ER-mitochondria cross-talk is regulated by the Ca2+ sensor NCS1 and is impaired in Wolfram syndrome
The ER-mitochondria connection for Ca2+
Loss-of-function mutations in the ER protein WFS1 result in Wolfram syndrome; however, some of the defining symptoms of this disorder, such as diabetes and optic atrophy, are due to mitochondrial dysfunction. Using fibroblasts from Wolfram syndrome patients or normal individuals, Angebault et al. found that WFS1 deficiency was associated with decreased Ca2+ uptake by mitochondria, reduced mitochondrial contact with the ER, and decreased mitochondrial respiration. WFS1 interacted with a Ca2+-sensing protein, called NCS1, and NCS1 abundance was lower in patient fibroblasts than in control fibroblasts. Reconstituting NCS1 in patient fibroblasts restored mitochondrial respiration and Ca2+ signaling dynamics. These results explain how a deficiency in an ER protein impairs mitochondrial activity and suggest that defective ER-mitochondria association may contribute to the pathogenesis of neurodegenerative disorders.
Abstract
Communication between the endoplasmic reticulum (ER) and mitochondria plays a pivotal role in Ca2+ signaling, energy metabolism, and cell survival. Dysfunction in this cross-talk leads to metabolic and neurodegenerative diseases. Wolfram syndrome is a fatal neurodegenerative disease caused by mutations in the ER-resident protein WFS1. Here, we showed that WFS1 formed a complex with neuronal calcium sensor 1 (NCS1) and inositol 1,4,5-trisphosphate receptor (IP3R) to promote Ca2+ transfer between the ER and mitochondria. In addition, we found that NCS1 abundance was reduced in WFS1-null patient fibroblasts, which showed reduced ER-mitochondria interactions and Ca2+ exchange. Moreover, in WFS1-deficient cells, NCS1 overexpression not only restored ER-mitochondria interactions and Ca2+ transfer but also rescued mitochondrial dysfunction. Our results describe a key role of NCS1 in ER-mitochondria cross-talk, uncover a pathogenic mechanism for Wolfram syndrome, and potentially reveal insights into the pathogenesis of other neurodegenerative diseases.
INTRODUCTION
Mitochondria play a role in various essential cellular functions, including bioenergetics, metabolism, ion homeostasis, and apoptosis. Mitochondrial dysfunction has been linked to many common neurodegenerative disorders that are disabling and are often fatal. However, the cause of mitochondrial dysfunction remains largely undefined. Mitochondria interact physically and functionally with endoplasmic reticulum (ER) through mitochondria-associated membranes (MAMs) (1–3) to influence cellular physiology and viability (4, 5). Accordingly, impaired MAM function is increasingly recognized as a contributor to neurodegeneration (6, 7).
Loss of function of the ER protein WFS1 leads to a neurodegenerative disease called Wolfram syndrome that is associated with diabetes, optic atrophy, and deafness (8). Wolfram syndrome was originally described as a mitochondriopathy due to a clinical phenotype that resembled other mitochondrial disorders (9). However, this hypothesis has been challenged because WFS1 localized to the ER (10). A relationship between cytosolic Ca2+ disturbance, impaired mitochondrial dynamics, and delayed neuronal development in WFS1-deficient neurons has been suggested (11). Here, we report the identification of a WFS1 binding partner, neuronal calcium sensor 1 (NCS1), a cytoplasmic Ca2+ binding protein that is abundant in neurons. Our data suggest that NCS1 promoted ER-mitochondria contact and Ca2+ transfer, subsequently controlling mitochondrial bioenergetics. This study shows that WFS1 and NCS1 may participate in the tethering of ER to mitochondria and the lack of this interaction may contribute to neurodegeneration.
RESULTS
Loss of function of WFS1 does not impair mitochondrial morphology
To investigate the effect of WFS1 deficiency, we took advantage of fibroblasts from Wolfram syndrome patients, which are WFS1 null. The patients carried predicted loss-of-function alleles in WFS1 (patients 1 and 2, V509-Y513del and F882fsX950; patient 3, homozygous for L432V; patient 4, c.1-43G>T and W867X; Fig. 1A) and exhibited symptoms typical for Wolfram syndrome (table S1). Although WFS1 protein abundance was decreased in all patients (Fig. 1B), WFS1 mRNA expression was only decreased in patients 3 and 4 (Fig. 1C). These results suggested that different mutations resulted in distinct effects on mRNA expression and that WFS1 loss-of-function mutations affected the production or stability of the protein. This conclusion is consistent with previous results, showing a decrease in the stability of mutant WFS1 protein overexpressed in COS-7 cells (12, 13). Measurement of the mitochondrial membrane potential (Δψm) using tetramethylrhodamine methyl ester (TMRM) dye after application of the mitochondrial uncoupler carbonyl cyanide p-trifluoromethoxyphenylhydrazone (FCCP) revealed similar Δψm in control and patient cells (Fig. 1D). We validated this result using the ratiometric JC-1 (5,5,6,6-tetrachloro-1,1,3,3-tetraethylbenzimidazolylcarbocyanine iodide) probe (Fig. 1, E and F), which confirmed that loss of function of WFS1 did not change Δψm. Ultrastructural examination of mitochondria with transmission electron microscopy (TEM) indicated that the overall structure (presence of a double membrane and presence of mitochondrial cristae located perpendicular to their major axis) and their form (elongated) were similar in patient and control cells (Fig. 1G). The exception was patient 4, whose cells had expanded ER (Fig. 1G). This ER morphology (dilated and irregular lumen) has been previously noted in Wfs1- or Cisd2-deficient cells (14, 15).
Fig. 1 WFS1-deficient fibroblasts have normal mitochondria.
(A) Schematic representation of WFS1 showing the positions of the disease-causing mutations. (B) Immunoblot of lysates from control (C1) and patient fibroblasts (P1 to P4) for WFS1. Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) was used as a loading control (n = 5 independent experiments). (C) Transcript levels of WFS1 were quantified by reverse transcription quantitative polymerase chain reaction (RT-qPCR) in control (C1 and C2) and patient fibroblasts (P1 to P4). The mRNA levels were normalized to the reference gene L27. Error bars represent SEM. Controls, n = 9; P1, n = 6; P2, n = 6; P3, n = 6; P4, n = 5 independent experiments. (D) Assessment of Δψm with TMRM in control (C) (n = 36) and patient (P) (n = 34; P1, n = 10; P2, n = 7; P3, n = 9; P4, n = 8) fibroblasts. TMRM fluorescence was recorded continuously for 5 min before and after FCCP (1 μM) application. To enable comparison between groups, we calculated the percentage of TMRM fluorescence decreased by FCCP for each group. All data represent means ± SEM. NS, not significant. (E) Δψm in control (C) and patient (P) fibroblasts detected by the dye JC-1. Red fluorescence indicates the mitochondrial, aggregated form of JC-1. Green fluorescence indicates the monomeric form of JC-1. Scale bars, 10 μm. (F) Ratio of red to green fluorescence intensity, indicating the ratio of the aggregate to monomeric form of JC-1, in control (n = 24) and patient (n = 28) fibroblasts. Data are means ± SEM. (G) Ultrastructure of the mitochondria (top) and the ER (bottom) by TEM. Scale bars, 100 nm (top) and 500 nm (bottom).
WFS1 deficiency is associated with decreased mitochondrial Ca2+ uptake
WFS1 has been suggested to play a role in Ca2+ homeostasis by activating an inositol 1,4,5-trisphosphate receptor (IP3R)–mediated Ca2+ release pathway (16), inhibiting sarco/endoplasmic reticulum Ca2+-ATPase (SERCA) turnover and modifying the filling state of the ER Ca2+ store (17, 18). WFS1 deficiency has also been suggested to lead to IP3R dysfunction, disturbed cytosolic Ca2+ homeostasis, and altered mitochondrial dynamics (11). Accordingly, we hypothesized that the loss of function of the ER protein WFS1 affected Ca2+ flux between the ER-mitochondria through regulation of IP3R. Coimmunoprecipitation experiments demonstrated that mouse WFS1 interacted with IP3R (fig. S1). We then measured the effect of WFS1 loss of function on Ca2+ flux using the ER-targeted aequorin (erAEQ; Fig. 2A) (19). There were no significant differences in steady-state ER Ca2+ ([Ca2+]ER) between control and WFS1 patient fibroblasts (Fig. 2B). Next, we investigated the ER response to the Ca2+-mobilizing agonist bradykinin, which activates Gq-coupled plasma membrane receptors to trigger the production of IP3 and release of Ca2+ from the ER through the IP3R. Before bradykinin stimulation, fibroblasts were transduced with the Ca2+-sensitive FRET (fluorescence resonance energy transfer)–based cameleon protein D1-ER (Fig. 2C) (20). We found that mutated fibroblasts released 32% less ER Ca2+ than did control fibroblasts (Fig. 2D), suggesting that Wolfram syndrome patient fibroblasts release less Ca2+ through the IP3R. To clarify whether loss of function of WFS1-suppressed IP3-mediated cytosolic [Ca2+] signals was upstream or at the level of the IP3Rs, we performed aequorin experiments in permeabilized cells stimulated with IP3. In response to IP3, patient fibroblasts released less Ca2+ from the ER than did control fibroblasts (Fig. 2, E and F). These results suggest that WFS1 loss impairs Ca2+ signaling directly at the level of IP3R.
Fig. 2 Loss of function in WFS1 impairs Ca2+ homeostasis.
(A) ER Ca2+ concentration ([Ca2+]ER) in control (black line; n = 19 fibroblasts) and P3 patient (red line; n = 19 fibroblasts) fibroblasts transiently expressing erAEQmut. (B) Histogram plotting [Ca2+]ER steady-state level in control (black) and P3 patient (red) fibroblasts. (C) ER Ca2+ release induced by bradykinin (100 μM) in control (black; n = 17) and P3 patient (red; n = 16) fibroblasts transduced with the Ca2+-sensitive FRET-based cameleon protein D1-ER. (D) Histogram plotting ER Ca2+ release in control (black) and P3 patient (red) cells. Data are means ± SEM. *P < 0.05. (E) ER Ca2+ concentration ([Ca2+]ER) in control (black line) and P3 patient (red line) fibroblasts transiently expressing erAEQ. After depletion of intracellular Ca2+ stores and aequorin reconstitution, cells were perfused in immunoblotting buffer. CaCl2 was added to give a calculated free [Ca2+] of 500 nM. After steady state was achieved, IP3 (5 M) was introduced to evoke ER-Ca2+ release. (F) Histogram plotting mean rates of Ca2+ release for the first 50 s after stimulation in control (n = 12) and patient (n = 13) fibroblasts. Data are presented as means ± SEM. *P < 0.05. (G) Representative cytosolic Ca2+ measured by Fluo-4 fluorescence in control (black line) and patient (red line) fibroblasts under histamine stimulation. (H) Maximum amplitude of Fluo-4 fluorescence in control (n = 26) and patient (n = 35) fibroblasts. (I) Rate of cytosolic Ca2+ rise in control (n = 25) and patient (n = 35) fibroblasts. (J) Rate of cytosolic Ca2+ decay in control (n = 25) and patient (n = 35) fibroblasts. (K) Representative mitochondrial Ca2+ ([Ca2+]m) as measured by pericam-mt fluorescence in control (black line) and patient (red line) fibroblasts stimulated with histamine. (L) Maximum amplitude of pericam-mt fluorescence in control (n = 23) and patient (n = 36) fibroblasts. (M) Rate of [Ca2+]m entry in control (n = 23) and patient (n = 36) fibroblasts. (N) Rate of [Ca2+]m decay in control (n = 23) and patient (n = 36) fibroblasts. (O) [Ca2+]m uptake rate with 4 μM Ca2+ in control (black line; n = 8 fibroblasts) and P3 patient (red line; n = 8 fibroblasts) permeabilized cells expressing mitochondrial aequorin (mt-AEQ). (P) Histogram plotting [Ca2+]m uptake rate in control (black) and P3 patient (red) fibroblasts. (Q) [Ca2+]m uptake rate with 1 μM Ca2+ in control (black line; n = 9 fibroblasts) and P3 patient (red line; n = 15 fibroblasts) permeabilized cells expressing mt-AEQ. (R) Histogram plotting the [Ca2+]m uptake rate in control (black) and P3 patient (red) cells. Data are means ± SEM. *P < 0.05, **P < 0.01, and ***P < 0.001.
Depletion of ER Ca2+ with thapsigargin, a specific inhibitor of the SERCA pump, did not reveal differences in ER Ca2+ content between controls and patients (fig. S2, A to D). Histamine induces IP3R-dependent ER Ca2+ release from ER stores (21). As demonstrated by confocal imaging of cytoplasmic and mitochondrial [Ca2+] with Fluo-4 and the genetically encoded reporter pericam-mt, respectively, histamine-induced ER Ca2+ release in patient cells was decreased relative to that in control cells (Fig. 2G). In addition to the maximum peak amplitude, the rate of cytosolic Ca2+ rise (but not the rate of cytosolic Ca2+ decay) was also decreased in patient cells compared to control cells (Fig. 2, H to J). Ca2+ uptake by mitochondria was reduced in patient fibroblasts (Fig. 2, K and L). The rates of [Ca2+]m entry and of [Ca2+]m decay were similar in patient and control cells (Fig. 2, M and N). Because Δψm was not affected, the lack of [Ca2+]m uptake in patient cells was not due to reduced driving force. To confirm these results, we performed experiments in digitonin-permeabilized cells perfused with medium at fixed [Ca2+]. When Ca2+ was added back to the perfusion medium, the rates and peaks of [Ca2+]m uptake were similar between control and patient cells (Fig. 2, O to R). Overall, these data suggest that the decrease in [Ca2+]m uptake of patient fibroblasts was due to decreased ER calcium release through IP3R.
WFS1 deficiency is associated with decreased ER-mitochondria interactions
Cross-talk between ER and mitochondria is essential for processes in eukaryotic cells such as lipid synthesis and transport (22), mitochondrial functions (23), regulation of Ca2+ homeostasis (24), autophagy (3), and apoptosis (25). The ER directly communicates with mitochondria through close contacts referred to as MAMs, which are microdomains that enable efficient Ca2+ transfer between the ER and mitochondria. Ca2+ transfer from the ER into the mitochondria is facilitated by proteins that tether the two organelles together. IP3R on the ER interacts with VDAC1 on the outer mitochondrial membrane through the molecular chaperone glucose-regulated protein 75 (GRP75) (26). The IP3R/GRP75/VDAC1 complex also involves the [Ca2+]m uniporter (MCU) on the inner mitochondrial membrane (26) to allow Ca2+ transfer from the ER to the mitochondrial matrix. To study the role of MAMs in Wolfram syndrome, we used in situ proximity ligation assay (PLA) to detect and quantify both VDAC1-IP3R1 and GRP75-IP3R1 interactions at the MAM interface in fixed fibroblasts, as previously described (27). We found significantly reduced VDAC1-IP3R1 and GRP75-IP3R1 interactions in patient fibroblasts compared to control fibroblasts (−70% and −56%, respectively), indicating reduced ER-mitochondria interactions in patient fibroblasts (Fig. 3A). To confirm this observation, we evaluated the percentage of mitochondria in close contact with ER in control and patient cells with TEM morphometric analysis (Fig. 3, B to D, and table S2). Mitochondria-ER associations were found in control and patient cells (Fig. 3B). However, the percentage of mitochondria with ER contact was significantly reduced by 49% in patient cells (Fig. 3C). To evaluate more precisely the importance of these contacts, we measured the percentage of mitochondria perimeter in contact with the ER, and this value was significantly reduced by 38% in patient cells (Fig. 3D). However, there were no significant differences in the protein abundance of IP3R (Fig. 3E), GRP75 (Fig. 3F), VDAC1, MCU (Fig. 3G), or MFN2 (Fig. 3H), another protein that regulates the tethering of the ER to mitochondria (28).
Fig. 3 Loss of function in WFS1 disruption MAM structure.
(A) Representative PLA images (left) and quantitative analysis (right) of VDAC1-IP3R1 and GRP75-IP3R1 interactions in control and patient fibroblasts (8 to 21 images/controls and patients, n = 56 images for VDAC1-IP3R1 and n = 22 to 43 images for GRP75-IP3R1). a.u., arbitrary units. ***P < 0.001 compared to controls. Scale bar, 10 μm. (B) TEM of control and patient fibroblasts. The red arrowheads indicate the perimeter of mitochondria (M) in contact with ER. Scale bars, 100 nm. (C) Quantitative analysis of ER-mitochondria contacts in control and patient fibroblasts. Bar graph shows the percentage of mitochondria in contact with ER (means ± SEM, n = 403 mitochondria in control cells and n = 659 mitochondria in patient cells). *P < 0.05 compared to controls. (D) Quantification of mitochondrial length adjacent to ER normalized by total mitochondrial perimeter in control and patient fibroblasts. Bar graph shows the percentage of mitochondrial perimeter in contact with ER (means ± SEM, n = 403 mitochondria in control cells and n = 659 mitochondria in patient cells). ***P < 0.001 compared to controls. (E) Western blot (top) and densitometric analysis (bottom) of IP3R protein abundance in control (C1 to C3, n = 4 experiments) and patient (P1, n = 3; P2, n = 3; P3, n = 4; P4, n = 4 experiments) fibroblasts. (F) Western blot (top) and densitometric analysis (bottom) of GRP75 protein abundance in control (C1 to C3, n = 5 experiments) and patient (P1, n = 5; P2, n = 5; P3, n = 5; P4, n = 5 experiments) fibroblasts. (G) Western blot (top) and densitometric analysis (bottom) of VDAC1 and MCU protein abundance in control (C1 to C3, n = 3 to 6 experiments) and patient (P1, n = 4; P2, n = 4; P3, n = 6; P4, n = 6 experiments) fibroblasts. (H) Western blot (top) and densitometric analysis (bottom) of MFN2 protein abundance in control (C1 to C3, n = 4 to 9 experiments) and patient (P1, n = 5; P2, n = 6; P3, n = 4; P4, n = 4 experiments) fibroblasts. GAPDH was used as a loading control (means ± SEM). n in (E) to (H) represents experiments.
WFS1 deficiency impairs mitochondrial functionality
Reduced [Ca2+]m uptake might adversely affect various metabolic pathways, resulting in altered energy production. We evaluated mitochondrial functionality by analyzing oxygen consumption and the enzymatic activity of complexes and citrate synthase (CS). WFS1 patient cells exhibited a significant decrease (30%) in complex II–driven respiration (Fig. 4A), and complex I–driven respiration was decreased in cells from three of four patients (fig. S3). To understand the cause of the decreased mitochondrial respiration in patient fibroblasts, we assessed the activities of respiratory chain complexes I [NADH (reduced form of nicotinamide adenine dinucleotide) ubiquinone reductase], II (succinate ubiquinone reductase), and IV (cytochrome c oxidase) in cell homogenates. The activity of complex I was significantly decreased in patient cells (40%), whereas the activity of complexes II and IV were similar between patient and control cells (Fig. 4B). No changes were observed in CS activity (Fig. 4B), suggesting that mitochondrial mass was normal. Therefore, we measured the protein abundance of selected subunits of each complex in the mitochondrial electron transport chain. The protein abundance of complex I (NDUFA13; Fig. 4C), complex IV (MTCO1; Fig. 4D), and complex V [adenosine 5′-triphosphate (ATP) synthase; Fig. 4E] subunits did not differ between control and patient cells. Patient cells showed a decrease in the protein abundance of complex II (SDHA; Fig. 4F) and complex III (UQCRC2) subunits (Fig. 4G). Protein abundance was quantified for each complex (Fig. 4H).
Fig. 4 Loss of function in WFS1 impairs mitochondrial functionality.
(A) Mitochondrial complex I– and complex II–dependent respiratory rate. Respiratory rate: nanomole of oxygen consumed per minute per milligram of protein (means ± SEM; controls, n = 5 experiments; P1, n = 3; P2, n = 4; P3, n = 6; P4, n = 6 experiments). *P < 0.05. (B) Complexes (Cx) I, II, and IV and CS enzymatic activities in control and patient fibroblasts (means ± SEM; controls, n = 3 to 6 experiments; P1, n = 3; P2, n = 3; P3, n = 5; P4, n = 5 experiments). *P < 0.05. (C) Western blot of NDUFA13 (complex I), (D) MTCO1 (complex IV), (E) ATP synthase (complex V), (F) SDHA (complex II), and (G) UQCRC2 (complex III) analysis in control (C1 to C3) and patient (P1 to P4) fibroblasts. GAPDH was used as a loading control. (H) Densitometric analysis in control and patient fibroblasts (means ± SEM; controls, n = 3 to 5 experiments; P1, n = 5; P2, n = 5; P3, n = 7; P4, n = 7 experiments). *P < 0.05 compared to control.
WFS1 regulates mitochondrial functionality through NCS1
We sought to identify protein partners of the cytosolic domain of WFS1 by performing a yeast two-hybrid analysis using residues 1 to 311 of murine Wfs1 as bait to screen a random-primed complementary DNA (cDNA) library prepared from mouse inner ear. A total of 29 positive colonies were obtained (table S3). We identified mouse Ncs1 as an interacting partner (Fig. 5A). NCS1 is an EF-hand cytosolic protein preferentially expressed in neurons (29) that regulates IP3R (30–32) and the dopamine D2 receptor (33). We confirmed the interaction of NCS1 with Myc-tagged WFS1 and with endogenous IP3R by coimmunoprecipitation (fig. S4, A and B). Although fibroblast NCS1 mRNA expression levels were similar between control and patient fibroblasts (Fig. 5B), NCS1 protein levels were decreased by almost 50% in patient fibroblasts (Fig. 5C). Consistent with this observation, knockdown of WFS1 using short interfering RNA (siRNA) in control fibroblasts decreased NCS1 protein levels by 60% (Fig. 5D and fig. S5). Together, these results suggest that WFS1 regulates NCS1 protein stability, which is consistent with the ability of WFS1 to regulate the stability of other proteins, such as HRD1 (34) and SERCA (18), through the ubiquitin proteasome pathway. Last, because IP3R and WFS1 are localized in MAMs (35–37), we looked at the cellular distribution of NCS1 in mouse brain subjected to subcellular fractionation. NCS1 was not only localized mainly in the ER and in MAMs (Fig. 5E) but also faintly detected in the mitochondrial fraction (Fig. 5E), which might be due to its interaction with PINK1 (38).
Fig. 5 WFS1 interacts with NCS1 and modulates its abundance.
(A) Schematic representation of the interaction site between mouse Wfs1 and Ncs1 identified by yeast two-hybrid screening. A fragment encompassing amino acids 1 to 311 of human WFS1 was used for the yeast two-hybrid screening. EF-hand motifs in Ncs1 are helix-loop-helix structural domains found in a large family of Ca2+ binding proteins. (B) Transcript levels of NCS1 were quantified by RT-qPCR in control (C1 and C2) and patient fibroblasts (P1 to P4). The mRNA levels were normalized to the reference gene L27 (means ± SEM; controls, n = 9; P1, n = 6; P2, n = 6; P3, n = 6; P4, n = 5 independent experiments). (C) Immunoblot of lysates from control (C1 to C3) and patient fibroblasts (P1 to P4) for NCS1. GAPDH was used as a loading control. The immunoblot bands were quantified by densitometry, and the NCS1/GAPDH ratios were calculated (controls, n = 4; P1, n = 4; P2, n = 4; P3, n = 5; P4, n = 5 independent experiments). (D) Western blot analysis and quantification of control fibroblasts transfected with scrambled siRNA (siScr) or WFS1 siRNA (siWFS1). WFS1 and NCS1 protein abundance were analyzed 72 hours after siRNA transfection. Data are means ± SEM from n = 7 independent experiments. *P < 0.05. (E) Subcellular fractions prepared from mouse brain tissue were immunoblotted for HSP60 (a mitochondrial marker), SIL1 (an ER marker), VDAC1 (a MAM marker), and NCS1. H, homogenate; ER, endoplasmic reticulum fraction; Mp, pure mitochondrial fraction; MAMs, mitochondria-associated membrane fraction.
Knockdown of NCS1 impairs Ca2+ homeostasis and mitochondrial function
To elucidate the role of NCS1 in mitochondrial function, we used siRNAs to deplete NCS1 in human fibroblasts (fig. S6). Similar to Wolfram syndrome patient cells, histamine-mediated Ca2+ release was reduced in cells with NCS1 knockdown (Fig. 6A), specifically the maximum peak amplitude and the rate of cytosolic Ca2+ rise (Fig. 6, B and C) but not the rate of cytosolic Ca2+ decay (Fig. 6D). In addition, [Ca2+]m uptake was significantly reduced in NCS1-deficient cells (Fig. 6E). Although the maximum peak amplitude was reduced in cells with NCS1 knockdown (Fig. 6F), [Ca2+]m entry and decay were similar between siNCS1- and siScr-treated cells (Fig. 6, G and H). Knockdown of NCS1 led to a decrease in both complex I– and complex II–driven respiration, similar to that observed in patient fibroblasts (Fig. 6I). Moreover, NCS1 knockdown also decreased VDAC1 and SDHA (complex II) protein abundance compared to controls (Fig. 6J) and the proportion of mitochondria contacting ER (Fig. 6K). These results demonstrated that NCS1 modulated [Ca2+]m uptake and could be a key determinant to maintain mitochondrial function and MAM integrity.
Fig. 6 NCS1 knockdown impairs Ca2+ homeostasis and mitochondrial function.
(A) Representative cytosolic Ca2+ measured by Fluo-4 fluorescence in control fibroblasts treated with nontargeting siScr (black line) or siNCS1 (red line) and stimulated with histamine. (B) Maximum amplitude of Fluo-4 fluorescence in control fibroblasts treated with nontargeting siScr (black line; n = 13 cells) or siNCS1 (red line; n = 12 cells). (C) Rate of cytosolic Ca2+ rise in control fibroblasts treated with nontargeting siScr (black line; n = 13 cells) or siNCS1 (red line; n = 12 cells). (D) Rate of cytosolic Ca2+ decay in control fibroblasts treated with nontargeting siScr (black line; n = 13 cells) or siNCS1 (red line; n = 12 cells). (E) Representative [Ca2+]m measured by pericam-mt fluorescence in control fibroblasts treated with nontargeting siScr (black line) or siNCS1 (red line). (F) Maximum amplitude of pericam-mt fluorescence in control fibroblasts treated with nontargeting siScr (black dots; n = 25 cells) or siNCS1 (red dots; n = 24 cells). (G) Rate of [Ca2+]m entry in control fibroblasts treated with nontargeting siScr (black line; n = 25 cells) or siNCS1 (red line; n = 24 cells). (H) Rate of [Ca2+]m decay in control fibroblasts treated with nontargeting siScr (black line; n = 25 cells) or siNCS1 (red line; n = 24 cells). (I) Complex I– and complex II–mediated respiration of control fibroblasts transfected with nontargeting siRNA (siScr) or NCS1 siRNA (siNCS1) (means ± SEM, n = 3 experiments). (J) Effect of NCS1 and WFS1 knockdown on NCS1, VDAC1, and SDHA (complex II) protein abundance in control fibroblasts transfected with nontargeting (siScr), NCS1 (siNCS1), or WFS1 (siWFS1) siRNA. NCS1 knockdown effect is shown 72 and 96 hours after siRNA transfection. GAPDH was used as a loading control. The immunoblot bands were quantified by densitometry, and the intensity of the NCS1, VDAC1, and SDHA bands was calculated (means ± SEM, n = 7 experiments). (K) Effect of NCS1 knockdown on ER-mitochondria contact. Bar graph shows the relative percentage of mitochondria in contact with ER (n = 257 mitochondria for siScr and n = 239 mitochondria for siNCS1). *P < 0.05, **P < 0.01, and ***P < 0.001.
Reexpression of NCS1 rescues the defective mitochondrial phenotype of WFS1-deficient cells
Given the effect of NCS1 on ER-[Ca2+]m uptake and respiratory chain function and to validate our hypothesis, we investigated whether overexpression of NCS1 in patient fibroblasts could rescue mitochondrial dysfunction. Transfection and subsequent expression of NCS1 in WFS1-deficient cells were confirmed by immunoblotting (Fig. 7A). Histamine-induced ER Ca2+ release was not statistically significant between patient cells overexpressing NCS1 and control cells (Fig. 7B and fig. S7A). NCS1 overexpression did not alter the rate of cytosolic Ca2+ rise but increased the rate of Ca2+ decay (Fig. 7, C and D). In addition, NCS1 overexpression in patient cells restored normal [Ca2+]m uptake (Fig. 7E and fig. S7B) by correcting the rate of [Ca2+]m entry and decay (Fig. 7, F and G). Moreover, NCS1 expression increased the number of VDAC1-IP3R1 interactions in MAMs by 40% in patient cells compared to control cells (Fig. 7H). Although overexpression of NCS1 in patient cells did not significantly modify the protein abundance of VDAC1 (Fig. 7I) and that of other proteins of the respiratory chain complexes (fig. S8, A to F), mitochondrial respiration analysis showed that the NCS1 overexpression in WFS1-deficient cells was associated with a statistically significant increase in complex I– and complex II–driven respiration (Fig. 7J). These data indicate that NCS1 rescue could protect WFS1-deficient cells and could be used as a tool to restore mitochondrial function.
Fig. 7 NCS1 overexpression restores mitochondrial function.
(A) Expression of NCS1 in patient fibroblasts transfected with Flag alone (Flag) or NCS1-Flag (NCS1). GAPDH was used as a loading control. (B) Maximum amplitude of histamine-stimulated Fluo-4 fluorescence in control fibroblasts (black dots; n = 23), P3 patient fibroblasts (red dots; n = 8), P3 patient fibroblasts expressing Flag alone (red squares; n = 9), and P3 patient fibroblasts expressing NCS1-Flag (red triangles; n = 8). (C) Rate of cytosolic Ca2+ rise in control fibroblasts (black dots), P3 patient fibroblasts (red dots), P3 patient fibroblasts expressing Flag alone (red squares), and P3 patient fibroblasts expressing NCS1-Flag (red triangles). Same n values as (B). (D) Rate of cytosolic Ca2+ decay in control fibroblasts (black dots), P3 patient fibroblasts (red dots), P3 patient fibroblasts expressing Flag alone (red squares), and P3 patient fibroblasts expressing NCS1-Flag (red triangles). Same n values as (B). (E) Maximum amplitude of histamine-stimulated pericam-mt fluorescence in control fibroblasts (black dots; n = 23), P3 patient fibroblasts (red dots; n = 12), P3 patient fibroblasts expressing Flag alone (red squares; n = 10), and P3 patient fibroblasts expressing NCS1-Flag (red triangles; n = 10). (F) Rate of [Ca2+]m entry in control (black dots), P3 patient (red dots), P3 patient treated with Flag alone (red squares), and P3 patient treated with NCS1-Flag (red triangles). Same n values as (E). (G) Rate of [Ca2+]m decay in control fibroblasts (black dots), P3 patient fibroblasts (red dots), P3 patient fibroblasts expressing Flag alone (red squares), and P3 patient fibroblasts expressing NCS1-Flag (red triangles). Same n values as (E). Means ± SEM; n = 8 to 28 cells. *P < 0.05, **P < 0.01, ***P < 0.001. (H) Representative PLA images (top) and quantitative analysis (bottom) of VDAC1-IP3R1 interactions in patient fibroblasts transfected with Flag alone or NCS1-Flag (NCS1; n = 94 and 156, respectively). ****P < 0.0001 compared to controls. Scale bar, 10 μm. (I) Effect of NCS1 overexpression on VDAC1 protein expression in patient fibroblasts with Flag alone (Flag) or NCS1-Flag (NCS1). GAPDH was used as a loading control. The immunoblot bands were quantified by densitometry, and VDAC1 intensity was calculated (means ± SEM; n = 6 experiments). (J) Complex I– and complex II–mediated respiration of patient fibroblasts transfected with Flag alone (Flag) or NCS1-Flag (NCS1) (means ± SEM; n = 4 experiments). **P < 0.01.
DISCUSSION
Our study provides new insights into mechanisms underlying the mitochondrial dysfunction in WFS1-deficient cells involving NCS1, a WFS1 interacting protein. We demonstrated that WFS1 interacted with the Ca2+ sensor NCS1 (39) and that NCS1 protein abundance was substantially decreased in WFS1-deficient cells. Addition of NCS1 to planar lipid bilayers or overexpression of NCS1 in PC12 cells results in increased IP3R channel activity (30–32), which should lead to increased Ca2+ release from the ER. In fibroblasts from our Wolfram syndrome patients, a decrease in Ca2+ flux from the ER was observed only under histamine application, whereas ER Ca2+ load was unchanged, suggesting a role of NCS1 in regulating IP3R activity. In addition, there was a concomitant decrease in the [Ca2+]m uptake. We further showed that ER-mitochondria structural interactions were reduced in patient cells, as evidenced by reduced interactions of two pairs of proteins at the MAM interface (VDAC1-IP3R1 and GRP75-IP3R1). This reduced tethering likely contributed to reduced Ca2+ transfer from ER to mitochondria in these patient cells, as has been previously described in MFN2-deficient cells that show a decrease in MAM tethering (28, 40). Dysregulation of Ca2+ homeostasis and impaired MAM formation have been previously described in WFS2-deficient cells (41, 42), suggesting a common pathophysiological mechanism. In many neurodegenerative diseases, the contact between ER and mitochondria is decreased (43), similar to what we observed in WFS1 patients. Specifically, in Alzheimer’s disease, the number of contacts is increased (44), similar to what is observed in Wolfram syndrome patients. Moreover, in amyotrophic lateral sclerosis type 8 (ALS8), the number of ER-mitochondria contacts is increased, and in ALS16 (45), the number is decreased. These two observations are similar to what is observed in Wolfram syndrome. It seems that the number of ER-mitochondria contacts depends on the protein affected. MAM structure and function are tightly regulated to ensure proper functioning of the mitochondria. Any modification in the structure and/or function will lead to a dysfunction of mitochondrial respiration that will have repercussions on cellular physiology.
Although the reduction of [Ca2+]m uptake was not attributed to a change in Δψm, it may have resulted from two factors: the decrease in the ER Ca2+ release and disorganization of the MAMs. One limitation of the study was the small number of patients from whom fibroblasts were analyzed because of the extreme rarity of the pathology. Rescue experiments for WFS1 failed to express the protein; however, down-regulation or overexpression of NCS1 in cells confirms the results, reinforcing the robustness of analyses of patient cells and suggesting that NCS1 is involved in Ca2+ and MAM dysregulation in Wolfram syndrome. The reduced [Ca2+]m uptake impairs the function of the mitochondria, because some dehydrogenases of the tricarboxylic acid (TCA) cycle require Ca2+ to maintain complex I and complex III and V function. There is no evidence that Ca2+ controls the abundance of complex II. Nakamura et al. (46) have found that the loss of NCS1 in mouse cardiomyocytes alters the proper functioning of the mitochondria by decreasing ATP levels and the respiratory rate, which is associated with a decrease in complex I, III, and V protein abundance. Consistent with this work, cells from our Wolfram syndrome patients showed a substantial decrease in mitochondrial respiratory chain activity. The decrease in complex II–driven respiration is probably linked to the decrease in complex II expression.
The clinical hallmark of Wolfram syndrome is the association of diabetes mellitus and optic atrophy, due respectively to degeneration of pancreatic β cells and retinal ganglion cells (47), two cell types in which NCS1 is abundant (48, 49). In β cells, NCS1 localizes mainly in the cytosol, ER, and secretory granules. NCS1 promotes the priming of the secretory granules for release and increases the number of granules residing in the readily releasable pool. This action is mediated by an activation of the phosphatidylinositol 4-kinase β (48). In MIN6 pancreatic β cells, WFS1 knockdown reduces insulin secretion, which is associated with a decrease in cytosolic Ca2+ (50). It is tempting to speculate that this dysfunction could be due to the possible loss of NCS1. In retinal ganglion cells, NCS1 may be involved in synaptogenesis during development and in synaptic transmission upon maturation (49). These two processes depend on energy supplied by mitochondria transported along axons to the synaptic terminals (51, 52) notably to maintain Ca2+ homeostasis (53, 54). Efficient axonal transport of mitochondria depends on their own energy production, because axonal transport is ATP dependent and, on cytoskeletal-mitochondrial interaction, because axonal transport is microtubule based (55). Our data, combined with the work of Cagalinec et al. (11), suggest that altered Ca2+ homeostasis due to decreased NCS1 protein abundance may be responsible for the impaired mitochondrial dynamics. Our data provide a potential explanation for the defective transport of mitochondria along the retinal ganglion cell axons because when NCS1 is diminished, as would be expected in WFS1-deficient retinal ganglion cells, mitochondrial function is impaired. This defect may be responsible for the retinal ganglion cell degeneration observed in Wolfram syndrome patients.
Wolfram syndrome patients may develop psychiatric disorders (56), such as bipolar disorder, schizophrenia, anxiety, and depression (57–59). Surprisingly, changes in NCS1 abundance provoke similar symptoms in human and mice (60, 61). It is tempting to speculate that the neurological and psychiatric disorders are linked to destabilization of NCS1 expression levels. In line with this, Schlecker et al. (31) have demonstrated that lithium promotes the interaction of IP3R and NCS1 and in bipolar disorder.
In this study, we demonstrated that WFS1, NCS1, and IP3R may be part of a complex associated with ER-mitochondria contact sites. In healthy cells, WFS1 interacts with NCS1 and may prevent its degradation. Our studies of the interactions between IP3R and NCS1 have used coimmunoprecipitation (31) or PLA (30), approaches that do not allow us to conclude whether the interaction is direct. It is possible that another protein interacts with either NCS1 or IP3R to bridge the two proteins. Nevertheless, the modulation of IP3R function by NCS1 favors a direct interaction. When the WFS1/NCS1/IP3R complex is functional, Ca2+ can be properly transferred from the ER to mitochondria and activate the TCA cycle and the mitochondrial respiratory chain. When WFS1 is lost, the WFS1/NCS1/IP3R complex is disrupted and NCS1 is partially degraded or destabilized. Consequently, altered ER-[Ca2+]m transfer leads to mitochondrial bioenergetic dysfunction that may result in the activation of cell death in the tissues affected by Wolfram syndrome (Fig. 8). We show that NCS1 overexpression in Wolfram syndrome patient fibroblasts reverses most of the observed defects. As a consequence, NCS1 may be a drug target for the treatment of Wolfram syndrome.
Fig. 8 Model of mitochondrial alteration induced by loss of function of WFS1.
(Left) In healthy cells, WFS1 binds to NCS1 to form a complex with IP3R1 to activate ER-mitochondria Ca2+ transfer to promote mitochondrial function. When the WFS1/NCS1/IP3R1 complex and VDAC1 are functional, Ca2+ can properly transfer from the ER to mitochondria and activate the TCA cycle and mitochondrial respiratory chain. (Right) In Wolfram syndrome, the loss of function of WFS1 leads to NCS1 degradation and decreased ER-mitochondria Ca2+ transfer. As a consequence, the mitochondrial respiratory chain is altered.
MATERIALS AND METHODS
Cell cultures
Fibroblasts were cultured from skin biopsies taken after obtaining informed consent from three individuals without diabetes mellitus and four affected patients carrying mutations in WFS1 gene, as previously described (62). Human embryonic kidney (HEK) 293T cells were cultured in MegaCell Dulbecco’s Modified Eagle’s Medium (Sigma-Aldrich, St. Louis, MO, USA) supplemented with 10% fetal bovine serum (Sigma-Aldrich) and 20 mM penicillin/streptomycin (Sigma-Aldrich).
Cell transfection
Control fibroblasts were transfected with siRNA directed against WFS1 (ON-TARGETplus Human WFS1 siRNA SMARTpool, Thermo Fisher Scientific Biosciences, Waltham, MA, USA) and/or with siRNA directed against NCS1 (ON-TARGETplus Human NCS1 siRNA SMARTpool, Thermo Fisher Scientific Biosciences) using Lipofectamine 2000 (Invitrogen, Carlsbad, USA) according to the manufacturer’s instructions and processed 72 or 96 hours later for Western blot or oxygraphy analyses. Transfection with ON-TARGET Non-Targeting Pool (siScr) was used as controls.
To overexpress NCS1-Flag or pericam-mt vectors, fibroblasts were transfected by nucleofection. Nucleofection was performed according to an optimized protocol provided by the manufacturer (Amaxa Biosystems, Cologne, Germany). Cells were suspended in 100 μl of nucleofactor solution (Amaxa Biosystems); mixed with pericam-mt, Flag, or NCS1-Flag construct; and subjected to nucleofection, after which cells were transferred into prewarmed fresh medium. Cells were analyzed 24 hours after nucleofection.
Real-time RT-PCR
Real-time PCR was performed on total RNA extracted from cells using the RNeasy Mini Kit (Qiagen, Venlo, Netherlands) and reverse transcribed with the SuperScript III First-Strand Kit (Invitrogen) according to the manufacturer’s instructions.
Measurements of enzymatic activity and protein content
The activity of the mitochondrial respiratory chain complexes and respiratory rates was measured on cell homogenates, as described previously (62). The activity of the mitochondrial respiratory chain complexes was measured with cell homogenates in a cell buffer [250 mM saccharose, 20 mM tris(hydroxymethyl)aminomethane, 2 mM EGTA, and bovine serum albumin (BSA; 1 mg/ml) (pH 7.2)] at 37°C using a UV-SAFAS spectrophotometer (SAFAS). Cellular protein content was determined with the BCA (bicinchoninic assay) Kit (Pierce) using BSA as a standard. Complex I (NADH ubiquinone reductase, EC 1.6.5.3) activity was measured according to a procedure described elsewhere (63) and adapted using 2,6-dichloroindophenol (DCPIP) to avoid the inhibition of complex I activity by decylubiquinol (64). Cells were disrupted by two freezing-thawing cycles, washed, centrifuged for 1 min at 16,000g, and resuspended in cell buffer (50 μl per 106 cells). Complex II (succinate ubiquinone reductase) activity was measured according to James and colleagues (65). Specific enzymatic activities of complexes I and II were expressed in mIU (nanomoles of DCPIP per minute per milligram of protein). Complex IV (cytochrome c oxidase) activity was recorded according to a method that Rustin and colleagues (66) adapted in a 50 mM KH2PO4 buffer, using 15 μM reduced cytochrome c. Specific enzymatic activity was expressed in mIU (nanomoles of cytochrome c per minute per milligram of protein). CS activity was assayed by a standard procedure (67). Specific enzymatic activity was expressed in mIU [nanomoles of 5-5′-dithiobis(2-nitrobenzoic acid) per minute per milligram of protein].
Oxygen consumption
The respiratory rate was measured using cells resuspended in respiratory buffer [0.5 mM EGTA, 3 mM MgCl2⋅6H2O, 60 mM K-lactobionate, 20 mM taurine, 10 mM KH2PO4, 20 mM Hepes, 110 mM sucrose, and BSA (1 mg/ml) (pH 7.1)] and permeabilized by incubation with digitonin (15 μg per 106 cells). The respiratory rates of 3 × 106 to 5 × 106 cells were recorded at 37°C in 2-ml glass chambers using a high-resolution Oxygraph respirometer (Oroboros). Malate (5 mM) and pyruvate (5 mM) were added to provide NADH to complex I. The decarboxylation of pyruvate is catalyzed by pyruvate dehydrogenase (PDH), producing acetyl coenzyme A. Malate dehydrogenase (MDH) located in the mitochondrial matrix induces the oxidation of malate to oxaloacetate. The condensation of oxaloacetate and acetyl coenzyme A induces the formation of citrate by CS. Thus, the addition of malate and pyruvate results in the production of NADH by four enzymes: MDH, PDH, isocitrate dehydrogenase, and α-ketoglutarate dehydrogenase. Because of the activation of different malate shuttles, complex II is not activated. Citrate and α-ketoglutarate are depleted by antiport transport with malate. Thus, the addition of malate and pyruvate defines the respiration driven by complex I used to compensate the physiological proton leak. Activation of ATP synthesis was induced by the addition of 1.5 mM adenosine 5′-diphosphate (ADP). Thus, we obtained the respiration driven by complex I coupled to the ATP synthesis. The addition of succinate (10 mM) allowed reconstitution of TCA cycle function with the activation of succinate dehydrogenase. Thus, maximal respiration of the mitochondrial chain induced by the competition of the respiration driven by complexes I and II was determined. Addition of rotenone (10 μM) inhibited the electron transfer from complex I to coenzyme Q and allowed the measurement of respiration driven by complex II. Oligomycin supplementation (8 μg/ml) inhibited the ATP synthesis, and respiration uncoupled driven by complex II was determined. Last, FCCP (1 μM) was added to test the quality of the permeabilization of the plasma membrane by digitonin.
Western blot analysis
Protein abundance was detected by immunoblot using commercially available antibodies and revealed using chemiluminescence. Frozen fibroblast pellets were lysed by a hypoosmotic shock (10 μl of H2O per 106 cells). For coimmunoprecipitation, HEK293T pellets were mixed with 500 μl of lysis buffer consisting of 0.1 M NaCl, 1% Triton X-100, 1 mM phenylmethylsulfonyl fluoride (PMSF; Fluka), 1× SIGMAFAST protease inhibitor cocktail (Sigma-Aldrich), 5 mM EDTA, and 20 mM tris and rotated overnight at 4°C. The cellular protein content was determined with the BCA Kit.
Thirty micrograms of total fibroblasts or HEK293T-transfected protein was separated on SDS–polyacrylamide gel electrophoresis (PAGE) and transferred onto polyvinylidene difluoride membranes (Bio-Rad). Membranes were saturated with 5% nonfat milk dissolved in 0.1% tris-buffered saline (TBS)–Tween (pH 7.4, 137 mM NaCl, 2.7 mM KCl, 23 mM tris, and 0.1% Tween) for 1 hour at room temperature and incubated overnight at 4°C with polyclonal rabbit anti-WFS1 (1:500; Cell Signaling, USA), anti-MCU (1:1000; Abcam, UK), anti-MFN2 (1:1000; Cell Signaling), polyclonal chicken anti-NCS1 (1:1000; United States Biological, USA), monoclonal mouse anti-NDUFA13 (1:1000; Abcam), anti-SDHA (1:1000; Abcam), anti-UQCRC2 (1:1000; Abcam), anti-MTCO1 (1:1000; Abcam), anti–ATP synthase (1:1000; Invitrogen), anti-VDAC1 (1:1000; Abcam), anti-GAPDH (1:1000; Abcam), and anti-GRP75 (1:1000; Abcam). Anti-IP3R antibody was a gift from J. Parys (Leuven, Belgium). Membranes were then washed three times in 0.1% TBS-Tween and incubated with anti-rabbit immunoglobulin G (IgG), anti-chicken IgG, or anti-mouse IgG horseradish peroxidase–linked antibody (1:10,000; Sigma-Aldrich) for 1 hour at room temperature. Immunoreactive proteins were visualized with enhanced chemiluminescence (ECL+ Western Blotting Detection Reagents, Amersham Biosciences, UK). Band intensities were quantified with ImageJ [National Institutes of Health (NIH), USA; http://rsb.info.nih.gov/ij].
In situ PLA
Yeast two-hybrid screening
Yeast two-hybrid screening was performed by Hybrigenics Services SAS, Paris, France (www.hybrigenics-services.com). The coding sequence for Mus musculus–Wfs1 (amino acids 1 to 311; GenBank accession number GI:28422738) was PCR amplified and cloned into pB29 as a C-terminal fusion to LexA (N-Wfs1-LexA-C). The construct was checked by sequencing the entire insert and used as a bait to screen a random-primed mouse inner ear cDNA library constructed into pP6. pB29 and pP6 derive from the original pBTM116 (69, 70) and pGADGH (71) plasmids, respectively. Clones (54.3 million) (fivefold the complexity of the library) were screened using a mating approach with YHGX13 (Y187 ade2-101:loxP-kanMX-loxP, matα) and L40ΔGal4 (matα) yeast strains, as previously described (72). Twenty-nine His+ colonies were selected on a medium lacking tryptophan, leucine, and histidine. The prey fragments of the positive clones were amplified by PCR and sequenced at their 5′ and 3′ junctions. The resulting sequences were used to identify the corresponding interacting proteins in the GenBank database (National Center for Biotechnology Information) using a fully automated procedure. A confidence score [Predicted Biological Score (PBS)] was attributed to each interaction, as previously described (73).
The PBS relies on two different levels of analysis. First, a local score takes into account the redundancy and independency of prey fragments, as well as the distribution of reading frames and stop codons in overlapping fragments. Second, a global score takes into account the interactions found in all the screens performed at Hybrigenics using the same library. This global score represents the probability of an interaction being nonspecific. For practical purposes, the scores were divided into four categories, from A (highest confidence) to D (lowest confidence). A fifth category (E) specifically flags interactions involving highly connected prey domains previously found several times in screens performed on libraries derived from the same organism. Last, several of these highly connected domains have been confirmed as false positives of the technique and are now tagged as F. The PBS scores have been shown to positively correlate with the biological relevance of the interactions (74, 75).
Confocal imaging
All confocal experiments were performed on fibroblasts placed on the stage of a Zeiss LSM 510 inverted confocal microscope (Zeiss, Le Pecq, France). Pericam-mt/pcDNA was used to measure [Ca2+]m. To measure cytosolic Ca2+, fibroblasts were loaded with Fluo-4 AM (5 μM; Molecular Probes), Fluo-4 signal was obtained by excitation at 488 nm, and emitted light was collected at 515 nm. For pericam-mt, the excitation wavelength upon Ca2+ binding switches from 415 to 488 nm. Thus, the transfected cells were excited at 488, and images were collected at 515 nm. All the collected images were processed with ImageJ (NIH, USA; http://rsb.info.nih.gov/ij/). All confocal experiments were performed on fibroblasts placed on the stage of a Zeiss LSM 510 inverted confocal microscope (Zeiss) equipped with a 63× lens [oil immersion; numerical aperture (NA), 1.2].
ER Ca2+ release
Luminal Ca2+ dynamics were measured using the Ca2+-sensitive FRET-based cameleon protein D1-ER (76). Cells were cultured on 24-mm coverslips and transfected with D1-ER. Cells were imaged on a Zeiss Axiovert 200M microscope, equipped with a 40× oil objective (NA, 1.3). Emission ratio imaging of D1-ER was accomplished by using a 436DF20 excitation filter, a 450-nm dichroic mirror, and two emission filters [475/40 for enhanced cyan fluorescent protein (ECFP) and 535/25 for citrine]. Exposure times were 100 ms, and images were taken every 1 to 2 s. The FRET signal (yellow fluorescent protein/CFP) was normalized to CFP emission intensity, and changes in ER Ca2+ were expressed as the ratio of the emissions at 535 and 470 nm. To induce Ca2+ release from ER, the cells were challenged with an agonist that, through interaction with G protein–coupled receptors, evokes a rapid discharge from IP3Rs.
Ca2+ measurements with aequorin constructs
For mt-AEQ measurements, coverslips with control and mutated fibroblasts overexpressing MCU were incubated for 2 hours in KRB (Krebs-Ringer modified buffer) (135 mM NaCl, 5 mM KCl, 0.4 mM KH2PO4, 1 mM MgSO4, 5.5 mM glucose, and 20 mM Hepes (pH 7.4)] at 37°C supplemented with wild-type coelenterazine (5 M) and then transferred to the perfusion chamber. In the experiments with permeabilized cells, cells were perfused in intracellular buffer [140 mM KCl, 10 mM NaCl, 1 mM KH2PO4, 5.5 mM glucose, 2 mM MgSO4, 1 mM ATP, 2 mM Na+ succinate, and 20 mM Hepes (pH 7.05)] at 37°C. Cells were permeabilized through a 1-min perfusion with 100 μM digitonin during luminescence measurements. After a 2-min washing (to remove digitonin), [Ca2+]m uptake was estimated by mt-AEQ after cell perfusion with the same intracellular solution without EGTA and containing Ca2+ at different concentrations (1 and 4 μM). Calcium uptake speed was calculated as the first derivative by using the first derivative function smoothed for three time points.
To reconstitute erAEQ with high efficiency, luminal [Ca2+] of the ER was first reduced by incubating the cells for 45 min at 4°C in KRB supplemented with 5 μM coelenterazine, the Ca2+ ionophore ionomycin, and 600 μM EGTA. After incubation, the cells were extensively washed with KRB supplemented with 2% BSA and 2 mM EGTA before the luminescence measurement was initiated. Aequorin signals were measured in KRB supplemented with either 1 mM CaCl2 or 100 μM EGTA, using a purpose-built luminometer. The agonist (bradykinin at 100 μM) was added to the same medium.
Mitochondrial membrane potential
To measure Δψm, fibroblasts were loaded with 10 nM TMRM (Life Technologies) for 20 min at 37°C, followed by washout. TMRM dyes were excited at 565 nm, and emitted light was collected at 585 nm. At the end of each experiment using TMRM dyes, cells were exposed to the mitochondrial uncoupler FCCP (1 μM) to fully dissipate (Δψm) and to determine the dynamic range of the dye. TMRM fluorescence was normalized to the fluorescence signals obtained after FCCP application. After loading, cells were placed on the stage of a Zeiss LSM 510 inverted confocal microscope (Zeiss) equipped with a 63× lens (oil immersion; NA, 1.2). Collected images were processed with ImageJ (NIH, USA; http://rsb.info.nih.gov/ij/). Detection of Δψm was also performed by loading cells with a JC-1 staining kit (Thermo Fisher Scientific). JC-1 emissions were collected using a Zeiss LSM 510 confocal microscope equipped with a 40× oil objective (NA, 1.30). The green fluorescent/red fluorescent intensity ratios (540/595 nm) were calculated to determine Δψm levels.
Coimmunoprecipitation
pcDNA-IP3R1 was provided by K. Mikoshiba (Okasaki, Japan) (77). The plasmid expressing Wfs1 was obtained by subcloning Wfs1 into myc-tagged pCS2 + MT using Hind III and Bam H1. Ncs1 was subcloned into C-terminal p3XFLAG-CMV between Eco R1 and Bam H1. To assess interactions between IP3R1 and Ncs1-Flag and between Wfs1-myc and Ncs1-Flag, 100 μg of proteins from each transfection reaction was mixed with 2 μl of mouse anti-Flag M2 antibody (Sigma-Aldrich) containing 1 mM PMSF and 2% BSA in DWBa buffer [100 mM NaCl, 20 mM tris, 1 mM EDTA (pH 7.4), 1% Triton X-100]. For IP3R1 and Wfs1-myc antibody, 100 μg of proteins from each transfection reaction was mixed with 2 μg of a rabbit anti-myc (Sigma-Aldrich). The mixtures were vortexed and incubated at 4°C overnight. Thirty microliters of Sepharose A and 10 μl of Sepharose G (Immunoprecipitation Starter Pack, GE Healthcare) were used per sample and prepared following supplier instruction. Sepharose suspension was added to the tubes containing antibody/protein samples and mixed on a rotating wheel at 4°C for 1 hour. The samples were then centrifuged at 2000 rpm for 1 min and washed three times with 1 ml of DWBa and two times with 1 ml of DWBb [100 mM NaCl, 20 mM tris, and 1 mM EDTA (pH 7.4)] each. The pellets obtained after the last wash were resuspended in 25 μl of Laemmli sample buffer (Bio-Rad). Last, all the samples were boiled at 95°C for 5 min and centrifuged at 13,200 rpm for 1 min. The supernatants were loaded on an SDS-PAGE gel for Western blotting and analyzed with an anti-myc antibody for Wfs1-myc and Ncs1-Flag coimmunoprecipitation, with the rabbit Rbt476 pan anti-IP3R antibody (a gift from J. Parys, Leuven, Belgium) (78) for Ncs1-Flag and IP3R1 coimmunoprecipitation and Wfs1-myc and IP3R1 coimmunoprecipitation. Thirty micrograms of protein from Ncs1-Flag and Wfs1-myc transfection reactions was used as controls.
Transmission electron microscopy
Cells were immersed in a solution of 2.5% glutaraldehyde in PHEM buffer (1×, pH 7.4) overnight at 4°C, rinsed in PHEM buffer, and postfixed in a 0.5% osmic acid for 2 hours in the dark and at room temperature. After two rinses in PHEM buffer, cells were dehydrated in a graded series of ethanol solutions (30 to 100%) and were embedded in EMbed 812 using an Automated Microwave Tissue Processor for Electronic Microscopy (Leica). Ultrathin sections (70 nm; Leica-Reichert Ultracut E) were collected at different levels of each block. These sections were stained with uranyl acetate and lead citrate before examination in a Tecnai F20 TEM at 200 kV in the CoMET Montpellier Rio Imaging (MRI) facilities, Institute for Neurosciences of Montpellier, France. To evaluate the frequency of contact between the mitochondria and the ER in the different fibroblasts, we randomly took 20 images of cytoplasmic area from 20 different cells in each sample with a JEOL 1400 TEM at ×15,000 magnification. In each image, we counted the number of mitochondria (403 in control cells and 649 in patient cells for a total of 1052 mitochondria) and calculated the proportion of mitochondria in close contact with ER (<30 nm). Moreover, the perimeter of each mitochondria and the proportion of the mitochondrial surface closely associated with ER were calculated. For experiments with knockdown of NCS1, 257 mitochondria were analyzed for siScr and 239 mitochondria were analyzed for siRNA NCS1.
Statistical analysis
The Mann-Whitney was used to compare the fibroblasts from WFS1 patients and controls. The Kruskal-Wallis test was used to compare the effect of the overexpression of NCS1 or vector alone. Differences were considered significant at *P < 0.05, **P < 0.01, and ***P < 0.001. Proportions of ER-mitochondrial contacts between controls and patient fibroblasts were compared with their 95% confidence intervals.
Acknowledgments
We thank the patients and their families for their contribution to the clinical studies. We are indebted to the MRI for their technical support. We thank J. Parys for providing the pan IP3R Rbt476 antibody, K. Mikoshiba for providing the IP3R1 pcDNA vector, and N. Bendridi and M.-A. Chauvin for technical assistance. We thank A. Miyawaki who gifted us with the ratiometric pericam-mt plasmid. We thank V. Paquis-Flucklinger, C. Rouzier, and A. Chaussenot for their contributions. We thank P. Carroll for his contribution in reading the manuscript. We thank S. Cubaynes for help with statistics. Funding: We are indebted to the INSERM and Université Montpellier for providing institutional support and to the patient associations Union National des Aveugles et Déficients Visuels (UNADEV) and Association Syndrome de Wolfram for their financial support. This work was supported by grants from the Agence Nationale pour la Recherche (ANR-12-JSV1-0008-01), Fondation pour la Recherche Médicale, and Fondation de France. C.A. was supported by the Fondation de France. Author contributions: C.A. performed most of the experiments, analyzed the data, and contributed to writing the manuscript. J.F. performed Ca2+ imaging and wrote the paper. S.P. and A.D. performed aequorin experiments. J.R. performed in situ PLA and mitochondrial content quantification. C.A.A. and J.K. carried out coimmunoprecipitation experiments. J.J., C.M., M.Q., and D.B.-W. helped in the characterization of cells. C.C. and M.T. performed TEM. D.M. and C.H. provided reagents. P.P. analyzed data. A.L. provided reagents and analyzed the data. B.D. and C.D. conceived and designed the study, analyzed the data, and wrote the paper. Competing interests: The authors declare that they have no competing interests. Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper or the Supplementary Materials.
Supplementary Material
Summary
Fig. S1. WFS1 interacts with IP3R.
Fig. S2. Ca2+ imaging in thapsigargin-treated cells.
Fig. S3. Analysis of mitochondrial respiratory rate.
Fig. S4. NCS1 interacts with WFS1 and IP3R.
Fig. S5. Time course of siRNA-mediated knockdown of WFS1.
Fig. S6. Time course of siRNA-mediated knockdown of NCS1.
Fig. S7. Representative cytosolic and [Ca2+]m in control and patient fibroblasts expressing Flag or NCS1-Flag.
Fig. S8. Mitochondrial protein abundance in patient fibroblasts overexpressing NCS1.
Table S1. Clinical features of control subjects and patients with Wolfram syndrome.
Table S2. TEM image analysis results.
Table S3. Two-hybrid screening results.
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Science Signaling
Volume 11 | Issue 553
October 2018
October 2018
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Received: 5 October 2017
Accepted: 4 October 2018
Acknowledgments
We thank the patients and their families for their contribution to the clinical studies. We are indebted to the MRI for their technical support. We thank J. Parys for providing the pan IP3R Rbt476 antibody, K. Mikoshiba for providing the IP3R1 pcDNA vector, and N. Bendridi and M.-A. Chauvin for technical assistance. We thank A. Miyawaki who gifted us with the ratiometric pericam-mt plasmid. We thank V. Paquis-Flucklinger, C. Rouzier, and A. Chaussenot for their contributions. We thank P. Carroll for his contribution in reading the manuscript. We thank S. Cubaynes for help with statistics. Funding: We are indebted to the INSERM and Université Montpellier for providing institutional support and to the patient associations Union National des Aveugles et Déficients Visuels (UNADEV) and Association Syndrome de Wolfram for their financial support. This work was supported by grants from the Agence Nationale pour la Recherche (ANR-12-JSV1-0008-01), Fondation pour la Recherche Médicale, and Fondation de France. C.A. was supported by the Fondation de France. Author contributions: C.A. performed most of the experiments, analyzed the data, and contributed to writing the manuscript. J.F. performed Ca2+ imaging and wrote the paper. S.P. and A.D. performed aequorin experiments. J.R. performed in situ PLA and mitochondrial content quantification. C.A.A. and J.K. carried out coimmunoprecipitation experiments. J.J., C.M., M.Q., and D.B.-W. helped in the characterization of cells. C.C. and M.T. performed TEM. D.M. and C.H. provided reagents. P.P. analyzed data. A.L. provided reagents and analyzed the data. B.D. and C.D. conceived and designed the study, analyzed the data, and wrote the paper. Competing interests: The authors declare that they have no competing interests. Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper or the Supplementary Materials.
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Montpellier University
the Institut National de la Santé et de la Recherche Médicale
Union National des Aveugles et Déficients Visuels (UNADEV)
Association Syndrome de Wolfram
gence Nationale pour la Recherche: ANR-12-JSV1-0008-01
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