LRRK 2 regulates endoplasmic reticulum–mitochondrial tethering through the PERK ‐mediated ubiquitination pathway

Article10 December 2019Open Access Source DataTransparent process LRRK2 regulates endoplasmic reticulum–mitochondrial tethering through the PERK-mediated ubiquitination pathway Toshihiko Toyofuku Corresponding Author Toshihiko Toyofuku [email protected] orcid.org/0000-0002-7965-2583 Department of Immunology and Molecular Medicine, Graduate School of Medicine, Osaka University, Suita, Japan Search for more papers by this author Yuki Okamoto Yuki Okamoto Department of Immunology and Molecular Medicine, Graduate School of Medicine, Osaka University, Suita, Japan Search for more papers by this author Takako Ishikawa Takako Ishikawa Department of Immunology and Molecular Medicine, Graduate School of Medicine, Osaka University, Suita, Japan Search for more papers by this author Shigemi Sasawatari Shigemi Sasawatari Department of Immunology and Molecular Medicine, Graduate School of Medicine, Osaka University, Suita, Japan Search for more papers by this author Atsushi Kumanogoh Atsushi Kumanogoh Department of Respiratory Medicine and Clinical Immunology, Graduate School of Medicine, Osaka University, Suita, Japan Search for more papers by this author Toshihiko Toyofuku Corresponding Author Toshihiko Toyofuku [email protected] orcid.org/0000-0002-7965-2583 Department of Immunology and Molecular Medicine, Graduate School of Medicine, Osaka University, Suita, Japan Search for more papers by this author Yuki Okamoto Yuki Okamoto Department of Immunology and Molecular Medicine, Graduate School of Medicine, Osaka University, Suita, Japan Search for more papers by this author Takako Ishikawa Takako Ishikawa Department of Immunology and Molecular Medicine, Graduate School of Medicine, Osaka University, Suita, Japan Search for more papers by this author Shigemi Sasawatari Shigemi Sasawatari Department of Immunology and Molecular Medicine, Graduate School of Medicine, Osaka University, Suita, Japan Search for more papers by this author Atsushi Kumanogoh Atsushi Kumanogoh Department of Respiratory Medicine and Clinical Immunology, Graduate School of Medicine, Osaka University, Suita, Japan Search for more papers by this author Author Information Toshihiko Toyofuku *,1, Yuki Okamoto1, Takako Ishikawa1, Shigemi Sasawatari1 and Atsushi Kumanogoh2 1Department of Immunology and Molecular Medicine, Graduate School of Medicine, Osaka University, Suita, Japan 2Department of Respiratory Medicine and Clinical Immunology, Graduate School of Medicine, Osaka University, Suita, Japan *Corresponding author. Tel: +81 0662 108381; E-mail: [email protected] The EMBO Journal (2020)39:e100875https://doi.org/10.15252/embj.2018100875 Correction(s) for this article LRRK2 regulates endoplasmic reticulum–mitochondrial tethering through the PERK-mediated ubiquitination pathway15 September 2020 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract Mutations in the leucine-rich repeat kinase 2 (LRRK2) gene are the most common cause of familial Parkinson's disease (PD). Impaired mitochondrial function is suspected to play a major role in PD. Nonetheless, the underlying mechanism by which impaired LRRK2 activity contributes to PD pathology remains unclear. Here, we identified the role of LRRK2 in endoplasmic reticulum (ER)–mitochondrial tethering, which is essential for mitochondrial bioenergetics. LRRK2 regulated the activities of E3 ubiquitin ligases MARCH5, MULAN, and Parkin via kinase-dependent protein–protein interactions. Kinase-active LRRK2(G2019S) dissociated from these ligases, leading to their PERK-mediated phosphorylation and activation, thereby increasing ubiquitin-mediated degradation of ER–mitochondrial tethering proteins. By contrast, kinase-dead LRRK2(D1994A)-bound ligases blocked PERK-mediated phosphorylation and activation of E3 ligases, thereby increasing the levels of ER–mitochondrial tethering proteins. Thus, the role of LRRK2 in the ER–mitochondrial interaction represents an important control point for cell fate and pathogenesis in PD. Synopsis The contribution of mitochondrial dysfunction to Parkinson's disease remains unclear. Here, the mitochondrial ubiquitination system is identified as a new regulatory target of leucine-rich repeat kinase 2 (LRRK2), revealing a functional link between vulnerability to endoplasmic reticulum (ER) stress and dynamics of the ER-mitochondria interface. LRRK2 mutation decreases mitochondrial biogenesis, respiration and proteasomal degradation in mouse embryonic fibroblasts. LRRK2 mutation alters ER-mitochondrial Ca2+ transfer and sensitivity to ER stress. PERK phosphorylates mitochondrial E3 ubiquitin ligases and decreases the formation of mitochondria-associated ER membranes under stress. Kinase-dead LRRK2 binds E3 ubiquitin ligases and abrogates their PERK-mediated activation. Introduction Parkinson's disease (PD) is a neurodegenerative disorder with no cure. Genetic studies revealed that missense mutations in the protein LRRK2 are the most common cause of familial PD (Funayama et al, 2002; Paisan-Ruiz et al, 2004; Zimprich et al, 2004). In addition, genome-wide association studies have identified a common variation in the LRRK2 gene as a risk factor for sporadic PD (Satake et al, 2009; Simon-Sanchez et al, 2009). LRRK2 encodes a 2,527–amino acid protein consisting of an ankyrin-repeat (ANK) domain, a leucine-rich repeat (LRR), a Ras of complex proteins (ROC) domain, a C-terminal of Roc (COR) domain, and kinase and WD40 domains. Mitochondrial dysfunction has been implicated in a range of neurodegenerative diseases and in PD in particular (Winklhofer & Haass, 2010). The molecular pathogenesis of sporadic PD and the basis of selective dopaminergic neuronal loss remain unclear. Mutations in several genes, including SNCA (encoding alpha-synuclein), DJ-I, LRRK2, PINK1, and PRKN (encoding Parkin), cause forms of familial PD that are clinically indistinguishable from sporadic PD (Klein & Westenberger, 2012). PINK1 and PRKN encode mitochondrially located proteins that participate in mitochondrial quality control, further supporting the idea that mitochondrial dysfunction is sufficient to cause PD. Mitochondria play major roles in multiple cellular processes, including energy metabolism, calcium homeostasis, and lipid metabolism. Mitochondria are associated with the endoplasmic reticulum (ER), with 5–20% of the mitochondrial surface apposed to ER membranes (Rizzuto et al, 1998; Csordas et al, 2006). The regions of the ER associated with mitochondria are termed mitochondria-associated ER membranes (MAMs), and these contacts facilitate a variety of signaling processes between the two organelles, including calcium (Gincel et al, 2001; Rizzuto et al, 2012) and phospholipid exchange (Rowland & Voeltz, 2012), and impact diverse physiological processes including ATP production, autophagy, protein folding, and apoptosis (Simmen et al, 2010; Rowland & Voeltz, 2012; Hamasaki et al, 2013; Kornmann, 2013). Despite the fundamental importance of these interactions to cell metabolism, the mechanisms that mediate recruitment of ER membranes to mitochondria are not fully understood. Several protein complexes have been proposed as ER–mitochondrial tethers, implying that different ER–mitochondrial tethering protein complexes may permit selective recruitment of different domains of the ER, causing the distances between physiological ER–mitochondrial contacts to vary 10–30 nm (Csordas et al, 2006; Rowland & Voeltz, 2012). To maintain energy production and various cellular processes, mitochondrial protein quality control mechanisms are required to counteract the continuous accumulation of defective mitochondrial components. One such mechanism is the dynamic remodeling of mitochondrial membrane through fission and fusion (Karbowski & Youle, 2011), and the other is the ubiquitin/proteasome system, which removes damaged proteins in mitochondria and ER (Christianson & Ye, 2014; Ruggiano et al, 2014). The covalent attachment of ubiquitin to target proteins (substrates) is mediated by the sequential action of an E1-activating enzyme, an E2 conjugase, and an E3 ubiquitin ligase (Pickart & Eddins, 2004). E3 ligases have the ability to bind both E2 proteins (via a RING domain, Ubox, or HECT domain) and substrates. Mitochondria localized E3 ubiquitin ligases such as MARCH5, MULAN, and Parkin ubiquitinate MAM components to regulate MAM formation and mitochondrial morphology (Harder et al, 2004; Braschi et al, 2009; Lokireddy et al, 2012; Nagashima et al, 2014; Gladkova et al, 2018). On the other hand, recent reports have revealed the contribution of ER stress to the pathogenesis of PD (Mercado et al, 2013). ER stress activates the unfolded protein response (UPR), a complex signal-transduction pathway that mediates restoration of ER homeostasis (Doyle et al, 2011). Under chronic ER stress, the UPR triggers cell death by apoptosis, eliminating damaged cells. In mammalian cells, the UPR is initiated by activation of three distinct types of stress sensors located at the ER membrane: two transmembrane kinases, PERK and IRE1α, and transcription factor ATF6. Immunohistochemistry of post-mortem brain tissue from PD patients revealed that the phosphorylated forms of PERK and its substrate, eukaryotic initiation factor 2 α (eIF2α), are present in dopaminergic neurons of the substantia nigra (Hoozemans et al, 2007). However, the mechanisms leading to ER stress in PD and the actual impact of the UPR on this disease remain unclear. In this study, we investigated how LRRK2 is mechanistically involved in mitochondrial biogenesis. By analyzing metabolism and Ca2+ transport in MEFs genetically engineered using the CRISPR/Cas9 system, we identified the mitochondrial ubiquitination system as a key target in LRRK2-mediated mitochondrial biogenesis and showed that LRRK2 regulates ubiquitin ligase activity via PERK under ER stress. Thus, our findings reveal a new functional link between vulnerability to ER stress and mitochondrial biogenesis in the context of PD pathophysiology. Results Experiments were performed using genome-engineered mouse MEFs in which LRRK2 was deleted or replaced with either a kinase-active LRRK2 harboring the most common PD-related mutation (G2019S) or kinase-inactive LRRK2 with the D1994A mutation (Fig EV1A). Kinase assays using a synthetic substrate peptide (LRRKtide) revealed that LRRK2(D1994A) had lower activity, and LRRK2(G2019S) had higher activity, than wild-type LRRK2 (Fig EV1B–D). Click here to expand this figure. Figure EV1. Functional properties of CRISPR/Cas9-engineered MEFs Properties of LRRK2 in MEFs of the indicated genotypes. (Top panel) Immunoblot of endogenous LRRK2 in MEFs. Results confirm that LRRK2 expression is lost in LRRK2−/− MEFs, and show that LRRK2(D1994A) and LRRK2(G2019S) are expressed at levels similar to wild-type LRRK2. (Middle panel) Nucleotide sequences showing that LRRK2 was mutated at D1994 or G2019 in MEFs. Kinase activity of mutant LRRK2 from MEFs, as determined using the ADP-Glo Assay. Data represent kinase activities of LRRK2 prepared from MEFs of the indicated genotypes toward LRRK2-specific substrate (LRRKtide: RLGRDKYKTLRQIRQ). LRRK2(D1994A) had lower kinase activity, whereas LRRK2(G2019S) had higher kinase activity, than wild-type LRRK2. Error bars represent ± SD from eight independent experiments. *P < 0.05 vs. LRRK2. Kinase activity of mutant LRRK2 from transfected LRRK2−/− MEFs, as determined using the ADP-Glo Assay. Data represent kinase activities of LRRK2 from LRRK2−/− MEFs transfected with mutant LRRK2 toward LRRK2-specific substrate (LRRKtide: RLGRDKYKTLRQIRQ). LRRK2(D1994A) had lower kinase activity than LRRK2(G2019S). Error bars represent ± SD from eight independent experiments. Peak values of Ca2+ transients in LRRK2−/− MEFs transfected with mutant LRRK2. Peak values of Ca2+ transients in LRRK2−/− MEFs transfected with LRRK2(D1994A) were higher than those in LRRK2−/− MEFs transfected with control vector, whereas those in LRRK2−/− MEFs transfected with LRRK2(G2019S) were not significantly different from those in LRRK2−/− MEFs transfected with control vector. Error bars represent ± SD from six independent experiments. MEFs of the indicated genotypes were stained with Mitotracker (green) and DAPI (blue). After images were acquired, they were analyzed using a macro developed for the ImageJ software. Mitochondrial morphology was scored as fragmented, intermediate, or tubulated. Data represent the proportions of cells with the indicated mitochondrial morphologies, expressed as percentages of the total number of MEFs counted (≥ 100 cells per experiment). LRRK2−/− and LRRK2(G2019S) MEFs contained more fragmented and less tubulated mitochondria than LRRK2+/+ MEFs. Scale bar: 15 μm. Error bars represent ± SD from six independent experiments. *P < 0.05 vs. LRRK2+/+ MEFs. Extracellular acidification rate (ECAR) (mpH/min) of MEFs of the indicated genotypes was measured on an XF24 Analyzer. ECAR values were similar for all MEFs. Error bars represent ± SD from eight independent experiments. OCR and ECAR of MEFs of the indicated genotypes. OCR and ECAR were lower in LRRK2−/− and LRRK2(G2019S) MEFs, and higher in LRRK(D1994A) MEFs, than in LRRK2+/+ MEFs. Error bars represent ± SD from eight independent experiments. Data information: For graphs (B-G), the P values were determined by a Mann–Whitney U-test. ns = not significant, *P < 0.05. Source data are available online for this figure. Download figure Download PowerPoint Mitochondrial morphology In PD neurons, un-fragmented damaged materials accumulate, possibly due to impaired vesicular trafficking to the lysosome (Abeliovich & Rhinn, 2016). Phosphoproteomics has revealed that LRRK2 phosphorylates a subset of Rab GTPases thereby regulating intracellular endosome trafficking (Steger et al, 2016). Especially, the activity of Rab7 GTPase, a mediator for the late endosome–lysosome transport, is regulated by drosophila LRRK2 homolog (Dodson et al, 2012). Electron micrography revealed multiple large, electron-dense materials in the cytoplasm of both LRRK2−/−, LRRK2(D1994A) and LRRK2(G2019S)-expressing MEFs (Fig 1A), suggesting that LRRK2 mutation impairs the lysosomal degradation of cytosolic debris through defects in trafficking of endosome to lysosome. Visualization of mitochondrial morphology using Mitotracker revealed that the proportion of fragmented mitochondria was elevated in LRRK2−/− and LRRK2(G2019S)-expressing MEFs (Fig EV1E). Consistent with the differences in mitochondrial morphology among MEFs, the activity of citrate synthase, the initial enzyme of the tricarboxylic acid (TCA) cycle and an exclusive marker of the mitochondrial matrix, was reduced in LRRK2−/− and LRRK2(G2019S)-expressing MEFs but elevated in LRRK2(D1994A)-expressing MEFs (Fig 1B). Thus, LRRK2 mutations disturbed mitochondrial biogenesis and/or proteasomal degradation processing. Figure 1. LRRK2 regulates mitochondrial energetics and cellular vulnerability to ER stress A. Electron micrographs of MEFs of the indicated genotypes. Scale bar: 1 μm. High-magnification images depicting representative electron-dense materials are shown. Scale bar: 0.2 μm. B. Citrate synthase activity (absorbance/min) was measured in MEFs of the indicated genotypes. Error bars represent ± SD from eight independent experiments. C, D. Oxygen consumption rates (OCRs) of MEFs of the indicated genotypes were measured on an XF24 Analyzer. (C) Oxygen consumption profiles for MEFs of the indicated genotypes exposed sequentially to oligomycin (2 μg/ml) (Oligo), FCCP (2 μM), and rotenone (1 μM) plus actinomycin (2 μM) (R & A). (D) Maximal OCR (pmol O2/min) (n = 8). Error bars represent ± SD from eight independent experiments. E. ATP production (pmol/min) (n = 8). Error bars represent ± SD from eight independent experiments. F. Representative immunoblot of LC3 and p62 in MEFs of indicated genotype. MEFs were treated with tunicamycin (5 μg/ml) or vehicle control in the presence or absence of bafilomycin A1 (400 nM), and endogenous LC3 and p62 levels were measured by immunoblotting. Data represent the ratios of LC3-II to LC3-I and p62 to actin in the absence of bafilomycin, which were normalized against the corresponding values in LRRK+/+ MEFs. Error bars represent ± SD from four independent experiments. G. Survival rate of MEFs treated with tunicamycin (5 μg/ml), thapsigargin (1 μM), or hydrogen peroxide (100 μM). Error bars represent ± SD from four independent experiments. Data information: For graphs (B and D-G), the P values were determined by a Mann–Whitney U-test. ns = not significant, *P < 0.05, **P < 0.01. Source data are available online for this figure. Source Data for Figure 1 [embj2018100875-sup-0003-SDataFig1.pdf] Download figure Download PowerPoint Mitochondrial oxidative phosphorylation To determine the role of LRRK2 in mitochondrial energetics, we measured basal and maximal (i.e., uncoupled with FCCP) oxygen consumption rate (OCR), an indicator of mitochondrial OXPHOS, and the extracellular acidification rate (ECAR), an indicator of aerobic glycolysis in living MEFs (Figs 1C and D, and EV1F and G). Maximal OCRs were significantly lower in LRRK2(G2019S)-expressing and LRRK2−/− MEFs, but higher in LRKK2(D1994A)-expressing MEFs, indicating that OXPHOS was enhanced by LRRK2(D1994A) but suppressed by loss of LRRK2 or expression of LRRK2(G2019S) (Fig 1C and D). By contrast, ECAR did not differ significantly between MEFs expressing LRRK2 mutants (Fig EV1F), indicating that aerobic glycolysis was not altered by LRRK2 mutation. Relative utilization of OXPHOS and glycolysis, as indicated by the OCR/ECAR ratio, was higher in LRRK2(D1994A)-expressing MEFs and lower in LRRK−/− and LRRK2(D2019S)-expressing MEFs (Fig EV1G). These results suggested that OXPHOS is regulated by LRRK2 in a kinase-dependent manner. The reduced rate of OXPHOS in LRRK2(G2019S)-expressing MEFs could be due to less active mitochondria, a lower density of mitochondria, or a combination of both. Because the rate of OXPHOS predicts ATP production, we next estimated the relative contribution of mitochondrial OXPHOS to ATP production (Fig 1E). To this end, we measured intracellular basal ATP content in MEFs in the absence and presence of oligomycin, a specific inhibitor of the mitochondrial F1F0-ATP synthase, to confirm the involvement of OXPHOS as the source of ATP production. MEFs expressing LRRK2−/− or LRRK2(G2019S) had significantly lower oligomycin-sensitive ATP content than wild-type MEFs. Thus, the reduction in OXPHOS activity due to LRRK2(G2019S) resulted in a decrease in ATP production. Autophagy To determine whether LRRK2 regulates autophagy, we measured the LC3-II level in MEFs (Fig 1F). Under basal conditions, as well as under ER stress induced by tunicamycin, the LC3-II level was higher in LRRK2−/− and LRRK2(S2019S)-expressing MEFs than in LRRK2+/+ MEFs, but lower in LRRK2(D1994A)-expressing MEFs. When LC3-II degradation was blocked with 200 nM bafilomycin A1 (BFA), a specific inhibitor of autophagic degradation, the higher LC3-II levels in LRRK2−/− and LRRK2(S2019S)-expressing MEFs and the lower LC3-II levels in LRRK2(D1994A)-expressing MEFs than in that in LRRK2+/+ MEFs were also detected. Thus, the lower LC3-II levels in LRRK2(D1994A)-expressing MEFs indicated that autophagosome formation was suppressed. By contrast, the higher LC3-II levels in LRRK2−/− and LRRK2(G2019S)-expressing MEFs indicated that autophagosome formation was enhanced. Consistent with the results for LC3-II, the levels of p62, another substrate of autophagy, were reduced in LRRK2(D1994A)-expressing MEFs, but elevated in LRRK2−/− and LRRK2(S2019S)-expressing MEFs. Thus, LRRK2-mutant MEFs exhibited impaired autophagic flux, as previously demonstrated (Alegre-Abarrategui et al, 2009; MacLeod et al, 2013). Cell survival under ER stress To determine whether LRRK2 regulates cellular vulnerability to ER stress, we performed MTT assays to measure the viability response to ER stress inducers such as tunicamycin and thapsigargin, as well as oxidative stressors such as hydrogen peroxide (Fig 1G). Treatment with inducers of ER stress or oxidative stress decreased cell viability more strongly in LRRK2−/− and LRRK2(G2019S)-expressing MEFs than in MEFs expressing other LRRK2 mutants. Thus, LRRK2(G2019S) exhibited greater vulnerability to ER and oxidative stresses. Together, these biochemical analyses indicated that LRRK2 regulates the viability response to ER stress, whereas kinase-active LRRK2(G2019S) enhances cellular vulnerability to this type of stress. Calcium homeostasis Key enzymes of OXPHOS, such as F1F0-ATPase and pyruvate dehydrogenase, are regulated by mitochondrial matrix Ca2+ (Territo et al, 2000; Balaban et al, 2005). Autophagy has been implicated in the IP3R-mediated mechanism (Sarkar et al, 2005; Criollo et al, 2007; Vicencio et al, 2009) and is activated by defects in IP3-induced Ca2+ release (Cardenas et al, 2010). Cellular vulnerability to stress is associated with mitochondrial mishandling of Ca2+ (Orrenius et al, 2003). These findings suggested that changes in mitochondrial biogenesis of MEFs expressing mutant LRRK2 could be due to a defect in a mitochondrial Ca2+-dependent mechanism. Accordingly, we examined the Ca2+ machinery on both the ER and mitochondrial sides. Specifically, we measured Ca2+ transfer from ER to mitochondria by monitoring bradykinin-stimulated calcium release from ER. Mitochondrial calcium transfer To monitor mitochondrial Ca2+ concentration ([Ca2+]m), we targeted the protein-based Ca2+ indicator cameleon to mitochondria and then continuously visualized free Ca2+ in the mitochondrial matrix using fluorescence energy transfer (FRET; Miyawaki et al, 1997, 1999). Data in the figures are presented as absolute Ca2+ concentrations (Fig 2A–D). On average, basal [Ca2+]m in all MEFs was similar levels (2–3 μM). Treatment with 2.5 μM bradykinin significantly increased the [Ca2+]m transient in wild-type MEFs; the level was higher than that in LRRK2−/− and LRRK2(G2019S)-expressing MEFs, but lower than that in LRKK2(D1994A)-expressing MEFs (Fig 2B). Thus, mitochondrial Ca2+ transfer is inactivated by LRRK2−/− and LRRK2(G2019S), but activated by LRRK2(D1994A). By contrast, treatment with bradykinin significantly decreased ER Ca2+ concentration ([Ca2+]ER), as measured by the protein-based Ca2+ indicator ER-D1 targeted to the ER, in wild-type MEFs; the level was lower than that in LRRK2−/− and LRRK2(G2019S)-expressing MEFs (Fig EV2A and B). Thus, the magnitude of the change in [Ca2+]m was reciprocal with that of the change in [Ca2+]ER. Figure 2. LRRK2 regulates ER–mitochondrial Ca2+ transfer and tethering A–D. MEFs were transfected with mitochondrially targeted cameleon. Free Ca2+ dynamics in the mitochondrial matrix were visualized using FRET. Mitochondrial [Ca2+] ([Ca2+]m) was continuously monitored by FRET imaging; data are represented as absolute [Ca2+] in μmol. (A) Absolute [Ca2+]m changes in MEFs of indicated genotype in response to bradykinin (2.5 μM). (B) Basal and peak values of Ca2+ transients (μM). (C) Peak values of Ca2+ transients in MEFs transfected with IP3R or shRNA against IP3R, or treated with 2-AP (20 mM). (D) Peak values of Ca2+ transients in MEFs transfected with VDAC1 or shRNA against VDAC1. Error bars represent ± SD from six independent experiments. E. Representative electron micrographs of MEFs of the indicated genotypes. Full arrowheads designate the limits of the zone of intimate contact between ER and mitochondria (< 20 nm). Scale bar: 100 nm. Data represent percentage of ER–mitochondrial contacts per unit of mitochondrial perimeter. Error bars represent ± SD from 40 images from MEFs of the indicated genotypes. F. In situ PLA images using anti-IP3R and anti-VDAC1 antibodies. Scale bar: 20 μm. Data represent the number of fluorescent puncta in MEFs of the indicated genotypes, normalized against the value for LRRK2+/+ MEFs. Error bars represent ± SD from six independent experiments. G. Peak values of Ca2+ transients in MEFs transfected with synthetic tethering protein (TOM-mRFP-ER) to induce artificial tethering of the ER and mitochondria. Error bars represent ± SD from six independent experiments. Data information: For graphs (B-G), the P values were determined by a Mann–Whitney U-test. ns = not significant, *P < 0.05, **P < 0.01. Download figure Download PowerPoint Click here to expand this figure. Figure EV2. ER–mitochondrial interaction in MEFs expressing LRRK2 mutants MEFs were transfected with ER-targeted D1ER pcDNA3. Free Ca2+ dynamics in the ER matrix ([Ca2+] ER) were visualized by FRET. Data are represented as absolute [Ca2+] in μmol. Error bars represent ± SD from eight independent experiments. Peak values of [Ca2+] ER in MEFs of the indicated genotype. Peak [Ca2+] ER levels were lower in LRRK2−/− and LRRK2(G2019S) MEFs, and higher in LRRK2(D1994A) MEFs, than in LRRK2+/+ MEFs. Error bars represent ± SD from eight independent experiments. ER–mitochondrial contacts, as determined by electron microscopy of MEFs of the indicated genotypes. Bar graph shows ER–mitochondrial distance (nm), which was similar in all MEFs. Error bars represent ± SD from 40 images. Colocalization of IP3R and VDAC1 in MEFs. (Top panel) Immunofluorescence images of IP3R and VDAC1 in MEFs. MEFs were immunostained with mouse anti-IP3R and rabbit anti-VDAC1 antibodies, followed by anti-mouse IgG-FITC and anti-rabbit IgG-DsRed. Merged image shows the close apposition between ER-localized IP3R and mitochondria-localized VDAC1. Scale bar: 20 μm.(Bottom panel) In situ PLA images of MEFs transfected with IP3R shRNA or VDAC1 shRNA. Loss of either endogenous IP3R or VDAC1 expression dramatically decreased PLA intensity. Schematic of synthetic tethering protein (TOM-mRFP-ER) composed of the mitochondrial targeting domain of mouse TOM70, mRFP, and the ER-targeting domain of yeast UBC6. Data represent the results of quantitative analysis of the IP3R–VDAC interaction in MEFs, as determined from in situ PLA images using anti-IP3R and anti-VDAC antibodies. PLA intensities in LRRK2−/− and LRRK2(G2019S)-expressing MEFs increased to the intensity in LRRK2+/+ MEFs transfected with empty vector (Control). Error bars represent ± SD from six independent experiments. Data information: For graphs (B, C and E), the P values were determined by a Mann–Whitney U-test. ns = not significant, *P < 0.05. Download figure Download PowerPoint Close proximity between ER-localized IP3R and OMM-localized VDAC1 at the MAM potentiates rapid transfer of Ca2+ through the OMM. Mitochondrial Ca2+ accumulation is augmented by IP3-activated IP3R (Rizzuto et al, 1993) or over-expression of VDAC1 (Madesh & Hajnoczky, 2001; Rapizzi et al, 2002), but attenuated by down-regulation of either protein. To determine whether IP3R or VDAC1 is involved in disrupting mitochondrial Ca2+ accumulation in LRRK-modified MEFs, we measured [Ca2+]m in MEFs in which IP3R or VDAC1 was modified. shRNA-mediated down-regulation of IP3R or pretreatment with 20 μM 2-APB, a membrane-permeable blocker of IP3R, attenuated [Ca2+]m in all MEFs (Fig 2C), confirming the crucial role of IP3R in mitochondrial Ca2+ transfer. Over-expression of IP3R increased peak [Ca2+]m in LRRK2(D1994A)-expressing MEFs but not in LRRK2−/− or LRRK2(G2019S)-expressing MEFs. Down-regulation of VDAC1 decreased peak [Ca2+]m in all MEFs, whereas over-expression of VDAC1 increased peak [Ca2+]m in LRRK2(D1994A)-expressing MEFs but not in LRRK2−/− or LRRK2(G2019S)-expressing MEFs (Fig 2D). Thus, Ca2+ transfer through IP3R and VDAC1 was suppressed by LRRK2(G2019S), but enhanced by LRRK2(D1994A). Physical interaction and Ca2+ transfer between ER and mitochondria To obtain insight into the mechanism by which LRRK2 influences ER–mitochondrial Ca2+ transfer, we analyzed the relationship between the ER and mitochondria. Specifically, we performed ultrastructural analysis by electron microscopy to evaluate ER–mitochondrial contacts (Figs 2E and EV2C). Visual inspection of EM images acquired by facility personnel blinded to sample identity revealed a reduction in the number of ER–mitochondrial contact sites per unit of mitochondrial perimeter in LRRK2−/− and LRRK2(G2019S)-expressing MEFs. Thus, LRRK2 ablation and LRRK2(G2019S) block ER–mitochondrial contacts. Next, we examined the physical interaction between the ER and mitochondria by in situ proximity ligation assay (PLA) using two organelle-surface proteins involved in the calcium channeling complex: IP3R and VDAC1 at the MAM interface (Fig 2F; De Vos et al, 2012; Hedskog et al, 2013). IP3R and VDAC1 were in close proximity in wild-type MEFs, in which PLA intensity was higher than that in LRRK2−/− and LRRK2(G2019S)-expressing MEFs, but lower than that in LRKK2(D1994A)-expressing MEFs. Thus, LRRK2 might be involved in MAM formation in a kinase-dependent manner. To determine whether reduced ER–mitochondrial Ca2+ transfer in LRRK2−/− and LRRK2(G2019S)-expressing MEFs was indeed due to a decrease in ER–mitochondrial contacts, we performed a rescue experiment using a synthetic ER–mitochondrial tethering protein, TOM-mRFP-ER)(Csordas et al, 2006; Kornmann, 2013), which restores changes in PLA intensity in LRRK2−/− and LRRK2(G2019S)-expressing MEFs (Figs 2G and EV2E). Over-expression of TOM-mRFP-ER significantly rescued mi