefore, local ATP depletion in swollen, depolarized mitochondria at the nodes of Ranvier could explain the inhibited motility frequently observed in circularized mitochondria. It is also plausible that other axonal alterations induced by oxidative stress (e.g. microtubule disruption and/or local ionic imbalances) contribute to the reduction of mitochondrial transport. Further experiments are necessary to investigate the exact mechanism of mitochondrial arrest at the nodes of Ranvier after oxidative stress. The nodes of Ranvier have increased energetic demands due to their high density of ion channels. Upon ATP depletion, ATP-dependent pumps can no longer maintain the ionic homeostasis, which ultimately leads to toxic intraaxonal Ca2+ concentrations (Stys, 2005; Waxman, 2006). We therefore hypothesized that restoring axonal bioenergetics would help reduce the ionic imbalance at the nodes of Ranvier, and that this would prevent mitochondrial alterations and the onset of axon degeneration. Previously, Wang and colleagues had shown that ATP levels progressively decrease in injured nerves (Wang et al., 2005), and that administering NAD+ and pyruvate prevented the decrease in ATP and protected axons from degeneration (Park et al., 2013; Wang et al., 2005). When we applied these mitochondrial substrates in our transected axons, together with an oxidative insult, not only were mitochondrial alterations significantly prevented (Fig. 6), but axons were strikingly protected from degeneration (Fig. 7), as indicated by the absence of morphological signs of degeneration. Thus, these data suggest that oxidative damage to mitochondria might be a consequence of energetic deficits in the axon. However, from these experiments we cannot exclude the possibility that NAD+ and pyruvate may have protected mitochondria by other mechanisms besides increasing the energetic supply, such as scavenging ROS or directly inhibiting mPTP (Kerr et al., 1999). It would be thus interesting to elucidate in future experiments the exact role of axonal bioenergetic supply in mitochondrial damage and subsequent axon degeneration induced by oxidative stress. Conclusions Altogether, our results demonstrate that transected motor axons from murine peripheral roots represent a suitable ex vivo model system for studying mitochondrial changes upon exogenous application of oxidative stress. This model revealed that oxidative stress leads to mitochondrial dysfunction in PNS myelinated axons, which precedes axonal damage. We propose that the unique ionic specializations and high energy requirements at the nodes of Ranvier make them a key site of

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early mitochondrial damage in peripheral axons. In agreement with this, we demonstrated that enhancement of axonal bioenergetics had a positive effect on interrupting the degenerative cascade by protecting mitochondria from oxidative damage. Conflict of interest The authors declare that they have no conflict of interest. Acknowledgments This project was supported by a fellowship from La Caixa and the Deutscher Akademischer Austauschdienst to H. Bros (A/11/94367) and by the German Research Council (DFG Exc 257 to F. Paul). We thank Natalie Asselborn for technical assistance and Prof. Markus Schülke for valuable suggestions. We also thank the JIMI network for infrastructural imaging support. References Bajic, A., Spasic, M., Andjus, P.R., Savic, D., Parabucki, A., Nikolic-Kokic, A., Spasojevic, I., 2013. Fluctuating vs. continuous exposure to H(2)O(2): the effects on mitochondrial membrane potential, intracellular calcium, and NF-kappaB in astroglia. PLoS ONE 8, e76383. 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