Inhibition of excessive mitochondrial fission reduced aberrant autophagy and neuronal damage caused by LRRK2 G2019S mutation

Abstract

LRRK2 G2019S mutation is the most common genetic cause of Parkinson's disease (PD). Cellular pathology caused by this mutant is associated with mitochondrial dysfunction and augmented autophagy. However, the underlying mechanism is not known. In this study, we determined whether blocking excessive mitochondrial fission could reduce cellular damage and neurodegeneration induced by the G2019S mutation. In both LRRK2 G2019S-expressing cells and PD patient fibroblasts carrying this specific mutant, treatment with P110, a selective peptide inhibitor of fission dynamin-related protein 1 (Drp1) recently developed in our lab, reduced mitochondrial fragmentation and damage, and corrected excessive autophagy. LRRK2 G2019S directly bound to and phosphorylated Drp1 at Threonine595, whereas P110 treatment abolished this phosphorylation. A site-directed mutant, Drp1^T595A, corrected mitochondrial fragmentation, improved mitochondrial mass and suppressed excessive autophagy in both cells expressing LRRK2 G2019S and PD patient fibroblasts carrying the mutant. Further, in dopaminergic neurons derived from LRRK2 G2019S PD patient-induced pluripotent stem cells, we demonstrated that either P110 treatment or expression of Drp1^T595A reduced mitochondrial impairment, lysosomal hyperactivity and neurite shortening. Together, we propose that inhibition of Drp1-mediated excessive mitochondrial fission might be a strategy for treatment of PD relevant to LRRK2 G2019S mutation.

INTRODUCTION

Parkinson's disease (PD) is the second most common neurodegenerative disorder characterized by progressive and substantial loss of dopaminergic (DA) neurons in the substantia nigra (1,2). Leucine-rich repeat kinase 2 (LRRK2) is a protein encoded by the PARK8 locus. It has a conserved serine-threonine kinase mitogen-activated protein kinase kinase kinase (MAPKKK) domain, a member of the Roc (Ras of complex) GTPase family (3–5). While over 50 variants have been identified throughout the different LRRK2 domains in PD patients, the mutation G2019S (Gly2019 to Ser) that takes place in the MAPKKK domain has been recognized as the most common cause of dominant familial PD and accounts for up to 2% of sporadic PD cases (6). The G2019S mutant augments the kinase activity of LRRK2, which is associated with increased toxicity in DA neurons (7). However, the molecular mechanism by which LRRK2 G2019S causes neurodegeneration is poorly understood.

Mitochondria form a dynamic network of interconnected tubules which is maintained by the balance between fission and fusion events (8). These dynamic processes maintain mitochondrial function by enabling mitochondrial recruitment to critical subcellular compartments and mitochondrial quality control (8–10). A set of conserved large GTPases belonging to the dynamin family controls mitochondrial fusion and fission; Opa1 (optic atrophy 1) and mitofusin 1/2 regulate fusion, whereas dynamin-related protein 1 (Drp1) mediates fission (8). Drp1 is largely located in the cytosol. Upon activation, a pool of Drp1 translocates to the mitochondria, where it binds to its mitochondrial adaptors, such as Fis1, Mff or MIEF1, to sever the mitochondrial membranes (11–14). Upon stimulation, mitochondria become fragmented and, importantly, the fragmentation has been demonstrated to contribute to mitochondrial outer membrane permeabilization, ATP depletion, an increase in reactive oxygen species (ROS) and the release of apoptotic factors from the mitochondrial intermembrane space, leading to subsequent cell death (15–18).

Although the majority of LRRK2 is present in the cytoplasm, ∼10% of these proteins are associated with the outer mitochondrial membrane (7), raising the possibility that hyperactivity of LRRK2 mutations might directly affect mitochondrial function. Indeed, flies expressing LRRK2 G2019S display increased sensitivity to rotenone, a mitochondrial complex I inhibitor and a neurotoxin causing Parkinsonism in rodent (19). Fibroblasts harvested from PD patients carrying LRRK2 G2019S exhibited a decrease in both mitochondrial membrane potential (MMP) and intracellular ATP levels (20,21). Further, overexpression of LRRK2 G2019S resulted in mitochondrial fragmentation due to an increase in Drp1 association with the mitochondria (22,23). Moreover, this aberrant mitochondrial morphology induced by the G2019S mutant was accompanied with shorter neurites in both rat primary cortex neurons (24) and human DA neuronal line SH-SY5Y cells (25). Thus, augmented Drp1-mediated mitochondrial fission might play an important role in LRRK2 G2019S-induced neuronal damage. However, how LRRK2 G2019S regulates Drp1 is not yet clear, although protein–protein interaction between LRRK2 and Drp1 has been reported (22,23).

We have recently developed a selective peptide inhibitor of Drp1, P110 (26). P110 specifically inhibited Drp1 hyperactivation and blocked its association with the mitochondria in cultured SH-SY5Y cells exposed to MPP⁺, a neurotoxin causing Parkinsonism (27,28). Moreover, treatment with P110 in rat primary midbrain DA neurons reduced mitochondrial fragmentation and neuronal cell death induced by MPP⁺ (26). In the present study, we examined whether inhibition of Drp1-induced excessive mitochondrial fission by P110 reduces neurodegeneration caused by LRRK2 G2019S.

RESULT

Inhibition of Drp1 by P110 reduced LRRK2 G2019S-induced mitochondrial fragmentation

Accelerated translocation of Drp1 to the mitochondria leads to excessive mitochondrial fission and fragmentation and causes mitochondrial dysfunction (9,29). Consistent with previous studies (21–23), we found that the levels of Drp1 associated with the mitochondria were increased in HEK293 T cells expressing LRRK2 wild-type (WT) and were much more enhanced in cells with the G2019S mutant. In contrast, translocation of Drp1 to the mitochondria was abolished by treatment with a Drp1 peptide inhibitor, P110 (Fig. 1A and B). In parallel, we examined Drp1 localization on the mitochondria in dermal fibroblasts of three PD patients carrying the LRRK2 G2019S mutation. These patient cells enabled us to determine the effects of an endogenous LRRK2 mutation on mitochondrial morphology. Confocal imaging analysis revealed increased localization of Drp1 on the mitochondria in the three PD patient fibroblasts carrying LRRK2 G2019S, whereas Drp1 slightly overlapped with the mitochondria in normal adult fibroblasts (Fig. 1C). There was no significant change observed in healthy fibroblasts treated with P110. Treatment with P110 in these patient fibroblasts significantly reduced localization of Drp1 on the mitochondria (Fig. 1C).

Figure 1.

Treatment with P110 blocked Drp1 association with the mitochondria induced by LRRK2. (A) HEK293 T cells were transfected with LRRK2 or G2019S and treated with peptide P110 or control peptide TAT (1 µm for each). After 16 h of transfection, mitochondrial fractions were isolated, and the Drp1 level was detected by immuno-blotting (IB) with anti-Drp1 antibody. Valtage-dependent anion channel (VDAC) was used as a loading control. (B) A histogram of Drp1 translocation to the mitochondria. The data were expressed as means ± SD from three independent experiments. *P < 0.05 versus non-transfected control (mock); ^#P < 0.05 versus TAT-treated group. (C) PD patient fibroblasts with LRRK2 G2019S mutation were treated with P110 (1 µm/day for 4 days), and cells were then stained with anti-Drp1 (red) and anti-Tom20 (a mitochondrial marker, green) antibodies. Three regions of each sample were randomly captured by a confocal microscope. PD1: ND31960 from 47-year-old female patient; PD2: ND33995 from 53-year-old male patient; PD3: Huf6 from 60-year-old female patient. Normal fibroblast (Nor): human dermal fibroblast adult (HDFa) from a normal adult. Histogram: Drp1 localization on mitochondria was quantified with Olympus Fluoview FV1000 software. At least 100 cells/per group were analyzed. The data were presented as Pearson co-efficiency/cell and expressed as means ± SD from three independent experiments. *P < 0.05 versus normal fibroblast; ^#P < 0.05 versus mock-treated group. Scale bar is 5 µm.

Drp1 recruitment to the mitochondria is a critical step for mitochondrial fission; we examined effects of the G2019S mutant on mitochondrial morphology. HeLa cells exhibited an interconnected mitochondrial network and have been used as a cell culture model to observe the changes in the mitochondrial web (30–32). Overexpression of the G2019S mutant in HeLa cells resulted in ∼53% of cells with fragmented mitochondria, as evidenced by small and spherical mitochondrial clusters (Fig. 2A). The number of fragmented mitochondria was greatly reduced by treatment with P110 (from 53 to 18% relative to that in the mock group). Further, we observed extensive mitochondrial fragmentation in the three PD patient fibroblast lines carrying the LRRK2 G2019S mutation, which was reduced by treatment with P110 (1 µm/day for 4 days) (Fig. 2B). Note that treatment with P110 had no obvious effects on mitochondrial morphology under normal conditions (Fig. 2B), which was consistent with our previous observation in cultured neuronal cells (26). Thus, our data showed that LRRK2 G2019S mutant caused abnormal mitochondrial morphology by recruiting Drp1 to the mitochondria. More importantly, a selective peptide inhibitor of Drp1, P110, reduced excessive mitochondrial fragmentation in cells expressing LRRK2 G2019S exogenously or endogenously.

Figure 2.

P110 reduced mitochondrial fragmentation induced by LRRK2 G2019S. (A) HeLa cells were transfected with LRRK2 G2019S and the cells were then treated with TAT or P110 (1 µm, each). The cells were stained with anti-Tom20 antibody after a 24-h transfection. (B) PD patient fibroblasts with LRRK2 G2019S mutant were treated with P110 (1 µm/day for 4 days) and then stained with anti-Tom20 antibody. The number of cells with fragmented mitochondria was counted, and the data were expressed as means ± SD from three independent experiments. At least 200 cells/group were counted. *P < 0.05 versus non-transfected (mock) control or normal fibroblast (Nor); ^#P < 0.05 versus G2019S-transfected cells or patient fibroblasts. Scale bar is 5 µm.

Drp1 is required for LRRK2 G2019S-induced mitochondrial dysfunction

Mitochondrial dysfunction and oxidative stress have been implicated in the pathogenesis of PD (33,34). To assess the effect of P110 on LRRK2 G2019S-induced mitochondrial damage, we first determined the MMP and mitochondrial superoxide production. Overexpression of either LRRK2 WT or the G2019S mutant caused a dramatic loss of the MMP when compared with that in non-transfected cells (Fig. 3A). The anti-apoptotic Bcl2 family proteins Bcl-2 and Bcl-xL, which maintain the MMP, were decreased. As a result, cytochrome c was released from the mitochondria (Fig. 3B). In contrast, the cells treated with P110 under the same conditions not only kept the MMP, but also suppressed the decrease in the protein levels of Bcl-2 and Bcl-xL (Fig. 3B), suggesting an improvement of mitochondrial membrane integrity.

Figure 3.

Inhibition of Drp1 reduced LRRK2 G2019S-induced mitochondrial dysfunction. HeLa cells were transfected with LRRK2 or G2019S followed by treatment with TAT or P110 (1 µm, each). (A) 20 h after transfection, cells were stained with TMRM (0.25 µm). Three randomly picked regions of each sample were captured by microscope, and the density of TMRM fluorescence was expressed as means ± SD from three independent experiments. *P < 0.05 versus non-transfected control; #, P < 0.05 versus TAT-treated group. (B) 24 h after transfection, levels of Bcl-2, Bcl-xL and cytochrome c were examined in mitochondrial fractions by immunoblotting (IB); VDAC was used as a loading control. The protein levels of Bcl-2, Bcl-xL and cytochrome c were quantified (means ± SD; from three independent experiments). *, P < 0.05 versus non-transfected control; ^#P < 0.05 versus TAT-treated group. (C) Twenty-four hours after transfection, the protein levels of complex I–V of the mitochondrial electron transport chain were determined by IB using the MitoProfile^® Total OXPHOS Rodent WB Antibody Cocktail (Abcam, USA). Among five complexes, only protein levels in complex I and IV were changed. The levels of complex I and IV were then expressed as means ± SD from three independent experiments. *P < 0.05 versus non-transfected control; ^#P < 0.05 versus TAT-treated group. (D) Twenty hours after transfection, HeLa cells were stained with 5 µm MitoSox™ and 100 nm mitotracker for 10 min. The density of MitoSox™ fluorescence was expressed as means ± SD from three independent experiments. *P < 0.05 versus non-transfected control; ^#P < 0.05 versus TAT-treated group. Right panel: representative images of mitoSOX staining (red) and mitotracker (green). Fibroblasts either from a normal adult (Nor) or PD patients with the G2019S (PD1, PD2 and PD3) were treated with P110 (1 µm/day for 4 days) and then stained with TMRM (0.25 µm) (E) and 5 µm MitoSOX with 100 nm mitotracker (F). The density of fluorescence was quantified and expressed as means ± SD from three independent experiments. *P < 0.05 versus normal fibroblasts (Nor); #, P < 0.05 versus TAT-treated patient fibroblasts. Scale bar is 10 µm.

Assembly of the electron transport chain (ETC) complex depends on inner mitochondrial membrane integrity, which is maintained by fusion and fission processes (35), and deficits in the components of the ETC complex ultimately increases ROS production (36). We examined protein levels of the components of ETC complex I–V in cells expressing LRRK2 WT or the G2019S mutant. Protein levels of complex I and IV were significantly decreased when compared with those in non-transfected cells. In contrast, P110 treatment corrected these defects (Fig. 3C). Further, the production of mitochondrial superoxide, a major source of mitochondrial reactive oxygen species (mitoROS), was greatly elevated by ∼27-fold in cells expressing LRRK2 G2019S, which was abolished by treatment with P110 (Fig. 3D).

Consistent with previous studies (22,23), we found that the expression of LRRK2 WT caused disruption of mitochondrial function (Fig. 3A–C). To limit the possibility that these mitochondrial changes from LRRK2 and the G2019S mutant might be resulted from overexpression of proteins, we in parallel used patient fibroblasts to determine the effects of endogenous LRRK2 G2019S on mitochondrial functions. Similar to our above results (Fig. 3A and D), in the fibroblasts of PD patients carrying the LRRK2 2019S mutation, the loss of the MMP and increase in mitoROS in all three PD patient fibroblast lines were corrected by treatment with P110 (Fig. 3E and F). Thus, Drp1 hyperactivation might play an important role in LRRK2 G2019S-induced mitochondrial impairment.

LRRK2 G2019S exacerbated Drp1-dependent autophagy

Overexpression of LRRK2 G2019S in SH-SY5Y neuroblastoma cells led to an increase in the content of autophagosomes (25,37). Mice with the human LRRK2 G2019S transgene exhibited increased density of autophagic vacuoles in the brain cortex and striatum (24). These studies suggest that autophagy plays an active role in cellular toxicity induced by LRRK2 G2019S. Given that inhibition of Drp1 by a dominant-negative mutant Drp1^K38A reduced autophagy (38,39), we next examined whether LRRK2 G2019S leads to cellular damage via Drp1-dependent autophagy.

In HEK293T cells expressing LRRK2 WT or the G2019S mutant, a conversion of microtubule-associated protein light chain 3, LC3, (LC-I to LC3-II, a marker of autophagy) was evident; the level of LC3-II was significantly increased which was suppressed by either treatment with P110 or expression with Drp1^K38A (Fig. 4A). We found that knockdown of LRRK2 by silence RNA technique partially reduced the protein level of LC3-II in human normal fibroblasts (Supplementary Material, Fig. S1A). Together with the previous study showing that depletion of LRRK2 in mice led to a decreased LC3-II level in the kidney (40), these findings suggest a role of LRRK2 in regulation of autophagic process. However, the detailed mechanism remains to be examined. In addition, the activity of lysosomes measured by Lyso-ID Red dye was elevated by 5-fold in LRRK2 G2019S-expressing cells, and inhibition of Drp1 via either P110 treatment or expression of Drp1^K38A reduced this hyperactivity (Fig. 4B). These data indicate that inhibition of Drp1 reduces the excessive autophagy induced by LRRK2 G2019S.

Figure 4.

P110 treatment reduced LRRK2 G2019S-induced excessive autophagy. (A) Left panel: HEK293 T cells were transfected with LRRK2 or G2019S followed by treatment with TAT or P110 (1 µm, each). In parallel, cells were co-transfected with Drp1^K38A (a dominant-negative mutant of Drp1) and LRRK2 or G2019S. Thirty hours after transfection, LC3 I and LC3-II were determined by anti-LC3 antibody. Actin was used as a loading control. The level of LC3-II was quantified and expressed as means ± SD from three independent experiments. *P < 0.05 versus non-transfected control (mock); ^#P < 0.05 versus TAT-treated group with LRRK2 or G2019S. (B) HeLa cells were transfected with the indicated plasmids. Twenty hours after transfection, cells were stained with Lyso-ID Red dye. Three randomly picked regions of each sample were captured by microscope and the density of lyso-ID red was expressed as means ± SD from three independent experiments. *P < 0.05 versus non-transfected control (mock); ^#P < 0.05 versus TAT-treated group. (C) HEK293 T cells were transfected with LRRK2 or G2019S followed by treatment with TAT (1 µm), P110 (1 µm) or BFA (20 nm). Twenty-four hours after transfection, the activity of citrate synthase was determined using total cell lysates to indicate the mitochondrial mass. The activity of citrate synthase was calculated and compared with non-transfected cells (means ± SD; from three independent experiments). *P < 0.05 versus non-transfected control (mock); ^#P < 0.05 versus TAT-treated group with LRRK2 or G2019S. (D) HEK293 T cells were transfected with LRRK2 or G2019S followed by transfection with Myc-Parkin. At the indicated groups, the Myc-Parkin level on the mitochondria was determined by IB with anti-Myc antibodies. VDAC was used as a loading control. Histogram: the level of Myc-Parkin was quantified and expressed as means ± SD from four independent experiments. *P < 0.05 versus non-transfected control; ^#P < 0.05 versus TAT-treated group with LRRK2 or G2019S. Fibroblasts from a normal adult (Nor) or PD patient carrying LRRK2 G2019S were treated with P110 (1 µm/day for 4 days). Cells were stained with Lyso-ID Red dye to determine the lysosome activity. (E) Autophagic flux was assessed by the quantification of LC3-II levels in the presence or absence of BFA (F), and the citrate synthase activity was determined to reflect the mitochondrial mass (G). The data were expressed means ± SD from three independent experiments. *P < 0.05 versus normal fibroblast (Nor); ^#P < 0.05 versus PD patient fibroblasts treated with TAT. Scale bar is 10 µm.

Further, treatment with P110 improved the activity of citrate synthase, which reflects the mitochondrial mass (41), in cells expressing LRRK2 WT or the G2019S mutant (Fig. 4C). Treatment with bafilomycin A1 (BFA), which prevents maturation of autophagic vacuoles by inhibiting fusion between autophagosomes and lysosomes (42), also corrected the loss of the mitochondrial mass (Fig. 4C). Parkin recruitment to the mitochondrial outer membrane represents a critical step in mitochondria-associated autophagy (mitophagy) (43). In HEK293T cells co-expressing Myc-Parkin with LRRK2 WT or the G2019S mutant, a great increase in Parkin association with the mitochondria was observed when the cells were treated with BFA, whereas the accumulation of Parkin on mitochondria was blocked by treatment of P110 under the same conditions (Fig. 4D). However, in the absence of LRRK2, Parkin was still recruited to the mitochondria of cells exposed to FCCP (Supplementary Material, Fig. S1B), suggesting that LRRK2 is not required for Parkin translocation to the mitochondria. Thus, we reasoned that P110 treatment corrected the G2019S-induced mitochondrial damage, such as mitochondrial depolarization, which in turn prevented Parkin recruitment and subsequent mitochondria-related degradation.

Next, in LRRK2 G2019S PD patient fibroblasts, we found that the activity of lysosomes was dramatically increased by 27-, 15- and 12-fold in three lines of PD patient fibroblasts, relative to that in normal fibroblasts, which was consistently suppressed by treatment with P110 (Fig. 4E). Moreover, treatment with P110 inhibited enhanced autophagic flux and increased mitochondrial mass (Fig. 4F and G). Together, the above data suggest that Drp1 hyperactivation mediate LRRK2 G2019S-induced excessive mitochondria-associated autophagy.

Threonine595 phosphorylation of Drp1 by LRRK2 G2019S is required for Drp1-mediated mitochondrial fragmentation and excessive autophagy

How does LRRK2 G2019S activate Drp1? Consistent with previous studies (22,23), we found that both LRRK2 WT and the G2019S mutant interacted with Drp1 (Fig. 5A). The interaction was abolished in the cells expressing LRRK2 3XKD (a kinase-dead form of LRRK2) (Fig. 5A), suggesting that this interaction was dependent on LRRK2 kinase activity. Given that LRRK2 G2019S augments LRRK2 serine/threonine kinase activity (3,4,7) and that Drp1’s post-translational modification leads to mitochondrial fragmentation (18,22,23,44–46), we set out to determine whether Drp1 could be a target of LRRK2 for phosphorylation.

Figure 5.

Drp1 Thr595 phosphorylation is required for LRRK2 G2019S-induced mitochondrial damage and excessive autophagy. (A) HEK293 T cells were transfected with LRRK2 or G2019S. Total cell lysates were harvested after 16 h of transfection. Immunoprecipitation (IP) was carried out with anti-Drp1 antibodies followed by IB with anti-LRRK2 antibody. Input: 10% of protein. (B) Recombinant human GST-Drp1 was incubated with recombinant WT and mutant LRRK2 protein. Phosphor-LRRK2 and phosphor-Drp1 were examined by IB with anti-phosphor-threonine antibodies. (C) Mass spectrometry analysis identified Thr595 as a phosphorylation site of Drp1 by LRRK2 G2019S. Domain structure and sequence alignment of Drp1 indicated the location of the Thr595 phosphorylation site. GTP, GTPase domain; MID, middle domain; VD, variable domain; GED, GTPase effector domain. (D) HeLa cells were transfected with G2019S or 3XKD and treated with TAT or P110 (1 µm, each). Total cell lysates were subjected to IP with anti-Drp1 antibody followed by IB with the indicated antibodies. Histogram: quantification of Drp1 Thr-phosphorylation was expressed as means ± SD from three independent experiments. *P < 0.05 versus non-transfected control (mock); #, P < 0.05 versus the G2019S transfected group. HeLa cells were co-transfected with Drp1^T595A or Drp1^T595D with G2019S. (E) Twenty-four hours after transfection at the indicated group, cells were stained with anti-Tom20 antibodies. The number of cells with fragmented mitochondria was counted, and the data were expressed as means ± SD from three independent experiments. At least 200 cells/group were counted. (F) Total lysates were subjected with IB with anti-LC3 antibodies. Quantitation was provided in a histogram. The data were expressed as means ± S.D. of three independent experiments. (G) Cells were stained with Lyso-ID Red dye. Nuclei were stained with Hoechst (blue). Right panel: representative images. Scale bar is 5 µm. Left panel: the quantification of the Lyso-ID red fluorescence from three independent experiments was provided in a histogram (means ± SD of three independent experiments). (H) The mitochondrial mass was determined by the activity of citrate synthase as shown above. The data were expressed means ± SD from three independent experiments. *P < 0.05 versus non-transfected control (mock); ^#P < 0.05 versus the G2019S transfected group.

Recombinant human Drp1 (GST-Drp1) was subjected to an in vitro phosphorylation assay with recombinant human LRRK2 [GST-tagged wide-type, G2019S mutant, and D1994A mutant (kinase-dead) protein consisting of all the functional domains (amino acid residues 970–2527)]. We found that both LRRK2 and the G2019S mutant were auto-phosphorylated and that the G2019S mutant predominantly phosphorylated Drp1, when antibodies specific to recognize phosphor-threonine were used (Fig. 5B). Again, a kinase dead mutant (D1994A) abolished this phosphorylation. Note that we did not observe increased serine-phosphorylation of Drp1 under the same conditions. Further, mass spectrometry analysis revealed one conserved site, Threonine595 (Thr595), as a target of LRRK2 G2019S-dependent phosphorylation (Supplementary Material, Fig. S2). This Thr site is highly conserved among mammalian species and is located in the variable domain (VD) of Drp1 (Fig. 5C). In HeLa cells expressing LRRK2 G2019S, we confirmed that endogenous Drp1 was Thr-phosphorylated, whereas the phosphorylation was abolished in cells expressing LRRK2 3XKD (kinase dead form of LRRK2) (Fig. 5D). More importantly, the Drp1 peptide inhibitor P110, completely suppressed Drp1 Thr-phosphorylation when compared with that in the group treated with control peptide carrier TAT (Fig. 5D). Currently, the possibility that P110 interferes with Drp1 conformational changes, and in turn affects Drp1’s phosphorylation, is under investigation.

To further determine the functional importance of Thr595, we generated a Thr595 to alanine mutant of Drp1 (Drp1^T595A, non-phosphorylated) and to aspartic acid mutant (Drp1^T595D, phosphomimetic). We then transfected these phosphor-mutants of Drp1 to the HeLa cells in the presence of LRRK2 G2019S and observed mitochondrial morphology. When the cells with LRRK2 G2019S were co-transfected with Drp1^T595A, the broken mitochondrial web induced by the G2019S was corrected and mitochondrial network exhibited interconnected and elongated (Fig. 5E). Quantification analysis revealed that the expression of Drp1^T595A reduced the number of cells with the fragmented mitochondria from 53 to 15% relative to that in the mock-treated group (Fig. 5E). Drp1^T595D co-expressing with the G2019S increased mitochondrial fragmentation, the extent of which was similar to that of cells expressing the G2019S alone. Note that Drp1^T595D alone did not increase mitochondrial fragmentation in the absence of LRRK2 G2019S. This finding is consistent with the previous studies showing Drp1 phosphor-mimetic mutant Drp1 Ser616D per se did not change mitochondrial morphology (47). Consistently, in two lines of PD patient fibroblasts carrying the G2019S mutant, the expression of Myc-Drp1^T595A corrected disrupted mitochondrial network; the number of cells with the fragmented mitochondria was dramatically reduced, when compared with that in the normal fibroblast-expressing control vector (Fig. 6A). Together, our findings showed that Drp1 Thr595 phosphorylation mediated LRRK2 G2019S-induced mitochondrial fragmentation.

Figure 6.

Expression of Drp1^T595A reversed mitochondrial fragmentation and reduced augmented autophagy in PD patient fibroblasts carrying LRRK2 G2019S. Normal fibroblast (Nor, HDFa) and PD patient fibroblasts (PD1 and PD2) were transfected with either Myc-vector (Con Vec) or Myc-Drp1^T595A for 5 days using TransIT®-2020 Transfection Reagent (Mirus Bio LLC, Madison, WI, USA), according to manufacturer's instruction. (A) Fibroblasts were stained with anti-Tom20 (green) and anti-Myc (red) antibodies at the indicated groups. Left: representative images showing mitochondrial network. Scale bar is 5 µm. Insert: fibroblasts co-expressing Myc-vector or Myc-Drp1^T595A. Right top: western blot analysis confirmed the expression of Myc-Drp1^T595A in the fibroblasts with the indicated antibodies. Right left: the number of Myc-expressing cells with the fragmented mitochondria was counted, and the data were expressed as means ± SD from three independent experiments. At least 50 cells/group were counted. (B) Fibroblasts were stained with Lyso-ID Red dye as above. The quantification of the Lyso-ID red fluorescence from three independent experiments was provided in a histogram (means ± SD). (C) The mitochondrial mass was determined by measuring the fluorescent density of Mitotracker Green. The data were expressed as means ± SD from three independent experiments. *P < 0.05 versus normal fibroblast (Nor) with control vector; ^#P < 0.05 versus PD fibroblasts with control vector.

We next determined how Drp1 phosphorylation at Thr595 affects LRRK2 G2019S-induced autophagy. Co-transfection of Drp1^T595A with the G2019S mutant suppressed LC3 cleavage, which was abolished by co-transfection of Drp1^T595D with the G2019S (Fig. 5F). Moreover, the increased lysosomal activity (Fig. 5G) and loss of the mitochondrial mass (Fig. 5H) induced by LRRK2 G2019S were corrected in the cells in which Drp1^T595A was co-expressed, when compared with cells with co-expression of Drp1^T595D. Further, the expression of Myc-Drp1^T595A reduced lysosomal hyperactivity (Fig. 6B) and increased mitochondrial mass (Fig. 6C) in two lines of PD patient fibroblasts carrying LRRK2 G2019S. Collectively, these data suggest that Drp1 phosphorylation at Thr595 is required for LRRK2 G2019S-induced mitochondrial and cellular pathology.

Inhibition of Drp1 hyperactivation by P110 reduced mitochondrial and neuronal abnormalities in dopaminergic neuron derived from PD patient-induced pluripotent stem cells (iPS cells)

PD features the selective neurodegeneration of DA neurons. To further examine whether Drp1 hyperactivation contributes to LRRK2 G2019S-induced DA neuronal damage, we generated iPS cells derived from fibroblasts of PD patients carrying the LRRK2 G2019S mutation. We generated two lines of LRRK2 G2019S-iPS cells and two lines of iPS cells from normal adult fibroblast (Con-iPS cells). Of those, one line per subject was characterized and shown to be fully reprogrammed, as judged by colony morphology and sustained growth for long-term passaging (>30 passages) (Supplementary Material, Fig. S3). These iPS cell lines express antigens of embryonic stem cells, including OCT3/4, SOX2, SSEA-4, TRA-1-60 and TRA-1-81, and were positive in alkaline phosphatase (AP) staining (Supplementary Material, Fig. S3). Moreover, the iPS cell lines showed in vitro pluripotency and were able to differentiate into cells immunopositive for three embryo germ layer markers, α-SMA (mesoderm), βIII-Tubulin (ectoderm) and GATA-4 (endoderm) (Supplementary Material, Fig. S3).

We directly differentiated these iPS cell lines into DA neurons using the protocol (48) with some modifications. DA neuron generation was assessed by co-staining with the neuron-specific class III β-tubulin (Tuj1) and tyrosine hydroxylase (TH, a marker of DA neuron, see Fig. 8C). At 30 days following differentiation, we started to treat these neurons with a Drp1 peptide inhibitor P110 (1 µm/day for five consecutive days). In neurons derived from the LRRK2 G2019S-iPS cells, mitochondrial morphology along the branches of patient neurons immunopositive for anti-TH antibody was greatly altered, showing less mitochondria and longer distances between them. P110 treatment significantly improved the length of mitochondria along patient DA neuronal branches from 36 to 84%, relative to that of DA neurons from Con-iPS cells (Fig. 7A and B), indicating a reduction in mitochondrial fragmentation. Moreover, in patient neurons, a loss of MMP, increased mitoROS and decreased ATP levels as well as lysosome hyperactivity were evident (Figs 7C, E, F and 8A and B), which were corrected by a 5-day treatment with P110. In parallel, in DA neurons derived from LRRK2 G2019S-iPS cells, expression of Myc-Drp1^T595A for 7 days not only improved mitochondrial length along neuronal branches (Fig. 7D), but also corrected lower MMP, increased mitoROS and lysosomal hyperactivity (Figs 7E, F and 8G). Further, in agreement with the previous studies (49), the surviving DA neurons with the LRRK2 G2019S mutation displayed evidently morphological alterations, including fewer and shorter neurites compared with those from Con-iPS cells. In contrast, in patient DA neurons treated with either P110 or Myc-Drp1^T595A, the neurite length was increased significantly (from 50 to 84% in DA neurons treated with P110 and from 45 to 70% in DA neurons expressing Myc-Drp1^T595A, respectively) (Fig. 8C–F). In addition, treatment with NH₄Cl (ammonium chloride, an inhibitor to inhibit lysosome activity) partially increased neurite length of DA neurons derived from LRRK2 G2019S-iPS cells (Supplementary Material, Fig. S4), confirming that impaired autophagy-lysosome pathway contributes to neurodegeneration of DA neurons in PD (50,51). Together, these results support our findings (Figs 1–4), validating an important role of Drp1 hyperactivation in LRRK2 G2019S-induced mitochondrial and neuronal defects.

Figure 7.

Inhibition of Drp1 hyperactivation reduced mitochondrial fragmentation and damage in neurons derived from LRRK2 G2019S patient-iPS cells. DA neurons (Con or LRRK2 G2019S) were differentiated from iPS cells derived from normal fibroblasts (Con-iPS cell) and PD patient fibroblasts carrying LRRK2 G2019S (LRRK2 G2019S-iPS cell), respectively. At 30 days following differentiation, cells were treated with P110 (1 µm/day for 5 days). (A) Cells were stained with anti-TOM20 (a marker of mitochondria) and anti-TH (a marker of DA neuron). The bottom panels are enlarged images of boxed areas in each middle panel. Scale bar: 10 µm. (B) The quantitation of mitochondrial length along the neurites of DA neurons (immunopositive for TH) is provided in a histogram as means ± SE of three independent experiments. The NIH Image J software was used for the quantification. At least 30 DA neurons per group were analyzed. (C) Total ATP levels were measured using total lysates of mixed neuronal cells derived from Con- or LRRK2 G2019S-iPS cells. Data are presented as means ± SE of three independent experiments. (D) At 28 days following differentiation, cells were transfected with either Myc-vector or Myc-Drp1^T595A using TransIT®-2020 Transfection Reagent. Seven days after transfection, cells were stained with anti-TOM20 (green) and anti-Myc (red) antibodies. The middle panels are enlarged images of boxed areas in each upper panel. The insert confirmed the neuron immunopositive for anti-Myc antibody. Scale bar: 10 µm. Western blot analysis confirmed the expression of Myc-Drp1^T595A in the neurons derived from iPS cells with the indicated antibodies. Lower right: quantitation of mitochondrial length along the neurites of neurons (immunopositive for anti-Myc antibody) is provided in a histogram as means ± SE of three independent experiments. The NIH Image J software was used for the quantification. At least 30 DA neurons per group were analyzed. *P < 0.05 versus neurons derived from Con-iPS cells with control vector (Myc-vector); ^#P < 0.05 versus neurons derived from the G2019S-iPS cells with control vector (Myc-vector). (E) Neuronal cells derived from Con- and LRRK2 G2019S-iPS cells were labeled with TMRM fluorescence dye to indicate mitochondrial membrane potential (MMP). Density of TMRM red fluorescence only in cells with a neuronal-like morphology (multipolar cell bodies with at least two processes) was further quantitated by the NIH Image J software. (F) Neuronal cells from Con-and LRRK2 G2010S-iPS cells were stained with mitoSOX red (measurement of mitoROS). The density of mitoSOX red fluorescence was further analyzed. At least 50 neurons per group were analyzed. (*P < 0.05 versus neurons derived from Con-iPS cells; #P < 0.05 versus neurons derived from the G2019S-iPS cells.)

Figure 8.

Inhibition of Drp1 hyperactivation corrected lysosome abnormality and reduced neurite shortening in DA neurons derived from PD patient-iPS cells. DA neurons were derived from either Con-iPS cells or PD patient LRRK2 G2019S-iPS cells. At 30 days following differentiation, cells were treated with P110 (1 µm/day for 5 days). (A) Cells were stained with anti-LAMP2 (red) and anti-TH (green). Density of LAMP2 fluorescence in cells expressing TH was quantitated and expressed as means ± SE of three independent experiments. (B) Lysosome activity was measured by Lyso-ID Red dye staining. The quantification of the red fluorescence in cells with neuron-like morphology was provided in a histogram (means ± SE of three independent experiments). (C) Representative images on DA neurons derived from Con- and LRRK2 G2019S-iPS cells at the indicated groups. The DA neurons were confirmed by co-staining with anti-Tuj1 antibody (red, a mature neuronal marker) and anti-TH antibody (green). Upper panel shows a single neuron from each experimental group (×40); middle panel shows the corresponding imaging in black and white; lower panel shows imaging that contains a cluster of neurons (×20). Nuclei were stained with Hoechst (blue). Scale bar is 5 µm. (D) The neurite length of DA neurons is provided in a histogram as means ± SE of three independent experiments. At least 50 neurons per group were analyzed with the neurite tracer plugin for the NIH Image J software. *P < 0.05 versus neurons derived from Con-iPS cells; ^#P < 0.05 versus neurons derived from the G2019S-iPS cells. (E) At 28 days following differentiation, cells were transfected with either Myc-vector or Myc-Drp1^T595A using TransIT®-2020 Transfection Reagent. Seven days after transfection, cells were stained with anti-TH (green) and anti-Myc (red) antibodies. The bottom panels confirmed the neuron immunopositive for anti-Myc antibody. Scale bar is 5 µm. (F) The neurite length of DA neurons expressing Myc-Drp1^T595A is provided in the histogram as means ± SE of three independent experiments. At least 30 neurons per group were analyzed with the neurite tracer plugin for the NIH Image J software. (G) Lysosome activity was measured by Lyso-ID Red dye staining in neurons transfected with either Myc-vector or Myc-Drp1^T595A. The quantification of the red fluorescence in cells with neuron-like morphology was provided in a histogram (means ± SE of three independent experiments). *P < 0.05 versus neurons derived from Con-iPS cells with control vector (Myc-vector); ^#P < 0.05 versus neurons derived from the G2019S-iPS cells with control vector (Myc-vector).

DISCUSSION

In this study, we used P110, a selective peptide inhibitor of Drp1 that we recently developed (26), to elucidate the role of Drp1-mediated mitochondrial fission in mitochondrial and neuronal pathology induced by the LRRK2 G2019S mutation. We have demonstrated that (i) the G2019S mutant caused Drp1-mediated mitochondrial dysfunction via increasing the recruitment of Drp1 to the mitochondria; (ii) the G2019S exacerbated autophagy, which was dependent on Drp1 activation; (iii) the G2019S bound to and phosphorylated Drp1 at Thr595, which led to mitochondrial fragmentation and subsequent excessive autophagy. These defects caused by LRRK2 G2019S were suppressed by either treatment with P110 or expression of Drp1 phosphor mutant (Drp1^T595A), suggesting that Drp1 hyperactivation is an important element in the G2019S-induced cellular pathology. The neuro-protective efficacy of P110 treatment was further validated in DA neurons derived from patient LRRK2 G2019S-iPS cells.

A disturbance of mitochondrial dynamics was recently highlighted in both sporadic and genetic PD models (23,52–55). The G2019S, the most common mutation of LRRK2, leads to increased kinase activity and promotes neuronal toxicity, indicating that the G2019S may play a pathogenic role through a ‘gain-of-function’ mechanism (7,56,57). LRRK2 has been reported to interact with the fission protein Drp1, which increases mitochondrial fragmentation (22,23). In the present study, in addition to the interaction between LRRK2 and Drp1, we identified that the mutant G2019S activated Drp1 by specific phosphorylation at Thr595. To our knowledge, this is the first report that Drp1 can be phosphorylated at a threonine site besides Ser 616 by protein kinase C delta (18) and CDK1(46) and Ser 637 by protein kinase A (10,45,47,58). A site-directed mutant (Drp1^T595A) abolished the G2019S-induced disruption of mitochondrial integrity, suggesting that Thr595 phosphorylation of Drp1 by LRRK2 G2019S mediated the mutant-induced mitochondrial dysfunction. Therefore, our studies may suggest a new target of LRRK2 in the context of PD. So far, all identified phosphorylation sites of Drp1 are located in the VD of this protein, emphasizing the importance of this domain in regulating Drp1 activation. Strack and Cribbs found that post-translational modifications of Drp1 in or near the VD alter the conformation of a membrane-proximal oligomerization interface to influence Drp1 assembly rate (59). It is thus possible that Drp1 Thr595 phosphorylation by G2019S changed Drp1’s conformation of self-assembly, which in turn increased its fission activity. However, this possibility remains to be determined. In addition, we noticed that inhibition of Drp1 by P110 treatment attenuated mitochondrial defects caused by LRRK2 WT which does not phosphorylate Drp1 at Thr595. It remains to determine whether additional mechanism is involved in LRRK2-induced Drp1 activation.

A concomitant occurrence of mitochondrial morphology changes and excessive autophagy was documented in various models of LRRK2 G2019S. The SH-SY5Y neuronal cell line expressing the G2019S mutant showed a loss of MMP, which was associated with mitochondrial fragmentation and shorter neurites with increased autophagic vacuoles (22,23). Fibroblasts from PD patients with the G2019S mutation exhibited mitochondrial dysfunction and higher autophagic activity through activation of the MEK/ERK pathway (21,60,61). G2019S-PD-specific iPS cells-derived neurons show accumulation of autophagic vacuoles (49) and mitochondrial damage (62). LRRK2 G2019S transgenic mice exhibited abnormal, condensed, small and fragmented mitochondria in autophogosomes in the cortex and striatum of the brain (24). Together, these lines of evidence suggest a link between mitochondrial fission and autophagy, even though the underlying mechanism is not yet clear. In fact, mitochondrial fission is a prerequisite for autophagy (38,63). Electron microscope tomography shows that fission events can yield asymmetric daughter mitochondria that differ in membrane potential (64). Drp1-induced mitochondrial fission caused LC3 cleavage, which is required for PINK1/Parkin-mediated mitophagy (65), and overexpression of the Drp1^K38A reduces mitochondrial autophagy (38,39). Further, we recently showed that inhibition of Drp1 by P110 suppressed MPP⁺-induced excessive autophagy (26). Thus, mitochondrial fission might constitute a principal route for generating depolarized mitochondria that are later targeted by the autophagy machinery, the process of which might be accelerated by the LRRK2 G2019S mutation. This hypothesis was further supported by our finding that LC3 cleavage, loss of mitochondrial mass and lysosome hyperactivity induced by the LRRK2 G2019S mutation were corrected by inhibition of Drp1 activation using the Drp1 peptide inhibitor P110 or a dominant-negative mutant Drp1^K38A or a phosphor mutant Drp1^595A (Figs 4, 5 and 6).

To date, LRRK2 transgenetic Drosophila and C. elegans showed PD-like phenotypes and overexpression of the G2019S mutant induces more severe phenotypes (66–68). However, transgenic mice expressing LRRK2 pathogenic mutants do not show specific DA neurodegeneration or relative behavior deficits, which might be due to the differences between mouse and human genetic backgrounds (66,69,70). As an alternative, the recent development of PD patient iPS cells offers a unique model in vitro to study neuropathology specifically induced by PD-related disease genes and to assess the potential of drugs for treatment (48,49,62). The neurons derived from LRRK2 G2019S PD iPS cells exhibited mitochondrial damage, elevated autophagic activity and increased neuronal vulnerability to stressors (48,49,71). Therefore, this patient disease-derived neuronal culture recapitulated some neuropathology in the context of patient genotype. In the present study, in agreement with previous studies (48,49,62), we observed mitochondrial and lysosomal dysfunctions, such as loss of MMP, increased mitoROS and lysosome hyperactivity in DA neurons derived from patient LRRK2 G2019S-iPS cells (Figs 7 and 8). Importantly, treatment with the Drp1 inhibitor P110 not only corrected impaired mitochondrial function and lysosome hyperactivity, but also increased the outgrowth of neurites of these patient DA neurons, suggesting that P110 treatment improved the viability of DA neurons via inhibiting the G2019S mutation-induced mitochondrial pathology.

Mitochondrial fission impairment was observed at the early stage of cell damage and occurred before the appearance of symptoms of neurological deficits in some diseased animal models (17,72). Therefore, inhibitors that block pathological condition-induced mitochondrial fission might be attractive strategies to halt or slow down the underlying neuronal degenerative process of diseases such as PD. In this study, we found that treatment with the Drp1 selective peptide inhibitor P110 robustly inhibited LRRK2 G2019S-induced Drp1 hyperactivation-mediated cellular injury. P110 is conjugated with peptide carrier TAT, which is cell permeable and easily passes through the blood–brain barrier (73–75). Therefore, the peptide inhibitor P110 might be useful for the treatment of PD. Our findings also encourage further validation of this therapeutic strategy in animal models of PD.

MATERIALS AND METHODS

Cell culture

Human embryonic kidney cells HEK293T and human cervix carcinoma cells HeLa were maintained in Dulbecco's Modified Eagle Medium supplemented with 10% (v/v) heat-inactivated fetal calf serum and 1% (v/v) penicillin/streptomycin.

LRRK2 G2019S patient fibroblasts (ND31960 from 47-year-old female and ND33995 from 53-year-old male) were purchased from Coriell Institute. Patient fibroblast Huf-6 from 60-year-old female was a generous gift from Dr Renee Pera's lab at Stanford University. Human dermal fibroblast adult (HDFa) was purchased from Invitrogen. These fibroblasts were maintained in minimum essential medium supplemented with 15% (v/v) heat-inactivated fetal calf serum and 1% (v/v) penicillin/streptomycin. All cell cultures were maintained in a humidified atmosphere with 5% CO2 at 37°C.

Isolation of mitochondrial-enriched fraction and lysate preparation

Cells were washed with cold phosphate-buffered saline (PBS) and incubated on ice in lysis buffer (250 mm sucrose, 20 mm HEPES-NaOH, pH 7.5, 10 mm KCl, 1.5 mm MgCl2, 1 mm EDTA, protease inhibitor cocktail, phosphatase inhibitor cocktail) for 30 min. Cells were scraped and then disrupted 10 times by repeated aspiration through a 25-gauge needle, followed by a 30-gauge needle. The homogenates were spun at 800g for 10 min at 4°C and the resulting supernatants were spun at 10 000g for 20 min at 4°C. The pellets were washed with lysis buffer and spun at 10 000g again for 20 min at 4°C. The final pellets were suspended in lysis buffer containing 1% Triton X-100 and were mitochondrial-rich lysate fractions. The mitochondrial protein valtage-dependent anion channel (VDAC) was used as a loading control.

Immunoprecipitation

Cells were lysed in total cell lysate buffer (50 mm Tris–HCl, pH 7.5, containing 150 mm NaCl, 1% Triton X-100 and protease inhibitor). Soluble protein was incubated with the indicated antibody overnight at 4°C and protein A/G beads for 1 h. Immunoprecipitates were washed three times with cell lysate buffer and analyzed by SDS–PAGE and immunoblotting with antibodies.

Immunocytochemistry

Cells cultured on coverslips were washed with cold PBS, fixed in 4% formaldehyde and permeabilized with 0.1% Triton X-100. After incubation with 2% normal goat serum (to block non-specific staining), fixed cells were incubated overnight at 4°C with antibodies against Drp1 (1:500, BD bioscience, USA) and Tom20 (1:500, Santa Cruz Biotechnology, USA). Cells were washed with PBS and incubated for 60 min with FITC-labeled goat anti-rabbit antibody and rhodamine-labeled goat anti-mouse antibody (1:500, Invitrogen, USA), followed by incubation with Hoechst dye (1:10 000, Invitrogen, USA) for 10 min. Coverslips were mounted and slides were imaged by confocal microscopy (Olympus, Fluoview FV100). To determine mitochondrial superoxide production in cultures, cells were incubated with 5 µm MitoSOX™ red mitochondrial superoxide indicator (Invitrogen) for 10 min at 37°C. To measure the membrane potential of mitochondria in cultures, cells were incubated with 0.25 µm tetramethylrhodamine (TMRM) (Invitrogen) for 20 min at 37°C. To determine lysosomal activity, cells were incubated with Lyso-ID Red dye (Enzo, USA) for 30 min at 37°C. The staining was imaged by microscope, and quantification was carried out using the NIH Image J software.

Citrate synthase activity assay

The cells were harvested, and 50 µg of lysate was added to each well of a 96-well plate containing 160 µl assay buffer (60 mm Tris–HCl, pH 7.5, 200 µm Acetyl CoA, and 250 µm DTNB). The lysate was incubated for 5 min, and then 20 µl of 2 mm oxaloacetate was added to each well. OD was measured at 412 nm for 30 min using a spectrometry plate reader (spectraMAX340, Molecular Devices, USA). The rate of citrate synthase activity was quantified relative to control cells.

In vitro phosphorylation of Drp1 by LRRK2

One hundred nanograms of recombined human Drp1 (GST-tagged protein, Abnova, Taiwan) was incubated with 50 ng of recombined human LRRK2 (GST-tagged protein consisting of amino acid residues 970-2527; either wild-type, G2019S mutant, or D1994A mutant (Invitrogen, USA) in a 25-µl reaction mixture (40 mm Tris–HCl, pH 7.5, 2 mm dithiothreitol, 10 mm MgCl₂, and 100 µm ATP). After incubation at 30° C for 30 min, the reaction was stopped by boiling in the sample-loading buffer for SDS–PAGE. The phosphorylation of Drp1 was detected by anti-threonine phosphorylation antibody.

Generation of iPS cells

PD patient fibroblast Huf-6 and normal adult skin fibroblast HDFa were used to generate iPS cell lines with the protocol as described in (76). Briefly, 1 day before transduction, fibroblasts were seeded at 8 × 10⁴ cells per well of a 6-well plate. Equal volumes of concentrated viral supernatant (0.5 ml) of each of the four transcriptional factors including SOX2, OCT3/4, KLF4 and C-MYC, supplemented with 8 μg/ml Polyprene, were mixed to obtain a final concentration of ×10 and added to growing fibroblasts. The next day, viral supernatants were washed with PBS and replaced with fresh MEF medium. The same infection process was repeated a day later. Four days post-transduction, cells were detached with trypsin, resuspended in MEF medium, counted and seeded at 5 to 10 × 10⁴ cells onto 10-cm dishes preplated with irradiated MEFs. After overnight incubation, MEF medium was replaced with iPS cell medium [DMEM/F12 (1:1) medium (Invitrogen) containing 20% KnockOut™ SR (Invitrogen), 1 mm non-essential amino acids (Invitrogen), 3 mml-glutamine (Invitrogen), 0.1 mm β-mercaptoethanol (Sigma-Aldrich), 100 units/ml penicillin and 100 μg/ml streptomycin (Invitrogen) and 20 ng/ml bFGF (Perprotech, USA). Medium was changed every other day. hESC-like colonies appeared ∼14–20 days post-transduction. Colonies were manually picked and transferred to 12- or 24-well plates preplated with CF1 (Applied Stem Cells., Inc.) feeders for expansion. For enzymatic passaging, iPSCs were incubated with 1 mg/ml collagenase IV/dispase (Invitrogen) at 37°C for 10 min, washed with culture medium three times and scraped off the dish with a cell scraper. Colonies were split at a 1:5 ratio, and transferred to a new dish precoated with irradiated embryonic mouse CF1 feeders or a dish precoated with Matrigel (BD bioscience, for feeder free culturing). For characterization of iPS cells, alkaline phosphatase staining was carried out using the stem cell characterization kit (Millipore, MA, USA). Immunostaining was performed to immunolabel nuclear, cytoplasmic and surface markers of stem cells. These included: Oct ¾ (1:500, Santa Cruz Biotech), SOX2 (1:200, Abgent), SSEA4 (1:50), TRA-1-60 (1:50) and TRA-1-81 (1:50) were from Millipore (MA, USA).

Neuronal differentiation

We followed the differentiation protocol as shown in (48) with some modifications. Briefly, iPS cell colonies were disassociated with Accutase (Invitrogen), plated onto 6-well plates precoated with 2.5% Matrigel (BD Biosciences) and allowed to reach 80% confluence in feeder free medium (Applied stem cells, Inc.). For the first 7 days, cells were treated with SB431542 (10 μm; Tocris Bioscience) and Noggin (100 ng/ml) in the neural media (NM) with FGF2 (20 ng/μl) and EGF (20 ng/μl). NM contained: neurobasal and DMEM (1:1), B-27 supplement minus vitamin A (50×, Invitrogen), N2 Supplement (100×, Invitrogen), GlutaMAX (Invitrogen, 100×), 100 units/ml penicillin and 100 μg/ml streptomycin (Fisher); for the next 4 days, cells were treated with human recombinant Sonic hedgehog (SHH, 200 ng/ml) in neuronal differentiation medium. Neuronal differentiation medium contained neurobalsal and DMEM (1:3), B27, N2, GlutaMax and PS. In the following 3 days, cells were switched to BDNF (20 ng/ml), ascorbic acid (200 μm, Sigma-Aldrich), SHH (200 ng/ml), and FGF8b (100 ng/ml) in neuronal differentiation medium. Thereafter, cells were treated with BDNF, ascorbic acid, GDNF (10 ng/ml), TGF-b (1 ng/ml) and cAMP (500 μm, Sigma-Aldrich). All growth factors were purchased from Perprotech (Rocky Hill, NJ, USA). Neurons were passed onto fresh plates after 30 days of induction and were treated with peptide inhibitor P110 (1 μM/day for 5 days). At 35 days after differentiation, the cells were fixed for immunostaining or mitochondrial function assays.

For spontaneous embryoid body (EB) differentiation, iPS cell colonies were seeded into ultra low attachment hydrogel (Corning) plates containing DMEM:F12 supplemented with 20% FBS (HyClone), 2 mml-glutamine, 0.1 mm non-essential amino acids, and 0.1 mm 2-mercaptoethanol differentiation medium. After 8 days of growth in the suspension, the EBs were transferred to gelatin-coated dishes containing the same medium to allow the cells to expand. Seven days later, the cells were fixed and stained with alpha-SMA (1:500, Santa Cruz Biotech, Dallas, TX, USA), GATA-4 (1:200, Millipore, Billerica, MA, USA) and TUJ1 (1:500, Covance, Princeton, NJ, USA).

For neuronal immunostaining, the cells were fixed and stained with TH (1:200, Millipore or ImmuStar), Tuj1 (1:500, Covance), MAP2 (1:200, Cell Signaling), LAMP2 (1:500, Abcam) and Tom20 (1:500, Santa Cruz Biotech). The imaging was observed by microscope (Fluoview FV100, Olympus).

Western blot analysis

Protein concentrations were determined by the Bradford assay. Thirty micrograms of proteins was resuspended in Laemmli buffer, loaded on SDS–PAGE and transferred onto nitrocellulose membranes. Membranes were probed with the indicated antibody, followed by visualization by ECL.

SUPPLEMENTARY MATERIAL

Supplementary Material is available at HMG online.

FUNDING

The work was partly supported by a grant from the University Hospitals Case medical Center Spitz Brain Health Fund to X.Q.

ACKNOWLEDGEMENTS

We thank Dr Renee Pera's lab at Stanford University for providing PD patient fibroblasts (Huf-6).

Conflict of Interest statement. A patent on the design and application of mitochondrial fission peptide inhibitors has been filed. The authors report no conflicts of interest.