AMP-Activated Protein Kinase Mediates Activity-Dependent Axon Branching by Recruiting Mitochondria to Axon Kentaro Tao, Norio Matsuki, Ryuta Koyama Laboratory of Chemical Pharmacology, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Tokyo 113-0033, Japan Received 29 June 2013; revised 12 September 2013; accepted 7 November 2013

ABSTRACT: During development, axons are guided to their target areas and provide local branching. Spatiotemporal regulation of axon branching is crucial for the establishment of functional connections between appropriate pre- and postsynaptic neurons. Common understanding has been that neuronal activity contributes to the proper axon branching; however, intracellular mechanisms that underlie activity-dependent axon branching remain elusive. Here, we show, using primary cultures of the dentate granule cells, that neuronal depolarization-induced rebalance of mitochondrial motility between anterograde versus retrograde transport underlies the proper formation of axonal branches. We found that the depolarization-induced branch formation was blocked by the uncoupler p-trifluoromethoxyphenylhydrazone, which suggests that mitochondria-derived ATP mediates the observed phenomena. Real-time analysis of mitochondrial movement defined the molecular

INTRODUCTION The temporal and spatial regulation of axon branching is the basis for the establishment of functional neural Correspondence to: R. Koyama ([email protected]). Contract grant sponsor: Grant-in-Aid for Young Scientists (B), The Ministry of Education, Culture, Sports, Science and Technology; contract grant number: 21790059. Contract grant sponsor: The Takeda Science Foundation. Contract grant sponsor: Grant-in-Aid for JSPS Fellows, Japan Society for the Promotion of Science; contract grant number: 22-10641. Ó 2013 Wiley Periodicals, Inc. Published online 12 November 2013 in Wiley Online Library (wileyonlinelibrary.com). DOI 10.1002/dneu.22149

mechanisms by showing that the pharmacological activation of AMP-activated protein kinase (AMPK) after depolarization increased anterograde transport of mitochondria into axons. Simultaneous imaging of axonal morphology and mitochondrial distribution revealed that mitochondrial localization preceded the emergence of axonal branches. Moreover, the higher probability of mitochondrial localization was correlated with the longer lifetime of axon branches. We qualitatively confirmed that neuronal ATP levels decreased immediately after depolarization and found that the phosphorylated form of AMPK was increased. Thus, this study identifies a novel role for AMPK in the transport of axonal mitochondria that underlie the neuronal activity-dependent formation of axon branches. VC 2013 Wiley Periodicals, Inc. Develop Neurobiol 74: 557–573, 2014

Keywords: granule cell; mitochondria; axonal transport; axon branching; AMP-activated protein kinase

circuits. To form a final morphology, axons undergo the neuronal activity-dependent multiple processes such as the region-specific competition between axon branches in the retinogeniculate projections, that is, eye-specific segregation (Chen and Regehr, 2000; Jaubert-Miazza et al., 2005; Huberman et al., 2008), and the layer-specific branching of thalamocortical axons (Herrmann and Shatz, 1995; Uesaka et al., 2005, 2007). Deficits in the regulation of activitydependent axon branching reorganize neural circuits, increasing the risk of neurological disorders such as epilepsy (Shetty, 2002; Bausch and McNamara, 2004). Several extracellular factors, for example, the brain-derived neurotrophic factor (Cohen-Cory and 557

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Fraser, 1995; Koyama et al., 2004; Singh et al., 2008), have been suggested to mediate activity-dependent axon branching under normal and pathological conditions; however, intracellular mechanisms that translate activity to axon branching remain open questions. Mitochondria are highly dynamic organelles that are transported in anterograde and retrograde directions along the axons (Hirokawa and Takemura, 2005; Hollenbeck and Saxton, 2005). They are localized to subcellular regions with high metabolic demands, such as the axon initial segment, nodes of Ranvier, growth cones, and synapses (Li et al., 2004; Chang et al., 2006; Kang et al., 2008; Kiryu-Seo et al., 2010). In addition, to satisfy rapid changes in local metabolic demands, the localization of the mitochondria is strictly controlled on a fine time scale (MacAskill et al., 2009; Wang and Schwarz, 2009). A recent study demonstrated that mitochondrial immobilization leads to axon branching (Courchet et al., 2013), but it is still unknown whether the change in mitochondrial dynamics is involved in this process. In this study, we investigated whether and how mitochondrial localization and axon branching are cell autonomously regulated after depolarization. Furthermore, we investigated the involvement of AMP-activated protein kinase (AMPK) in the regulation of mitochondrial distribution. AMPK is activated when the intracellular levels of ATP decrease, and AMPK regulates signal transduction via phosphorylation as well as transcription and translation to maintain homeostasis of the intracellular metabolic state (Ramamurthy and Ronnett, 2006). It has recently been proposed that AMPK contributes to the formation of neuronal polarity (Williams and Brenman, 2008; Amato et al., 2011); however, its function in neuronal morphogenesis is largely unknown. To examine the role of mitochondria in neuronal activity-dependent axonal morphogenesis, we performed real-time simultaneous imaging of mitochondrial transportation and axon branching in primary cultures of dentate granule cells (GCs), whose axons, that is, the mossy fibers, are known to exhibit activity-dependent robust branching in the epileptic brain (Nadler, 2003; Koyama and Ikegaya, 2004; Sutula and Dudek, 2007). Our study identified a novel role for AMPK in which neuronal depolarization induced the activation of AMPK and increased the anterograde transport of mitochondria, which resulted in robust axon branching.

MATERIALS AND METHODS Animals Sprague-Dawley rat pups (SLC; Shizuoka, Japan) were maintained under controlled temperature and light Developmental Neurobiology

schedules with unlimited food and water. All experimental procedures conformed to the National Institutes of Health Guide for the Care and Use of Laboratory Animals and guidelines provided by the University of Tokyo, Tokyo.

DNA Constructs and Pharmacological Agents pAcGFP-Mem and pDsRed2-Mito were purchased from Takara Bio (Shiga, Japan). The pharmacological agents were used at the following concentrations: carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP), 0.1 or 1 lM (SigmaAldrich; St. Louis, MO); Compound C (CC), 10 lM (Calbiochem; San Diego, CA); 5-aminoimidazole-4-carboxamide ribonucleoside (AICAR), 2.5 mM (Calbiochem); and kainic acid, 2 lg/lL (Fujisawa Pharmaceutical; Osaka, Japan).

Primary Culture of GCs and Gene Transfection Primary GC cultures were prepared from P6 SpragueDawley rats as described previously (Yamada et al., 2008). To culture only immature GCs, the dentate hilar region, in which neonatal-generated GCs extensively proliferate and migrate during the postnatal period, was isolated and used. Briefly, the posterior part of the brain was sliced into 300lm-thick transverse slices in aerated, ice-cold Gey’s balanced salt solution that contained 25 mM glucose. The hilar region was carefully dissected from the slice using a microscalpel (Microfeather P-730; Feather, Osaka, Japan) under microscopic control. Following trituration and treatment with 0.25% trypsin (Nakarai Tesque; Kyoto, Japan) and 0.01% DNase I (Sigma-Aldrich), the cells were plated at a density of 5 3 103 cells/cm2 on poly-L-lysine-coated glass cover slips in Neurobasal media (Invitrogen; Gaithersburg, MD) supplemented with 0.5 mM L-glutamine and 2% B-27 (Invitrogen). At 1 day in vitro (DIV), the cultured GCs were transfected with pAcGFP-Mem and pDsRed-Mito by lipofection using Lipofectamine 2000 (Invitrogen) according to the manufacturer’s instructions. After 1 h, the medium was completely replaced. All pharmacological agents used in this study were washed out by replacing the preincubated medium every 10 min for three times.

Real-Time Imaging GCs that did not come into direct contact with other GCs and glial cells were carefully selected using the differential interference contrast images and observed. Data were acquired using an MRC-1024 confocal microscope (BioRad; Richmond, CA) with a 203 objective (numerical aperture, 0.75; Nikon; Tokyo, Japan). For real-time imaging, the cover slips were placed in a sterilized chamber filled with 3 mL Dulbecco’s modified Eagle medium (DMEM) (Nakarai Tesque) and incubated with 5% CO2 at 37 C. Images were captured every 5 s for 20 min. For high potassium ion (K1) stimulation, 60 lL DMEM that contained 2.2 3 103 mM KCl was added to the chamber 10 min after beginning

Role of Mitochondria in Axonal Morphogenesis observation to obtain a final K1 concentration of 50 mM. For long-term real-time imaging, GCs were plated on CellView cell culture dish (Greiner Bio-One; Frickenhausen, Germany) coated with poly-L-lysine and transfected as described above. At 5 DIV, dishes were set into CellVoyager CV1000 live cell confocal system (Yokogawa Electric; Tokyo, Japan) and incubated with 5% CO2 at 37 C. Images were captured every 5 min for up to 12 h.

Analysis of Mitochondrial Movement Kymographs were created using ImageJ as described previously (Miller and Sheetz, 2004; Kang et al., 2008; MacAskill et al., 2009). Briefly, stacked images were opened in ImageJ, and a region along the entire axon main shaft was straightened. The stacks of straightened images were resliced and z-projected with a max projection option to generate the kymographs. For all kymographs, the height represents time (10 min) and the width represents the distance along the axon (150 lm). The movement of the mitochondria in the kymographs (shown as diagonal lines) was traced using Adobe Photoshop. We defined a threshold velocity of 0.1 lm/s, which was equivalent to a displacement of one pixel between two adjacent frames. If a mitochondrion paused for more than 10 s (three consecutive frames) or reversed direction, it was counted and traced as a new mitochondrion. Mitochondrial motility was calculated as the summation of the total mitochondrial displacement and was normalized by the length of the axon in a single direction per minute. If this value is 50 (lm/min/mm axon), we can expect to observe one moving mitochondrion at any given point along the axon over 1000 lm/50 lm 5 20 min. This means that, if we continuously observe mitochondrial dynamics, we would see 72 moving mitochondria over 24 h.

Analysis of Axonal Morphology and Mitochondrial Localization At 5 DIV, cultured GCs were stimulated with high K1 medium ([K1] 5 50 mM) for 20 min. At 6 or 8 DIV, cells were fixed with 4% paraformaldehyde at 37 C for 30 min. After fixation, the cells were washed three times with phosphate-buffered saline (PBS). Images were acquired using a 203 objective (NA, 0.75; Nikon) with an ECLIPSE TE300 inverted microscope (Nikon) that was equipped with an ORCA II cooled CCD camera (Hamamatsu Photonics; Hamamatsu, Japan) and was controlled by AQUACOSMOS software (Hamamatsu Photonics). Axonal length was measured using ImageJ, and only protrusions that were longer than 2 lm were defined as axon branches.

Immunocytochemistry and Immunoblotting For immunocytochemistry, the cells were fixed and washed as described above and then incubated with blocking

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solution (PBS with 2% goat serum and 0.01% Triton X) for 1 h at room temperature. The following primary antibodies were used: rabbit antitranslocase of outer membrane 20 (Tom20; 1:500; Santa Cruz Biotechnology; San Francisco, CA), mouse anti-Prox-1 (1:1000; Millipore; Bedford, MA), and rabbit anti-phospho-AMPKa (Thr172; 1:1000; Cell Signaling Technology; Danvers, MA). After incubation with the primary antibodies at 4 C overnight, the cover slips were washed three times with PBS. The following secondary antibodies or staining reagents were used: antimouse IgG Alexa-350 (1:400; Invitrogen), anti-rabbit IgG Alexa-488 (1:400; Invitrogen), and rhodamine phalloidin (1:40; Invitrogen). For immunoblotting, GCs were cultured at a density of 6 3 105 cells/cm2 and were stimulated with high K1 media for 20 min at 5 DIV as described above. Immediately after stimulation, the cover slips were washed twice with cold PBS, homogenized, and lysed with lysis buffer that contained protease inhibitors (RIPA buffer; Nakalai Tesque) and phosphatase inhibitor cocktail (Nakalai Tesque). The extracted protein was denatured by heating in sample buffer (EzApply, ATTO; Tokyo, Japan), followed by sodium dodecyl sulfate polyacrylamide gel electrophoresis. The samples were separated electrophoretically in the polyacrylamide gel (e-PAGEL; ATTO) and then transferred to polyvinylidene difluoride membranes. The membranes were incubated with the following primary antibodies: rabbit anti-phospho-AMPKa (Thr172; 1:1000; Cell Signaling Technology) and mouse anti-b-actin (1:1000; SigmaAldrich). Next, horseradish peroxidase (HRP)-conjugated anti-rabbit IgG (Nakalai Tesque) or HRP-conjugated antimouse IgG (1:1000; Sigma) were used as the secondary antibodies. The membranes were washed, and proteins were detected using 3,30 ,5,50 -tetramethylbenzidine solution (EzWest Blue; ATTO). Equal protein loading was confirmed by staining with Coomassie Brilliant Blue or by probing the membranes with an anti-b-actin antibody. For semiquantitative analysis, band densitometry was performed with scanned images of the immunoblot membranes using ImageJ.

Monitoring the Intracellular ATP Levels Using Mg-Green To study the intracellular ATP levels, the neurons were incubated with 10 lM magnesium (Mg)-Green AM (Invitrogen) for 30 min. The emission intensity of this probe increased as a function of free intracellular Mg21; however, it decreased with ATP content because of the high binding affinity of this probe to ATP as compared to ADP (Lee and Peng, 2008). Mg-Green has a high binding affinity to Ca21 and therefore Ca21 is likely to interfere with Mg21 detection using this agent. However, there is a line of evidences which suggest that the increase in [Ca21]i does not affect the fluorescence intensity of Mg-Green (Lee and Peng, 2008). In ciliary neurons, ATP depletion by two distinct chemical agents elevates [Ca21]i up to 400 nM, which did not increase the fluorescence intensity of Mg-Green Developmental Neurobiology

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Figure 1

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Role of Mitochondria in Axonal Morphogenesis (Bernstein and Bamburg, 2003). Jha et al. (2002) also confirmed that an application of 1 mM caffeine, which elevates [Ca21]i up to 645 nM, does not increase the fluorescence intensity of Mg-Green. The same conclusion was stated in another article using cardiomyocytes (Leyssens et al., 1996). A demonstration that high K (56 mM) depolarization-induced increase in [Ca21]i is 500–600 nM at the highest even when the extracellular solution contains physiological concentration of Ca21 (an experiment using cerebellar granuel cell cultures; Ciardo and Meldolesi, 1991), which supports us to conclude that the increase in the fluorescence intensity of Mg-Green observed in our study is not merely induced by the increase in [Ca21]i. Labeled neurons were washed and incubated in the culture media for 30 min to maximize de-esterification of the probe before the pharmacological treatments. For imaging, the cover slips were incubated in Ca21-free artificial cerebrospinal fluid (ACSF) that was composed of 127 mM NaCl, 1.6 mM KCl, 1.24 mM KH2PO4, 1.3 mM MgSO4, 26 mM NaHCO3, and 10 mM glucose. Images were captured every 60 s for 20 min.

Data Representation and Statistical Analyses The data are represented as the mean 6 SEM. The MannWhitney U test or Steel-Dwass test after the Kruskal-Wallis test was used for nonparametric statistics, and Student’s t test or Tukey’s test after analysis of variance (ANOVA) were used for parametric statistics. ANOVA for repeated measurements was used for the statistical analysis of the Mg-Green intensity and long-term real-time imaging. Data were pooled from at least 3 independent culture preparations. Data were collected and statistically analyzed independently by two people in a blind manner to avoid bias.

RESULTS Depolarization Induced Mitochondrial Accumulation and Axonal Branch Formation To visualize mitochondrial distribution in the axons, primary GC cultures were prepared from P6 rat pups and cotransfected with mitochondria-targeted Disco-

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soma sp. red (Mito-DsRed; DsRed2 fused to the mitochondrial targeting sequence of cytochrome c oxidase) and membrane-targeted Aequorea coerulescens green fluorescent protein (GFP) (Mem-AcGFP; AcGFP1 fused to the membrane targeting sequence of GAP43) at 1 DIV [Fig. 1(A,B)]. All the MitoDsRed1 puncta were immunopositive for the mitochondrial marker Tom20 in the GC axons [Fig. 1(D)]. First, to examine whether neuronal depolarization cell autonomously affects axonal morphogenesis, we used purified primary GC cultures and selected GCs that did not contact another cell, which allowed for the examination of the effects of depolarization at the single-cell level. This process allowed us to exclude the influence of cell–cell interactions, which are known to affect mitochondrial motility (MacAskill et al., 2009; Kiryu-Seo et al., 2010). We depolarized the dissociated GC cultures with 50 mM KCl for 20 min at 5 DIV [Fig. 1(A)]. Three days later (8 DIV), the stimulated GCs exhibited aberrant branches [Fig. 1(B,G–I)] and longer axons [Fig. 1(B,E)]. We also observed that the number of axonal mitochondria was significantly increased [Fig. 1(J)]. To determine whether the increased mitochondria contribute to activity-dependent axon branching, we performed a detailed observation of mitochondrial localization after a depolarizing stimulation with high K1 [Fig. 2(A)]. The mitochondria were predominantly observed to be localized at the bases of the axonal branch points [Fig. 2(B)]. To further investigate the role of mitochondria in axonal branch formation, we classified the branches into two groups according to the difference in length that results from the alteration of the molecular components (Dent and Kalil, 2001); the two groups were actin-rich short (10 lm) branches, which were continuously added and retracted [Fig. 2(B), arrows], and microtubulerich long (>10 lm) stable branches [Fig. 2(B), arrowheads]. One day after stimulation (6 DIV), short branches with mitochondria present in their bases (w/ o Mito) significantly increased, whereas there was no difference in the number of short branches without mitochondria (w/o Mito) [Fig. 2(C)]. Three days after the stimulation (8 DIV), we observed a greater

Figure 1 Depolarization induces axonal branch formation and mitochondrial accumulation. (A) Experimental scheme. Cultured GCs were stimulated by high K1-medium (50 mM) for 20 min at 5 DIV. (B) Stimulated GCs (right) established aberrant axon branches 3 days after the depolarization. (C) Magnified view of the boxed region in (B). Mitochondria were localized at the branch points (arrows). (D) The distribution of Mito-DsRed was thoroughly colocalized with the expression pattern of Tom20. (E–K) Total axonal length (E), total branch length (G), number of axonal branches (I), and the total number of mitochondria (J) in the axons were significantly increased in depolarized GCs, while main shaft length (F), mean branch length (H), and the density of mitochondria (K) were comparable. *, P < 0.05; **, P < 0.01; Student’s t-test or Mann-Whitney U-test, n 5 12–16 cells. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.] Developmental Neurobiology

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Figure 2 Mitochondrial localization and ATP production are necessary for depolarization-induced formation of axon branches. (A) Experimental scheme. FCCP (0.1 or 1 lM) was applied from 6 to 8 DIV. (B) Mitochondria were localized at the bases of both short (10 lm; arrows) and long (>10 lm; arrowheads) branches. Open arrows indicate short branches without mitochondria. (C,D) The number of axon branches with mitochondria (w/o Mito), but not those without mitochondria (w/o Mito), significantly increased after high K1 stimulation. *, P < 0.05; **, P < 0.01; Steel-Dwass test after Kruskal-Wallis test, n 5 12–16 cells. (E–G) About 90% of long branches had mitochondria at their bases in both control and high K1 conditions (E), whereas high K1 stimulation significantly increased the proportion of short branches with mitochondria (F). (G) Significantly higher proportion of mitochondria was associated to branch points after the high K1 stimulation compared with control. *, P < 0.05; **, P < 0.01; Tukey’s test after two-way ANOVA, n 5 12–16 cells. (H–K) The increase in the total axon length (H), total branch length (I), number of branches (J), and the number of mitochondria (K) were significantly suppressed by FCCP application. *, P < 0.05; **, P < 0.01; SteelDwass test after Kruskal-Wallis test or Tukey’s test after one-way ANOVA, n 5 6–10 cells. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

number of long branches compared with 6 DIV [Fig. 2(B), arrowheads]. Furthermore, we observed that the number of long branches with mitochondria significantly increased in the high K1-treated GCs comDevelopmental Neurobiology

pared with control GCs [Fig. 2(D)]. We found that about 90% of long branches at 8 DIV had mitochondria at their base in both control and high K1 conditions [Fig. 2(E)], suggesting that mitochondrial

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localization is associated with the maintenance of such long axonal branches. In contrast, we found that significantly higher proportion of short branches at 8DIV had mitochondria at their base in high K1 condition compared with control [Fig. 2(F)]. This result suggests that mitochondrial localization is associated with the emergence of such short axonal branches. To further analyze the relationship between axonal branches and mitochondrial localization from the opposite direction, we calculated the proportion of mitochondria associated to branch points in control condition and upon high K1 stimulation at 1 day (6 DIV) or 3 days (8 DIV) after the treatment. As a result, we found that significantly higher proportion of mitochondria was associated to branch points after the high K1 stimulation compared with control [Fig. 2(G)]. Our results, including the observation that axonal mitochondria increased after the depolarizing stimulation [Fig. 1(J)], suggest that the mitochondria contribute to the activity-dependent emergence and retention of axon branches. To test this hypothesis, we pharmacologically inhibited the function of mitochondria with the uncoupler FCCP, which is known to inhibit the production of ATP by abolishing mitochondria membrane potential (Lee and Peng, 2008), after the depolarizing stimulation [Fig. 2(A)]. FCCP prevented high K1-induced branch formation and the net growth of axons [Fig. 2(H–K)], which indicates that mitochondrial ATP production is required for branch formation. We confirmed that the treatment of cultures with the reagents at the concentration used in this study did not affect the cell viability in our culture system.

Depolarization Modulated the Axonal Transport of Mitochondria Previous studies have suggested that both mitochondrial motility and localization are affected by neuronal activity. It has been reported that local stimulation of dendritic spines leads to the accumulation of mitochondria in an N-methyl-D-aspartate (NMDA) receptor-dependent manner within minutes to hours (Li et al., 2004; MacAskill et al., 2009); however, the chronic effect of neuronal depolarization on mitochondrial dynamics in axons and its morphological consequences are largely unknown. To directly examine whether depolarization affects mitochondrial dynamics and axonal morphogenesis, we conducted real-time imaging of mitochondria from cultured GCs. Images of Mito-DsRed1 mitochondrial puncta in Mem-AcGFP1 axons of the GCs were captured every 5 s for 20 min using confocal microscopy at 6 DIV. The dynamics and distance of the

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mitochondrial movement were assessed by drawing kymographs as described previously (Miller and Sheetz, 2004) [Fig. 3(A)]. Mitochondria were equivalently transported in the anterograde direction toward the axon tip (blue) and in the retrograde direction toward the cell body (red) in the control condition [Fig. 3(A,B)]. The average velocity of motile mitochondria in control GCs (anterograde: 0.27 6 0.09 lm/s, retrograde: 0.41 6 0.16 lm/s; mean 6 SD; n 5 534 anterograde and 560 retrograde traces of motile mitochondria from 12 axons) were within the range reported in previous studies using rodent hippocampal neurons (Kang et al., 2008; Wang and Schwarz, 2009; MacAskill and Kittler, 2010). To relate mitochondrial dynamics to their quantitative accumulation in the axons, we adopted total mitochondrial displacement (Miller and Sheetz, 2004) as an index. In our experimental system, the flux of mitochondria in the axons was 0.021 6 0.016 (mean 6 SD) mitochondrion/mitochondria in the anterograde direction and 0.023 6 0.016 mitochondrion/ mitochondria (mean 6 SD) in the retrograde direction, which were comparable to the previous report (Miller and Sheetz, 2004). To investigate the effect of depolarization on the dynamics of mitochondrial movement, we depolarized GCs with 50 mM high K1 on the previous day (5 DIV). We observed that the depolarizing stimulation increased the net anterograde distance of mitochondrial movement [Fig. 3(B)] and resulted in an increased number of mitochondria in the axons [Fig. 3(C)]. To further characterize the modulation of mitochondrial transport following the depolarizing stimulation, we performed long-term time-lapse imaging of mitochondrial localization up to 12 h after the stimulation [Fig. 3(D–F)]. Although we observed that the stimulation acutely and completely arrested mitochondria in both directions [Fig. 3(D)], dynamics of axonal mitochondria recovered 1 h after the stimulation [Fig. 3(E)]. By calculating net anterograde moving distance at each point, we found that the imbalance of mitochondrial transport appeared as early as 4 h after depolarization [Fig. 3(F)]. These results suggest that neuronal activity rebalanced the anterograde and retrograde transport of the mitochondria toward accumulation in the axons.

Mitochondrial Localization Was Correlated with Both Emergence and Retention of Axonal Branches As shown in Figure 2, mitochondria were predominantly localized at the bases of the axonal branch points. To shed light on the spatiotemporal Developmental Neurobiology

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relationships between mitochondrial localization and axon branching, we performed long-term real-time imaging of axonal morphology and mitochondrial distribution simultaneously between 5 and 6 DIV. Representative simultaneous imaging of mitochondria and axonal morphogenesis indicate that Mito-DsRed1 puncta were observed (open arrowhead at 25min) before the emergence of an axonal protrusion (solid arrow at 0 min) [Fig. 4(A)]. Importantly, real-time imaging allowed us to quantitatively analyze the probability that mitochondria were observed at the bases of axon branches (probability of mitochondrial localization: Pm) during their growth, showing that mitochon-

dria were observed (25 min, black arrow) before the emergence of axon branches (defined as 0 min, green arrow) [Fig. 4(B)]. Furthermore, the axon branches with the higher Pm throughout their lifetime tended to have the longer lifetime [Fig.4(C,D)], which is consistent with the finding that axon branches with mitochondria are maintained and elongated compared with those without mitochondria (Fig. 2). During time-lapse imaging with 5-min interval, we did not find significant increase in mitochondrial density at any time point (data not shown), which is consistent with the data from fixed samples 3 days after the depolarization [Fig. 1(J)]. This indicates that the

Figure 3 Developmental Neurobiology

Role of Mitochondria in Axonal Morphogenesis

mitochondrial delivery and the axonal growth occurred quite simultaneously and that the density of mitochondria is within a stable range as long as physiological development of axons undergoes, which was accomplished by instantaneous recruiting of mitochondria in correlation with axonal growth. These results suggest that mitochondrial localization precedes the emergence of axon branches and that mitochondria are necessary for the retention of branches.

Activation of AMPK Was Related to Mitochondrial Accumulation and Branch Formation We focused on AMPK, which is activated by AMP, as an intracellular signal that mediates the depolarization-dependent mitochondrial accumulation in axons for two reasons. First, activated (depolarized) neurons have been shown to consume large amounts of ATP to restore and maintain the membrane potential or to release and uptake neurotransmitters, resulting in the production of AMP (Ronnett et al., 2009; Weisova et al., 2009). Second, activation of AMPK has been shown to facilitate mitochondrial biogenesis and increase the total amount of mitochondria (Reznick and Shulman, 2006; J€ager et al., 2007). Therefore, we hypothesized that AMPK activation underlies the modification of mitochondrial transport and accumulation. To test this hypothesis, we first examined whether neuronal depolarization activates AMPK immedi-

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ately after the 20-min high K1 stimulation at 5 DIV (Fig. 5). Considering that 20 min is too short to induce changes in the total expression levels of AMPK, we assume that the increase in pAMPK signals in this time course directly reflect the phosphorylation levels of AMPK. Immunocytochemical [Fig. 5(A,B)] and immunoblot [Fig. 5(C)] analyses using antibodies against phosphorylated (activated) AMPK (pAMPK) revealed that stimulation increased the phosphorylation of AMPK in cultured GCs. These phenomena were pharmacologically confirmed by either a specific activator or an inhibitor of AMPK. The AMPK inhibitor CC (10 mM) blocked high K1induced increase in pAMPK immunoreactivity, whereas AICAR (2.5 mM), an AMPK activator, was able to increase the activity [Fig. 5(A,B)]. To determine whether the depolarizing stimulation enhanced ATP consumption in neurons, the intracellular ATP levels were determined by monitoring the fluorescence of the ATP-probe Mg-Green in GCs; the fluorescence intensity negatively correlates with the intracellular ATP levels (see Materials and Methods section). We observed an increase in Mg-Green fluorescence signals in GCs during stimulation, which suggested a decrease in intracellular ATP concentration [Fig. 5(D,E)]. Depolarization also activates several intracellular signals by Ca21 influx through voltage-gated Ca21 channels; however, high K1induced AMPK activation was not blocked using Ca21-free media [Fig. 5(F,G)]. Additionally, previous studies demonstrated that the increase in intracellular Ca21 levels induced by the ATP depletion or

Figure 3 The dynamics of axonal mitochondria are chronically modulated by the activation of AMPK. (A) Representative kymographs (6 DIV) 1 day after 20 min application of vehicle (Control), high K1 media (high K1; 50 mM), and an AMPK activator AICAR (2.5 mM) at 5 DIV. Mito-DsRed signals were captured with real-time imaging and are represented as kymographs (height, 10 min; top). Anterograde movement (blue) and retrograde movement (red) were digitally traced on the kymographs (bottom). (B) Net anterograde distance of mitochondrial movement (anterograde moving distance—retrograde moving distance) showed that the balance of mitochondrial dynamics significantly shifted toward the anterograde direction after depolarization. This shift was blocked by application of the AMPK antagonist, CC (10 lM). AICAR reproduced the change in dynamics by depolarization. **, P < 0.01 between anterograde distance and retrograde distance; Tukey’s test after two-way ANOVA, n 5 8–12 cells. (C) The number of mitochondria in the axons at 8 DIV. The increase by depolarization was suppressed by CC, and AICAR application resulted in an accumulation of mitochondria. *, P < 0.05 versus control and #, P < 0.05 versus high K1; Tukey’s test after one-way ANOVA, n 5 10 cells. (D) Depolarization with high K1 rapidly and completely arrested mitochondrial movement in both directions. Anterograde movement (blue) and retrograde movement (red) were digitally traced on the kymographs (right). (E) Kymographs (left) of 1, 2, 4, 6, 9, and 12 h after depolarization from an identical axon. Anterograde movement (blue) and retrograde movement (red) were digitally traced on the kymographs (right). (F) The balance of the dynamics were significantly shifted toward the anterograde direction as early as 4 h after depolarization. *, P < 0.05; **, P < 0.01 between anterograde moving distance and retrograde moving distance; Tukey’s test after two-way ANOVA, n 5 6 cells. Gray lines represent net anterograde moving distance of each cell. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.] Developmental Neurobiology

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Figure 4 Mitochondrial localization is correlated with both emergence and retention of axon branches. (A) GCs were observed every 5 min for up to 12 h between 5 and 6 DIV. Localization of a mitochondrion (middle; open arrowheads) preceded the emergence of an axon branch (top; solid arrowheads) by 5 min (bottom; open and solid arrows). (B) Probability that mitochondria were observed (Pm) rose 5 min before (time 25; black arrow) the emergence of axon branches (time 0; green arrow). Gray area shows the mean 6 SD of the Pm at randomly-selected ROIs along the axons. n 5 25 axon branches from eight cells. At each time point, the probability was defined as 100% if axon branches owned mitochondria at their bases and 0% if the branches were without mitochondria. Values represent the mean 6 SEM. (C,D) Axon branches with the longer lifetime showed the higher Pm. The number of axon branches is indicated by the size of circles in (C). The Pm is an average value during the lifetime of each axon branch. (D) Branches were divided into two groups according to their Pm (threshold is median value). Branches with high Pm showed the significantly longer lifetime than branches with low Pm. **, P < 0.01; Mann-Whitney U-test, n 5 32 for high Pm and 33 for low Pm branches. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

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Figure 5 Depolarization activates AMPK through ATP consumption in a calcium-independent manner. (A) Immunocytochemistry by anti-pAMPK (Thr172) antibody of the control, high K1 (50 mM for 20 min)-, high K1 1 CC (10 lM)- and AICAR (2.5 mM for 20 min)-treated GCs. The soma and nuclei were detected by F-actin (stained with rhodamine–phalloidin; red) and Prox1 (blue), respectively. (B) Both depolarization and AICAR activated AMPK, and the simultaneous application of CC blocked activation. **, P < 0.01 versus control, ##, P < 0.01 versus high K1; Tukey’s test after one-way ANOVA, n 5 11–15 cells. (C) Immunoblot analysis also confirmed that depolarization induced AMPK activation. **, P < 0.01 versus control; Student’s t-test, n 5 6. (D,E) ATP consumption after depolarization visualized by Mg-Green (arrowheads). **, P < 0.01 versus control; repeated measure ANOVA, n 5 5–10 cells. (F,G) Depolarization activated AMPK under a calcium-free condition. **, P < 0.01 versus control; Tukey’s test after one-way ANOVA, n 5 26– 43 cells. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

the depolarizing stimulation does not affect the fluorescence intensity of Mg-Green (Lee and Peng, 2008). These results suggest that neuronal activity induces the consumption of ATP and activates AMPK, probably via an increase in intracellular AMP.

Next, we determined that the activation of AMPK is involved in the modification of mitochondrial transport and subsequent axonal growth. The inhibition of AMPK with 10 mM CC blocked the depolarizationinduced increase in the net anterograde distance Developmental Neurobiology

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Figure 6 AMPK activation is required for depolarization-induced axonal branch formation. (A) Representative images of control, high K1 (50 mM for 20 min)-, high K1 1 CC (10 lM)-, and AICAR (2.5 mM for 20 min)-treated GCs at 3 days after stimulation. Both high K1- and AICARtreated GCs exhibited aberrant axonal arborization. CC inhibited depolarization-induced phenomena. (B–D) Quantification of the morphological properties shown in (A). Total axon length (B), total branch length (C), and number (D) were significantly increased in both high K1- and AICARtreated groups. CC inhibited depolarization-induced phenomena. *, P < 0.05; **, P < 0.01 versus control, and #, P < 0.05 versus high K1; Tukey’s test after one-way ANOVA or Steel-Dwass test after Kruskal-Wallis test, n 5 9–16 cells.

of mitochondrial movement [Fig. 3(B)] and the number of axonal mitochondria [Fig. 3(C)]. In contrast, the activation of AMPK with a 20-min application of 2.5 mM AICAR at 5 DIV resulted in an increase in the anterograde distance [Fig. 3(A,B)] and the number of mitochondria [Fig. 3(C)]. Similarly, increase in the number and the length of axon branches induced by depolarization was blocked by CC and reproduced by AICAR (Fig. 6). These results suggest that AMPK underlies neuronal activity-induced mitochondrial accumulation and axonal growth. Developmental Neurobiology

DISCUSSION Overview We revealed that the neuronal depolarization cellautonomously induced the rebalance of mitochondrial transport and their eventual accumulation in axons through AMPK activation, which resulted in axonal branch formation in an ATP-dependent manner. Such a mechanism may enable neurons to establish proper axonal projections that reflect neuronal activity.

Role of Mitochondria in Axonal Morphogenesis

Causality between Mitochondrial Accumulation and Axonal Morphogenesis We showed that only the axon branches with mitochondria at their bases significantly increased after depolarization (Fig. 2). We acknowledge that it is difficult to conclude whether mitochondria found at branch points are stationary or just passing through or pausing at when cultures were fixed for immunostaining. We found that it is still difficult to completely discriminate stationary mitochondria from moving ones that are associated with branch formation even with a time-lapse imaging (Fig. 4). Considering that mitochondrial localization preceded the branch formation [Fig. 4(B)], that unstable axonal branches (lifetime 5 0) were not usually accompanied by mitochondria [Fig. 4(C)], and that over 80% of long branches had mitochondria at their branch points [Fig. 2(E)], we assume that most of mitochondria found at branch points in fixed neurons, especially those with long branches, were stationary. We still do not exclude the possible involvement of moving mitochondria in the immunostained images. However, the most important point here is that branching points had significantly higher Pm compared with nonbranching points (Fig. 4) when the regions of interest (ROIs) were randomly selected along the axon at the initial time point [Fig. 4(B,C)], which indicates that future branch points had more chance to be supplied with mitochondria-derived ATP than nonbranching points regardless of mitochondria are stationary or moving. To perform the time-lapse imaging of both mitochondrial dynamics and axon branching simultaneously and also to keep the phototoxicity at the minimum, we adopted 5 min as an interval during time-lapse imaging. Therefore, the mitochondria observed in Figure 4(A) are not identical. However, we think that the mitochondria observed at branch points do not need to be identical. Indeed, we repeatedly observed the phenomena that localized mitochondria at a specific branch point frequently turn over, while overall probability of mitochondrial localization (Pm) remains significantly higher than randomly selected ROIs along the axon [Fig. 4(C)]. This indicates that, regardless of whether or not the localized mitochondria are identical, the higher Pm underlies the longer lifetime of axonal branches probably via ATP supply [Fig. 4(E)]. Most of the branches with mitochondria were short at 6 DIV and elongated 2 days later (Fig. 2). These observations at two separate time points imply the possibility that mitochondrial accumulation precedes branch formation and elongation. Given that artificial

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extension-induced elongation of axon branches is not followed by mitochondrial localization (Ruthel and Hollenbeck, 2003), this possibility seems feasible. There have been previous reports on mitochondrial dynamics and neuronal morphogenesis, which revealed that (1) anterograde transport and mitochondrial accumulation preceded the establishment of neuronal polarity (Bradke and Dotti, 1997), (2) augmentation of ATP production and mitochondrial fission increased the number of dendritic spines (Li et al., 2004), and (3) local application of nerve growth factor (NGF) localized the mitochondria and induced branch formation (Chada and Hollenbeck, 2004). These reports along with our further finding that mitochondria were localized beforehand at the regions where axon branches to emerge (Fig. 4), support our hypothesis that accumulated mitochondria induce branch formation. However, our experiments using FCCP do not allow us to readily conclude that local, but not global, ATP levels are critical for axon branching. Localized mitochondria may contribute to axon branching possibly through control of ATP production and/or Ca21 homeostasis or through other unknown mechanisms as previously pointed out (Courchet et al., 2013). Our observation raises two questions on the relationship between colocalization of mitochondria and axon branches. First, why are not all mitochondria colocalized with branches? This may be a result of the functional heterogeneity of each mitochondrion along the axon, such as the variation in the energy supply of each axonal mitochondrion. Although it was recently reported that there was no difference in the transmembrane potential of each axonal mitochondrion, except those localized at growth cones (Verburg and Hollenbeck, 2008), there is still the possibility that mitochondria that are freshly fragmented and transported into axons have a relatively high rate of ATP production. Second, why do not all branches, especially the short branches, accompany mitochondria? Such short branches without mitochondria are destined to disappear, which is supported by our observation that there was almost no long branch without mitochondria at three days after depolarization. Additionally, by longterm real-time imaging, we confirmed that axon branches with low Pm were transitory (Fig. 4). This finding also supports our hypothesis that mitochondrial localization is necessary for the elongation and retention of newly emerged axon branches. To further confirm that mitochondrial localization is sufficient to induce axon branching, artificial trapping of mitochondria should be demonstrated in future studies. There are at least two possible mitochondrial dynamics that are required for the proper axonal Developmental Neurobiology

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development. First one is to increase the fraction of stationary mitochondria, as Courchet et al. (2013) showed that axon branches are induced when mitochondria are recruited to branch points and remained stationary, rather than changing motility. Second one is to increase the number of axonal mitochondria to satisfy metabolic demands. The motivation of our study which focused on the balance of mitochondrial motility originates from the latter: if the more axonal branches emerge after the depolarization, do the more mitochondria have to be recruited into axons? Such recruited mitochondria can be trapped at nascent branch points and would contribute to the emergence and maintenance of new axonal branches. Although future studies using in vivo animals and/or organotypic cultures, in which interaction between both neuronal and non-neuronal cells are maintained, are necessary to examine whether axonal branches induced by growth factors and neuronal activity are through different signaling pathways, we speculate that both the increase in the fraction of stationary mitochondria and the recruitment of mitochondria into axons are required for the proper axonal development under the physiological conditions. It is also possible that growth factor-induced local metabotropic demands and depolarization-induced robust metabotropic demands in axons may cause different mitochondrial dynamics.

Downstream of AMPK We found that AMPK activation following neuronal depolarization rebalanced the mitochondrial dynamics, which led to an accumulation in the axons and resulted in axonal branch formation. Although the molecular link between AMPK and mitochondrial transport has yet to be elucidated, there are several possible candidates. First, the function and expression level of molecular motors may be modified. Axonal transport of mitochondria in an anterograde direction is dependent primarily on kinesin superfamily protein (KIF)5 and KIF1B motor proteins (Tanaka et al., 1998; Hirokawa and Takemura, 2005). Importantly, the KIF5 kinesin light chain was phosphorylated by AMPK (Amato et al., 2011). In Aplysia neurons, activity-dependent upregulation of the kinesin subunit leads to the facilitation of anterograde transport, which requires cAMP response element-binding protein (CREB) activity (Puthanveettil et al., 2008). Similarly, considering that AMPK regulates the expression levels of various proteins, including peroxisome proliferator-activated receptor gamma coactivator 1a (PGC-1a) through CREB (Herzig et al., 2001; J€ager et al., 2007; Developmental Neurobiology

Thomson et al., 2008) and KIF5, which is upregulated by AMPK through CREB, it is hypothesized that AMPK activation increases axonal mitochondria cooperatively with PGC-1a. Second, modification of microtubule-associated proteins (MAPs), such as tau or MAP1B, may affect mitochondrial dynamics and branch formation. They are phosphorylated directly or indirectly by AMPK, which can alter the stability of microtubules and axonal transport (Jimenez-Mateos et al., 2006; Qiang et al., 2006; Dixit et al., 2008). Katanin is a microtubule severing protein that facilitates axon branch formation in an ATP-dependent manner. Given that phosphorylated tau detaches from microtubules, which removes its protection to the microtubules against severing proteins, it is conceivable that AMPK-dependent phosphorylation of tau induces katanin-dependent disruption of local microtubules and can lead to branch formation. To support this hypothesis, further experiments to examine the phosphorylation of MAPs after depolarization will be necessary. In this study, we did not determine whether transcriptional changes induced by high K1 stimulation was involved in the rebalance of mitochondrial motility and axon branching. Although there remains a possibility that upregulation of the local biogenesis of axonal mitochondria is involved in their accumulation, the contribution has been estimated to be minor, which is based on a previous study in which the authors assessed mitochondrial biogenesis in axons (Amiri and Hollenbeck, 2008). Therefore, we propose more presumable hypothesis that the augmented biogenesis of mitochondria in the soma, along with upregulated KIFs, drive the accumulation of mitochondria in the axons. Previous studies reported that both AICAR and CC could exert AMPK-independent effects. Although AMPK-independent pathways that are sensitive to both AICAR and CC have not been reported so far, our experiments using pharmacological reagents do not exclude the possible involvement of unknown pathways that are independent of AMPK. LKB1, one of the upstream kinase of AMPK, might be involved in activity-dependent axon branching. Our results also do not exclude the possible involvement of AMPK-related kinases, such as NUAK1 (Courchet et al., 2013) in activity-dependent axon branching.

Activity Dependency and Developmental Specificity We adopted depolarizing stimulation using 50 mM KCl for the reliable and reproducible activation of

Role of Mitochondria in Axonal Morphogenesis

cultured neurons. Although it is intriguing to examine whether axon branching induced by physiological neuronal activity and aberrant activity such as under epileptic conditions cause different modulation of AMPK function in vivo, especially because the axons of dentate GCs, that is, the mossy fibers, are known to exhibit aberrant branching called as the mossy fiber sprouting in the epileptic brain (Nadler, 2003; Koyama and Ikegaya, 2004; Sutula and Dudek, 2007). Considering that genetic deletion of AMPK has little or no consequences on brain development (Williams et al., 2011), AMPK-dependent modulation of mitochondrial dynamics and axon branching might be prominent when neurons undergo hyperactivity. Previous studies demonstrated that the prolonged activation of AMPK disrupts neuronal polarization (Amato et al., 2011; Williams et al., 2011) and is deleterious to dendritic spines of mature neurons (Mairet-Coello et al., 2013). In contrast to these studies, our method using depolarization or a pharmacological agent for 20 min may activate AMPK in a relatively transient manner. Moreover, we used only immature GCs specifically prepared from the dentate hilar region of the postnatal rat hippocampus. The neurons were cultured for 5–8 days, which is enough long to establish neuronal polarity but still too short for neurons to be completely mature. We observed that mitochondrial dynamics in older GCs were less prominent (data not shown), which is consistent with the previous study showing that the number of moving mitochondria in neuronal processes are lower in older cortical neurons in vitro (Chang and Reynolds, 2006). These properties of mitochondria may underlie the developmental specificity of AMPKdependent axon branching. Future studies will need to explore this hypothesis. In conclusion, this study identified AMPK as a mediator that translates neuronal activity to axonal morphogenesis in a cell-autonomous manner through modification of mitochondrial transport. The authors thank the members of Yakusaku Lab (Laboratory of Chemical Pharmacology, Graduate School of Pharmaceutical Sciences, The University of Tokyo, Tokyo, Japan) for helpful and constructive discussions on this study.

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