Mitochondria play an essential role in regulating insulin secretion from beta cells by providing the ATP needed for the membrane depolarization that results in voltage-dependent Ca2+ influx and subsequent insulin granule exocytosis. Ca2+, in turn, is also rapidly taken up by the mitochondria and exerts important feedback regulation of metabolism. The aim of this study was to determine whether the distribution of mitochondria within beta cells is important for the secretory capacity of these cells. We find that cortically localized mitochondria are abundant in rodent beta cells, and that these mitochondria redistribute towards the cell interior following depolarization. The redistribution requires Ca2+-induced remodeling of the cortical F-actin network. Using light-regulated motor proteins, we increased the cortical density of mitochondria twofold and found that this blunted the voltage-dependent increase in cytosolic Ca2+ concentration and suppressed insulin secretion. The activity-dependent changes in mitochondria distribution are likely to be important for the generation of Ca2+ microdomains required for efficient insulin granule release.

Pancreatic beta cells sense increases in the surrounding glucose concentration and secrete glucose-lowering insulin to counteract the elevation. The mechanism by which glucose controls insulin secretion involves uptake across the plasma membrane, followed by glycolysis and mitochondrial oxidative phosphorylation to produce ATP. This nucleotide closes ATP-sensitive K+ channels in the plasma membrane, leading to depolarization, opening of voltage-dependent Ca2+ channels and a rise in cytosolic Ca2+ that triggers insulin secretion (Rorsman and Ashcroft, 2018). Mitochondrial metabolism is thus required for glucose-stimulated insulin secretion, but this organelle also regulates insulin secretion downstream of ATP production by controlling the cytosolic Ca2+ concentration (Wiederkehr and Wollheim, 2008). The outer mitochondrial membrane is Ca2+ permeable, and Ca2+ is further transported into the mitochondrial matrix by the mitochondrial Ca2+ uniporter (mCU), in a process that is driven by the negative membrane potential (Patron et al., 2013). The mCU has a low affinity for Ca2+, and its activity under resting conditions is very low, but it can be activated by elevations in the cytosolic Ca2+ concentration and mitochondria contribute to Ca2+ homeostasis by acting as buffers. For the mCU to work efficiently, it needs Ca2+ in concentrations in excess of 10 µM. Such high concentrations are rarely seen in the bulk cytosol, but are frequently reached at sites of Ca2+ release or influx (Parekh, 2008). In recent years, numerous studies have shown that mitochondria localized close to Ca2+-release channels of the endoplasmic reticulum (ER) or voltage-dependent Ca2+ channels of the plasma membrane are exposed to much higher Ca2+ concentrations than those localized further away and can buffer changes in cytosolic Ca2+ (Giacomello et al., 2010; Malli et al., 2003; Quintana et al., 2011). Mitochondria are highly dynamic in cells, and their motility is largely dependent on GTPase-mediated transport along microtubules. The distribution of mitochondria in the cell is nonhomogenous, and it may also vary over time. This is particularly evident in cells with an asymmetric morphology, such as neurons. In these cells, mitochondria transport to axonal segments is required for local ATP production, which, in turn, is used for synaptic vesicle release (Verstreken et al., 2005). However, the presence of mitochondria in the presynaptic compartment also negatively regulates vesicle release by local buffering of synaptic Ca2+ elevations triggered by depolarization, in a process that is regulated by neuronal activity (Vaccaro et al., 2017). The basic mechanism of synaptic vesicle exocytosis and insulin granule exocytosis is the same, but whether mitochondria play a direct, local role in the regulation of insulin release is not known. Most studies on the interplay between mitochondria and Ca2+ have focused on the regulation of mitochondrial metabolism by the ion. The increase in Ca2+ that occurs in the mitochondrial matrix following voltage-dependent Ca2+ influx has been shown to have both stimulatory and inhibitory effects on ATP production (Alam et al., 2012; Kennedy et al., 1999; Krippeit-Drews et al., 2000; Li et al., 2013; Wiederkehr et al., 2011). Since insulin granules require microdomains of high Ca2+ for efficient exocytosis, it is possible that mitochondria play a more direct role in controlling insulin secretion by exerting a local buffering effect. In this study, we show that beta-cell mitochondria are abundant in the subplasmalemmal space under resting conditions, but that the density decreases following Ca2+ influx, in a process that requires intact cortical actin. Using light-driven molecular motors, we increased the number of subplasmalemmal mitochondria and found that this results in both reduced cytosolic Ca2+ elevations and exocytosis following depolarization.

Near-plasma membrane mitochondria redistribute in response to voltage-dependent Ca2+ influx

Total internal reflection fluorescence (TIRF) microscopy imaging of MIN6 beta cells expressing mitochondria-targeted mApple (mApple-Tomm20-N1) revealed that 18±2% of the subplasma membrane area was occupied by mitochondria. Most mitochondria exhibited dynamic movements, but there were sites that were occupied by mitochondria throughout the time course of the experiments. Similar observations have been made in neurons, in which only one-third of the mitochondrial population in axons is mobile (Kang et al., 2008; Sheng and Cai, 2012). To test whether the cortical mitochondria constitute a stable population in beta cells, we stimulated the cells with 20 mM glucose, the main physiological stimulus of insulin secretion. This stimulation was without effect on the overall cortical mitochondria density (Fig. 1A,B). However, simultaneous recordings of the intracellular Ca2+ concentration with GCaMP5G revealed that the cortical mitochondria exhibited frequent transient displacements that mostly (83%) coincided with temporary increases in the cytosolic Ca2+ concentration (Fig. 1C–F). To more directly test the effect of Ca2+ on cortical mitochondria, we depolarized the cells with 30 mM KCl. We found that the cortical mitochondria density was significantly reduced in depolarized cells (from 18±2% to 9±1%, n=22; P<0.01) (Fig. 1G,H). Indeed, depolarization induced a rapid 13±2% (n=32; P<0.01) reduction in subplasma membrane mitochondria fluorescence within 1 min (Fig. 1I), indicating that the localization of mitochondria within beta cells is regulated under conditions that promote insulin secretion.

Fig. 1.

Characterization and modulation of subplasma membrane mitochondria. (A) TIRF microscopy images of a MIN6 cell expressing mApple-Tomm20 before and during stimulation with 20 mM glucose. (B) Quantification of the subplasma membrane density change following 15 min exposure to 20 mM glucose (n=36 cells; P>0.1, paired two-tailed Student's t-test). (C) TIRF microscopy recording of the cytosolic Ca2+ concentration (GCaMP5G, black) and cortical mitochondria (mApple-Tomm20, red) in a glucose-stimulated MIN6 cell. (D) Means±s.e.m. for changes in cytosolic Ca2+ concentration (black) and subplasma membrane mitochondria density (red) during glucose-induced Ca2+ oscillations (n=40 oscillations from 13 cells). (E,F) Relative change in subplasma membrane mitochondria density following glucose-induced Ca2+ oscillations (n=40 oscillations from 13 cells; P<0.01, paired two-tailed Student's t-test). (G) TIRF microscopy images showing displacement of mitochondria following depolarization with KCl. (H) Quantification of TIRF microscopy images of MIN6 cells expressing mApple-Tomm20 under control conditions (basal) and following depolarization with 30 mM K+ (n=22 cells; P<0.001, paired two-tailed Student's t-test). (I) Average trace showing the time course of KCl-induced mitochondrial displacement from the plasma membrane (mean±s.e.m., n=32 cells). (J) Top: TIRF microscopy images of the time-dependent recruitment of mitochondria to the plasma membrane in response to illumination. Bottom: schematic of the optogenetic principle. Mitochondria are tagged with CRY2, while Kif1a is tagged with CIBN. Kif1a travels along the microtubules in the anterograde direction to the plasma membrane. Upon blue-light illumination, CRY2 and CIBN bind to each other, resulting in mitochondria transportation towards the plasma membrane. (K) Quantifications of the response in J (means±s.e.m., ncontrol=18, nopto-mito=14; P=0.0092, unpaired two-tailed Student's t-test). # denotes the timepoint after which there is a stable, statistically significant (P<0.05) difference between the two groups. Control cells did not express the Kif1a plasmid and were exposed to blue light in the same way as the optogenetic group. Opto-mito, cells expressing Tomm20-CRY2 and CIBN-Kif1a. (L) Quantification of mitochondrial density at the plasma membrane before illumination, after 1 h of blue-light illumination and after 2 h recovery in the absence of illumination (control=10.8±0.0.7683%, light=19.17±1.139%, 2 h recovery=11.9±0.9286%; ncontrol=112, nlight=114, n2h recovery=75; P<0.0001, one-way ANOVA, followed by Tukey's multiple comparison test, Pcontrol/light<0.0001, Plight/2h recovery<0.0001). (M) Nocodazol (1 µM) inhibits microtubule polymerization and prevents light-induced mitochondrial transport (control=12.67%±0.5255, light=16.58%±0.689, nocodazol=11.48%±0.5533, light+nocodazol=13.31±0.7077; ncontrol=172, nlight=176, nnocodazol=144, nnocodazol+light=127; P<0.0001, one-way ANOVA, followed by Tukey's multiple comparison test, Pcontrol/light<0.0001, Plight/nocodazol<0.0001, Plight/nocodazol+light=0.0018). PM, plasma membrane. Dashed lines in panels A,G,J define cell outline.

Fig. 1.

Characterization and modulation of subplasma membrane mitochondria. (A) TIRF microscopy images of a MIN6 cell expressing mApple-Tomm20 before and during stimulation with 20 mM glucose. (B) Quantification of the subplasma membrane density change following 15 min exposure to 20 mM glucose (n=36 cells; P>0.1, paired two-tailed Student's t-test). (C) TIRF microscopy recording of the cytosolic Ca2+ concentration (GCaMP5G, black) and cortical mitochondria (mApple-Tomm20, red) in a glucose-stimulated MIN6 cell. (D) Means±s.e.m. for changes in cytosolic Ca2+ concentration (black) and subplasma membrane mitochondria density (red) during glucose-induced Ca2+ oscillations (n=40 oscillations from 13 cells). (E,F) Relative change in subplasma membrane mitochondria density following glucose-induced Ca2+ oscillations (n=40 oscillations from 13 cells; P<0.01, paired two-tailed Student's t-test). (G) TIRF microscopy images showing displacement of mitochondria following depolarization with KCl. (H) Quantification of TIRF microscopy images of MIN6 cells expressing mApple-Tomm20 under control conditions (basal) and following depolarization with 30 mM K+ (n=22 cells; P<0.001, paired two-tailed Student's t-test). (I) Average trace showing the time course of KCl-induced mitochondrial displacement from the plasma membrane (mean±s.e.m., n=32 cells). (J) Top: TIRF microscopy images of the time-dependent recruitment of mitochondria to the plasma membrane in response to illumination. Bottom: schematic of the optogenetic principle. Mitochondria are tagged with CRY2, while Kif1a is tagged with CIBN. Kif1a travels along the microtubules in the anterograde direction to the plasma membrane. Upon blue-light illumination, CRY2 and CIBN bind to each other, resulting in mitochondria transportation towards the plasma membrane. (K) Quantifications of the response in J (means±s.e.m., ncontrol=18, nopto-mito=14; P=0.0092, unpaired two-tailed Student's t-test). # denotes the timepoint after which there is a stable, statistically significant (P<0.05) difference between the two groups. Control cells did not express the Kif1a plasmid and were exposed to blue light in the same way as the optogenetic group. Opto-mito, cells expressing Tomm20-CRY2 and CIBN-Kif1a. (L) Quantification of mitochondrial density at the plasma membrane before illumination, after 1 h of blue-light illumination and after 2 h recovery in the absence of illumination (control=10.8±0.0.7683%, light=19.17±1.139%, 2 h recovery=11.9±0.9286%; ncontrol=112, nlight=114, n2h recovery=75; P<0.0001, one-way ANOVA, followed by Tukey's multiple comparison test, Pcontrol/light<0.0001, Plight/2h recovery<0.0001). (M) Nocodazol (1 µM) inhibits microtubule polymerization and prevents light-induced mitochondrial transport (control=12.67%±0.5255, light=16.58%±0.689, nocodazol=11.48%±0.5533, light+nocodazol=13.31±0.7077; ncontrol=172, nlight=176, nnocodazol=144, nnocodazol+light=127; P<0.0001, one-way ANOVA, followed by Tukey's multiple comparison test, Pcontrol/light<0.0001, Plight/nocodazol<0.0001, Plight/nocodazol+light=0.0018). PM, plasma membrane. Dashed lines in panels A,G,J define cell outline.

To obtain control over mitochondria distribution, we developed an optogenetic tool to guide mitochondria to the plasma membrane based on the CRY2-CIBN dimerization system. Briefly, we fused the anterograde motor protein Kif1a to CIBN and co-expressed it with GFP-tagged Tomm20 fused to CRY2. Blue-light illumination promotes reversible CRY2-CIBN interaction, resulting in anterograde transport of mitochondria along microtubules (Fig. 1J). Blue-light illumination of MIN6 cells expressing these tools resulted in a time-dependent increase in subplasma membrane mitochondria density, with a maximal increase of up to 100% (Fig. 1J,K). This increase was reversed following interruption of the illumination for 2 h (Fig. 1L). Moreover, inhibition of microtubule polymerization with nocodazole prevented the recruitment of mitochondria to the plasma membrane upon illumination, demonstrating that the mitochondrial accumulation is due to active transport (Fig. 1M).

Accumulation of cortical actin in response to voltage-dependent Ca2+ influx controls subplasma membrane mitochondrial density

Beta cells, like most cells, have an extensive cortical actin network that can limit organelle accessibility to the plasma membrane (Hsieh et al., 2017; Orci et al., 1972; Papadopulos et al., 2015). To visualize cortical actin, we expressed an mCherry-tagged version of the calponin homology domain from Utrophin (mCherry-CHUtr), which selectively labels F-actin, in clonal beta cells and imaged them by TIRF microscopy. Under basal conditions, cortical F-actin was seen as a meshwork with densely spaced focal contacts (Fig. 2A). Following KCl depolarization, there was a striking accumulation of cortical actin, often in distinct patches, evident as increased subplasma membrane F-actin reporter fluorescence (Fig. 2A–D). Addition of latrunculin A to depolymerize F-actin prevented this effect (Fig. 2A–D). A possible explanation for the cortical F-actin patches is local changes in Ca2+ concentration beneath the plasma membrane. To explore this possibility, we co-expressed the plasma membrane-immobilized Ca2+ indicator 5HT6-G-GECO and mCherry-CHUtr. Depolarization resulted in an immediate increase in subplasma membrane Ca2+ concentration, accompanied by F-actin accumulation (time difference 19±3 s, n=13), and, although we observed local changes in Ca2+ concentration, these did not coincide with the location of F-actin patch formation (Fig. 2E,F). Elevation of the glucose concentration from 3 mM to 11 mM resulted in periodic accumulations of cortical F-actin that, in most cases (82%), were preceded by increases in the cytosolic Ca2+ concentration. Addition of the KATP-channel opener diazoxide completely suppressed glucose-induced Ca2+ concentration changes and dramatically inhibited the F-actin rearrangements (Fig. 2G–I; Fig. S1). As for KCl, the glucose-induced cortical F-actin accumulations occurred nonhomogenously across the plasma membrane, but tended to reoccur in the same location during prolonged stimulations (Fig. 2G; Fig. S1). The magnitude of cortical actin accumulation positively correlated with the relative increase in cytosolic Ca2+ concentration, indicating a direct role of Ca2+ in actin redistribution (Fig. 2I). Addition of latrunculin A prevented the glucose-induced periodic accumulations of F-actin but did not affect the Ca2+ fluctuations (Fig. 2J). Interestingly, the latrunculin A-induced actin depolymerization, like microtubule disruption, prevented the light-induced recruitment of mitochondria to the plasma membrane, indicating a role of cortical actin in the transport process (Fig. 2K,L). The relationship between actin accumulation beneath the plasma membrane and the disappearance of mitochondria from the same volume indicates that actin biding to the plasma membrane might displace mitochondria. TIRF microscopy imaging of cortical mitochondria (GFP-Tomm20) and F-actin (mCherry-CHUtr) showed that the depolarization-induced increase in cortical actin density coincided with a reduction in mitochondria density in the same compartment (Fig. S1), similar to what has been shown for actin and cortical ER (Hsieh et al., 2017). To test whether the observed accumulation of cortical actin is directly caused by Ca2+ influx, we depolarized cells in the absence or presence of the L-type voltage-dependent Ca2+ channel blocker verapamil. In the presence of the inhibitor, depolarization failed to induce either Ca2+ influx or redistribution of cortical actin (Fig. 3), demonstrating direct effects of Ca2+ on the cortical actin network.

Fig. 2.

Cortical F-actin controls cortical mitochondria accumulation. (A) TIRF microscopy images of a MIN6 cell expressing mCh-CHUtr following depolarization in the absence or presence of the F-actin depolymerizing drug Latrunculin A (LatA; 1 μM). (B) Kymograph from a line drawn across the cell in A. (C) Relative change in fluorescence of mCh-CHUtr in the subplasma membrane region, as visualized by TIRF microscopy. (D) Quantifications of the responses in B and C (means±s.e.m., n=27 cells; ***P<0.001, paired Student's t-test). (E) TIRF microscopy recordings of subplasma membrane [Ca2+] (5HT6-G-GECO, black) and F-actin (mCherry-CHUtr, magenta) fluorescence during depolarization with 30 mM KCl (means±s.e.m. for 13 cells from four separate experiments). (F) TIRF microscopy image (left) and recordings (right) of the changes in mCherry-CHUtr (F-actin, magenta) and 5HT6-G-GECO (Ca2+, black) fluorescence in response to KCl depolarization. Traces from two subregions of the same cell are shown. ROI, region of interest. (G) TIRF microscopy images (top) and recording (middle) from a MIN6 cell expressing the Ca2+ reporter GCaMP5G (black) and the F-actin marker mCh-CHUtr (magenta). The cell was exposed to a rise in the surrounding glucose concentration from 3 mM to 11 mM, followed by addition of the KATP-channel opener diazoxide to silence electrical activity. Red triangles on the recording indicate the time points to which the images on top are related. A kymograph from a line drawn across the cell showing the formation of spatially restricted actin patches at the plasma membrane of a glucose-stimulated MIN6 cell is also shown (bottom; see Movie 1 for original data from which this was recorded). The red dashed lines indicate the cell periphery. (H) Changes in cytoplasmic Ca2+ concentration (GCaMP5G, black) and cortical actin density (mCh-CHUtr, magenta) during glucose-induced oscillations (means±s.e.m. for 94 oscillations from ten cells). Pie chart to the right shows the fraction of cells for which the increase in cortical actin is correlated with (dark gray) or not correlated with (light gray) a rise in cytosolic Ca2+ concentration. (I) Scatter plot showing the correlation between relative increase in cytosolic Ca2+ concentration and increase in cortical actin density in glucose-stimulated MIN6 cells (n=94 oscillations from 10 cells; P<0.05). (J) TIRF microscopy recording of GCaMP5G (black) and mCh-CHUtr (magenta) fluorescence from a MIN6 cell exposed to 20 mM glucose and 1 μM LatA (representative of 12 cells from two independent experiments). (K,L) TIRF microscopy images (J) and quantifications (K) of mApple-Tomm20 fluorescence in the subplasma membrane region of INS-1E cells co-expressing opto-mito. Data are presented as mitochondrial density at the plasma membrane (means±s.e.m., control=33.06±1.747%, light=46.76±1.769%, LatA=32.02±2.084%, LatA+light=36.11±2.328%; ncontrol=110, nlight=104, nLatA=68, nLatA+light=93, cells from three independent experiments; one-way ANOVA with Tukey's multiple comparison test, P<0.0001 for control versus light and LatA versus light, P=0.0007 for light versus LatA+light).

Fig. 2.

Cortical F-actin controls cortical mitochondria accumulation. (A) TIRF microscopy images of a MIN6 cell expressing mCh-CHUtr following depolarization in the absence or presence of the F-actin depolymerizing drug Latrunculin A (LatA; 1 μM). (B) Kymograph from a line drawn across the cell in A. (C) Relative change in fluorescence of mCh-CHUtr in the subplasma membrane region, as visualized by TIRF microscopy. (D) Quantifications of the responses in B and C (means±s.e.m., n=27 cells; ***P<0.001, paired Student's t-test). (E) TIRF microscopy recordings of subplasma membrane [Ca2+] (5HT6-G-GECO, black) and F-actin (mCherry-CHUtr, magenta) fluorescence during depolarization with 30 mM KCl (means±s.e.m. for 13 cells from four separate experiments). (F) TIRF microscopy image (left) and recordings (right) of the changes in mCherry-CHUtr (F-actin, magenta) and 5HT6-G-GECO (Ca2+, black) fluorescence in response to KCl depolarization. Traces from two subregions of the same cell are shown. ROI, region of interest. (G) TIRF microscopy images (top) and recording (middle) from a MIN6 cell expressing the Ca2+ reporter GCaMP5G (black) and the F-actin marker mCh-CHUtr (magenta). The cell was exposed to a rise in the surrounding glucose concentration from 3 mM to 11 mM, followed by addition of the KATP-channel opener diazoxide to silence electrical activity. Red triangles on the recording indicate the time points to which the images on top are related. A kymograph from a line drawn across the cell showing the formation of spatially restricted actin patches at the plasma membrane of a glucose-stimulated MIN6 cell is also shown (bottom; see Movie 1 for original data from which this was recorded). The red dashed lines indicate the cell periphery. (H) Changes in cytoplasmic Ca2+ concentration (GCaMP5G, black) and cortical actin density (mCh-CHUtr, magenta) during glucose-induced oscillations (means±s.e.m. for 94 oscillations from ten cells). Pie chart to the right shows the fraction of cells for which the increase in cortical actin is correlated with (dark gray) or not correlated with (light gray) a rise in cytosolic Ca2+ concentration. (I) Scatter plot showing the correlation between relative increase in cytosolic Ca2+ concentration and increase in cortical actin density in glucose-stimulated MIN6 cells (n=94 oscillations from 10 cells; P<0.05). (J) TIRF microscopy recording of GCaMP5G (black) and mCh-CHUtr (magenta) fluorescence from a MIN6 cell exposed to 20 mM glucose and 1 μM LatA (representative of 12 cells from two independent experiments). (K,L) TIRF microscopy images (J) and quantifications (K) of mApple-Tomm20 fluorescence in the subplasma membrane region of INS-1E cells co-expressing opto-mito. Data are presented as mitochondrial density at the plasma membrane (means±s.e.m., control=33.06±1.747%, light=46.76±1.769%, LatA=32.02±2.084%, LatA+light=36.11±2.328%; ncontrol=110, nlight=104, nLatA=68, nLatA+light=93, cells from three independent experiments; one-way ANOVA with Tukey's multiple comparison test, P<0.0001 for control versus light and LatA versus light, P=0.0007 for light versus LatA+light).

Fig. 3.

Cortical F-actin is modulated by Ca2+. (A) Kymographs from TIRF microscopy recordings of INS-1E cells expressing GCaMP5G and mCh-CHUtr following depolarization in the absence or presence of verapamil (100 µM). (B) TIRF microscopy recordings of the changes in GCaMP5G and mCh-CHUtr fluorescence in response to the indicated stimuli. A.U., arbitrary units. (C) Quantifications of the area under the curve (AUC) for the traces in B (unpaired Student's t-test: nGCAMP5G=5, PGCAMP5G=0.0241; nmCh-CHUtr=6, PmCh-CHUtr=0.02).

Fig. 3.

Cortical F-actin is modulated by Ca2+. (A) Kymographs from TIRF microscopy recordings of INS-1E cells expressing GCaMP5G and mCh-CHUtr following depolarization in the absence or presence of verapamil (100 µM). (B) TIRF microscopy recordings of the changes in GCaMP5G and mCh-CHUtr fluorescence in response to the indicated stimuli. A.U., arbitrary units. (C) Quantifications of the area under the curve (AUC) for the traces in B (unpaired Student's t-test: nGCAMP5G=5, PGCAMP5G=0.0241; nmCh-CHUtr=6, PmCh-CHUtr=0.02).

Ca2+ buffering by cortical mitochondria limits voltage-dependent Ca2+ influx

Mitochondria are important Ca2+ regulators in many cells, acting as low-affinity buffers under conditions of elevated cytosolic Ca2+ concentrations. Next, we therefore determined whether the subplasma membrane mitochondria in beta cells might shape the Ca2+ response following voltage-dependent Ca2+ influx. To this end, we depolarized cells with normal or light-induced increase in subplasma membrane mitochondrial density and compared the resulting Ca2+ elevation. We found that the Ca2+ response gradually decreased as mitochondria redistributed to the cell cortex, stabilizing at 50% reduction after 30 min of illumination (Fig. 4A,B). The typical Ca2+ response to depolarization was a mixture between stable elevation, reflecting voltage-dependent influx and superimposed transient peaks due to Ca2+ release from the ER (Xie et al., 2018). The latter phenomenon is secondary to stimulated insulin secretion and co-release of ATP that acts on purinergic P2Y1 receptors to generate Ca2+-mobilizing inositol 1,4,5-trisphosphate (IP3) (Wuttke et al., 2013). Increased cortical mitochondrial density suppressed both components (Fig. 4B–F), indicating that the mitochondria buffer Ca2+ from both sources. Similar suppression following light-induced recruitment of mitochondria to the cell cortex was also seen in beta cells stimulated with 20 mM glucose (Fig. 4G,H). To directly show that mitochondria sequester Ca2+ in response to depolarization, we performed simultaneous confocal microscopy recordings of cytosolic (GCaMP5G) and mitochondrial (mito-LAR-GECO) Ca2+ concentrations, and observed parallel changes in Ca2+ concentration in the two compartments (Fig. S2).

Fig. 4.

Cortical mitochondria buffer against changes in cytosolic [Ca2+]. (A) Top: TIRF microscopy recordings of changes in Ca2+ concentration in MIN6 cells expressing opto-mito (mApple-Tomm20-CRY2 and CIBN-GFP-Kif1a) or CRY2-mApple-Tomm20-CRY2 (control) following repeated brief depolarizations with 30 mM K+. Bottom: images showing the time-dependent recruitment of mitochondria towards the plasma membrane in cells expressing opto-mito but not in control cells (representative of 22 cells from three experiments). (B) Depolarization-induced Ca2+ influx in MIN6 cells expressing opto-mito in the absence of blue light (control) or following 30 min exposure to blue light (means±s.e.m., ncontrol=45, nlight=43). (C) Example recording of the change in R-GECO (Ca2+ indicator) fluorescence in a control cell exposed to 30 mM K+, showing a compound response consisting of an elevated basal level with superimposed spikes. The example is used to schematically describe the parameters analyzed. (D–F) Quantifications of depolarization-induced changes in R-GECO fluorescence in cells expressing opto-mito in the absence (control) or presence (light) of blue-light illumination: plateau height (D), mean peak height (E), peak frequency (F) (base level: ncontrol=48, nlight=46; P=0.037; mean peak height: ncontrol=38, nlight=33; P=0.0225; number of peaks: ncontrol=48, nlight=46; P=0.3159; unpaired Student's t-test, data from six individual experiments). (G) TIRF microscopy recording of R-GECO fluorescence from a glucose-stimulated MIN6 cell co-expressing opto-mito following blue-light illumination. (H) Normalized changes in R-GECO fluorescence in glucose-stimulated MIN6 cells co-expressing CRY2-Tomm20 (control) or CRY2-Tomm20 and CIBN-Kif1a (opto-mito) before and after 20 min of blue-light exposure (ncontrol=11 cells, nopto-mito=10 cells, two independent experiments; two-tailed paired Student's t-test was used for comparisons within groups and two-tailed unpaired Student's t-test was used for comparison between groups).

Fig. 4.

Cortical mitochondria buffer against changes in cytosolic [Ca2+]. (A) Top: TIRF microscopy recordings of changes in Ca2+ concentration in MIN6 cells expressing opto-mito (mApple-Tomm20-CRY2 and CIBN-GFP-Kif1a) or CRY2-mApple-Tomm20-CRY2 (control) following repeated brief depolarizations with 30 mM K+. Bottom: images showing the time-dependent recruitment of mitochondria towards the plasma membrane in cells expressing opto-mito but not in control cells (representative of 22 cells from three experiments). (B) Depolarization-induced Ca2+ influx in MIN6 cells expressing opto-mito in the absence of blue light (control) or following 30 min exposure to blue light (means±s.e.m., ncontrol=45, nlight=43). (C) Example recording of the change in R-GECO (Ca2+ indicator) fluorescence in a control cell exposed to 30 mM K+, showing a compound response consisting of an elevated basal level with superimposed spikes. The example is used to schematically describe the parameters analyzed. (D–F) Quantifications of depolarization-induced changes in R-GECO fluorescence in cells expressing opto-mito in the absence (control) or presence (light) of blue-light illumination: plateau height (D), mean peak height (E), peak frequency (F) (base level: ncontrol=48, nlight=46; P=0.037; mean peak height: ncontrol=38, nlight=33; P=0.0225; number of peaks: ncontrol=48, nlight=46; P=0.3159; unpaired Student's t-test, data from six individual experiments). (G) TIRF microscopy recording of R-GECO fluorescence from a glucose-stimulated MIN6 cell co-expressing opto-mito following blue-light illumination. (H) Normalized changes in R-GECO fluorescence in glucose-stimulated MIN6 cells co-expressing CRY2-Tomm20 (control) or CRY2-Tomm20 and CIBN-Kif1a (opto-mito) before and after 20 min of blue-light exposure (ncontrol=11 cells, nopto-mito=10 cells, two independent experiments; two-tailed paired Student's t-test was used for comparisons within groups and two-tailed unpaired Student's t-test was used for comparison between groups).

Subplasma membrane mitochondria participate in the regulation of insulin secretion

Exocytosis of insulin-containing granules is triggered by Ca2+ influx. Since we found that Ca2+ has an influence on mitochondrial position, and that the mitochondrial position, in turn, has an impact on the cytosolic Ca2+ concentration, we next investigated the role of subplasma membrane mitochondria in the regulation of exocytosis. We used vesicle-associated membrane protein 2 (VAMP2) fused to the pH-sensitive fluorescent protein pHluorin as a detector of exocytosis. In cells co-expressing the mitochondrial marker mApple-Tomm20, we observed mitochondria in the proximity of, but not overlapping with, insulin granule fusion sites at the plasma membrane (Fig. 5A,B). To directly test the role of cortical mitochondria in the regulation of insulin secretion, we used the light-driven molecular motors to increase the cortical mitochondria density. To measure insulin secretion in real time at the single-cell level, we utilized a well-characterized feedback mechanism. The above-mentioned autocrine effect of ATP co-secreted with insulin in addition to IP3 also generates diacylglycerol (DAG) in the plasma membrane (Fig. 5C) (Wuttke et al., 2013). DAG recordings can therefore be used as a proxy for insulin secretion. Depolarization of MIN6 cells expressing the DAG biosensor mCherry-C1aC1bPKC resulted in pronounced DAG spikes, reflecting individual granule fusion events, and application of the P2Y1 receptor antagonist MRS2179 completely inhibited this response (Fig. 5D,E). In cells with light-induced increase in cortical mitochondria density, there was a 50% reduction in DAG spike frequency, indicating that cortical mitochondria can restrict insulin secretion (Fig. 5F). Consistent with this notion, we found that glucose-induced reductions in cortical mitochondria density were associated with increased DAG spike frequency (Fig. S2).

Fig. 5.

Cortical mitochondria limit insulin secretion. (A) TIRF microscopy images of a MIN6 cell expressing mApple-Tomm20 (mitochondria) and VAMP2-pHluorin (exocytosis sites). Images from different time points during depolarization with 30 mM K+ are shown. Arrows indicate insulin granule fusion events occurring in the proximity of cortical mitochondria. (B) Magnification of a single fusion site showing lack of colocalization between mitochondria and insulin granules. Graphs show the average VAMP2-pHluorin and mApple-Tomm20 fluorescence at sites of fusion (top) or at random sites (bottom) (nmito=12, nVamp2=12). (C) Principle of the indirect measurement of insulin secretion using ATP-dependent feedback activation of phospholipase C and diacylglycerol (DAG) formation. (D) TIRF microscopy recordings of the depolarization-induced DAG formation (mCh-C1aC1b) in MIN6 cells that can be inhibited by the P2Y1 receptor antagonist MRS2179 (10 μM). (E) Quantifications of the DAG spike frequency (n=29 cells; P<0.001, unpaired Student's t-test). (F) TIRF microscopy recordings of depolarization-induced DAG spikes in MIN6 cells co-expressing CRY2-Tomm20 (control) or CRY2-Tomm20 and CIBN-GFP-Kif1a (opto-mito) (ncontrol=26, nopto=14, cells from five individual experiments; P=0.0104, unpaired Student's t-test).

Fig. 5.

Cortical mitochondria limit insulin secretion. (A) TIRF microscopy images of a MIN6 cell expressing mApple-Tomm20 (mitochondria) and VAMP2-pHluorin (exocytosis sites). Images from different time points during depolarization with 30 mM K+ are shown. Arrows indicate insulin granule fusion events occurring in the proximity of cortical mitochondria. (B) Magnification of a single fusion site showing lack of colocalization between mitochondria and insulin granules. Graphs show the average VAMP2-pHluorin and mApple-Tomm20 fluorescence at sites of fusion (top) or at random sites (bottom) (nmito=12, nVamp2=12). (C) Principle of the indirect measurement of insulin secretion using ATP-dependent feedback activation of phospholipase C and diacylglycerol (DAG) formation. (D) TIRF microscopy recordings of the depolarization-induced DAG formation (mCh-C1aC1b) in MIN6 cells that can be inhibited by the P2Y1 receptor antagonist MRS2179 (10 μM). (E) Quantifications of the DAG spike frequency (n=29 cells; P<0.001, unpaired Student's t-test). (F) TIRF microscopy recordings of depolarization-induced DAG spikes in MIN6 cells co-expressing CRY2-Tomm20 (control) or CRY2-Tomm20 and CIBN-GFP-Kif1a (opto-mito) (ncontrol=26, nopto=14, cells from five individual experiments; P=0.0104, unpaired Student's t-test).

In this study, we set out to determine the importance of mitochondrial positioning for the regulation of insulin secretion from beta cells. We identified a pool of subplasma membrane-localized mitochondria that undergoes dynamic changes in localization in a Ca2+- and F-actin-dependent manner and contributes to the regulation of insulin secretion by buffering cytosolic Ca2+.

The importance of mitochondrial metabolism for normal glucose-stimulated insulin secretion is well established; ATP derived from oxidative phosphorylation is required for inducing the membrane depolarization that triggers exocytosis of insulin granules. ATP is also crucial for priming and replenishing the release-competent pool of insulin granules (Rorsman and Ashcroft, 2018). In addition, the mitochondria act as low-affinity Ca2+ buffers that sequester the ion under conditions of elevated cytosolic Ca2+, such as those triggering insulin secretion (Tarasov et al., 2012; Wiederkehr and Wollheim, 2012). To what extent the ensuing increase in mitochondrial Ca2+ impacts mitochondrial metabolism and subsequent insulin secretion is still a matter of debate, with studies supporting both stimulatory and inhibitory roles of the ion on metabolism (Kennedy et al., 1999; Krippeit-Drews et al., 2000; Li et al., 2013; Wiederkehr et al., 2011). Another layer of complexity in mitochondrial regulation of beta-cell function is the control of their position within the cells. The mitochondria are highly dynamic organelles that move along microtubules, and it has been shown, in neurons, that the presence of mitochondria in the synapses is required for normal neurotransmission (Vaccaro et al., 2017). The beta cells lack the complex morphology of neurons but still have regions in the plasma membrane that resemble the synaptic active zones (Low et al., 2014). Cortical mitochondria have been described in many cell types, and often consist of a mobile and more stable population. We find that beta cells under resting conditions have a high density of mitochondria beneath the plasma membrane, similar to what has been described in the related chromaffin cell (Villanueva et al., 2014). Glucose stimulation did not result in a stable change in mitochondrial density at the plasma membrane, but the glucose-induced Ca2+ oscillations were frequently associated with transient displacements of cortical mitochondria. Consistent with this, depolarization with ensuing Ca2+ influx resulted in a reduction in the number of cortical mitochondria. We find that this Ca2+-induced redistribution of mitochondria is likely due to an increase in the cortical F-actin density. Increased association between cortical F-actin and the plasma membrane under conditions promoting exocytosis has previously been shown in chromaffin cells, and found to be important for the replenishment of plasma membrane-docked granules (Papadopulos et al., 2015; Wen et al., 2011). The redistribution involves activation of the GTPase Cdc42 and the Arp2/3 complex (Gasman et al., 2004). Interestingly, Cdc42 activity was recently shown to be stimulated by Ca2+/calcineurin, providing a possible mechanism for the depolarization-induced actin remodeling (Ly and Cyert, 2017). F-actin has also been found to regulate insulin secretion through multiple pathways, including a direct interaction with syntaxin 4, a protein that is important for granule docking (Jewell et al., 2008; Tomas et al., 2006). In the case of secretory granules, the redistribution of F-actin results in increased granule density at the plasma membrane (Papadopulos et al., 2015), whereas we observe reduced mitochondrial density under the same conditions. The ER also shows, in many cell types, a prominent cortical distribution that is similarly restricted by the presence of F-actin (Hsieh et al., 2017). Acute increase in cortical actin density, as observed here, might therefore simply displace mitochondria that are not stably attached to the plasma membrane. Some mitochondria remain at the plasma membrane, and these might reflect a recently described population that is stably tethered (Lackner et al., 2013). Ca2+ increases have previously been shown to reduce the motility of mitochondria along microtubules (Yi et al., 2004), and it has been suggested that the stalling at sites of high Ca2+ would enable local ATP production. This mechanism could be of importance in asymmetrical cells in which free diffusion of ATP is restricted, but it is less clear as to what extent ATP microdomains exist in small symmetrical cells, such as beta cells (Kennedy et al., 1999).

To determine the role of cortical mitochondria in the regulation of beta-cell function, we developed a light-regulated transport system, similar to what has recently been described (van Bergeijk et al., 2015), and, using this, we were able to increase the cortical mitochondria density twofold. Since the light-induced mitochondria translocation is based on kinesin-dependent transport along microtubules, it was completely blocked by the microtubule destabilizing drug nocodazole. The reaction was reversed within 2 h, when illumination was interrupted. The light-induced changes in cortical mitochondria operate on a significantly longer timescale than the rapid dynamics induced by Ca2+, likely reflecting both the slow reversibility of the optogenetic dimers (Kennedy et al., 2010) and the requirement of active, retrograde transport. Surprisingly, the light-induced transport also required intact F-actin, as it was completely inhibited by the actin depolymerizing drug latrunculin A. It has been shown that, although transport along microtubules is the main mechanism, short-distance mitochondrial movements and immobilization of the organelle at the cell cortex often require the actin cytoskeleton (Boldogh and Pon, 2006; Chada and Hollenbeck, 2004; Saxton and Hollenbeck, 2012). Furthermore, mitochondrial movement is enhanced in the absence of F-actin (Morris and Hollenbeck, 1995). The light-induced accumulation of cortical mitochondria resulted in pronounced suppression of both glucose- and depolarization-induced elevation of the cytosolic Ca2+ concentration and of insulin granule exocytosis. Mitochondria have previously been ascribed local buffering capacity at ER–mitochondria contact sites, where they limit the spread of Ca2+ released by IP3 receptors (Rizzuto et al., 1998). The findings here suggest a similar role of mitochondria in controlling a pool of Ca2+ of importance for regulated secretion, and that this buffering capacity is under the control of Ca2+ itself. We propose a model in which cortical mitochondria accumulate close to sites of exocytosis, perhaps to fuel energy-demanding processes related to granule priming and release. Upon influx of Ca2+, the cortical actin density is increased, causing rapid displacement of mitochondria with resulting loss of Ca2+ buffering capacity near the influx sites, thereby supporting continued exocytosis (Fig. 6).

Fig. 6.

Proposed model for the involvement of actin filaments and mitochondria in the regulation of beta-cell Ca2+ homeostasis. Mitochondria are transported by microtubules, but the final transport towards the plasma membrane depends on F-actin. Mitochondria occupy a certain volume beneath the plasma membrane under resting conditions. Voltage-dependent Ca2+ influx results in an increase in cortical F-actin density that displaces mitochondria from the same volume. Sites of exocytosis are devoid of mitochondria, which instead seem to accumulate in proximity to the fusion site. An increase in cortical mitochondria density buffers against changes in Ca2+ concentration and thereby limits insulin secretion.

Fig. 6.

Proposed model for the involvement of actin filaments and mitochondria in the regulation of beta-cell Ca2+ homeostasis. Mitochondria are transported by microtubules, but the final transport towards the plasma membrane depends on F-actin. Mitochondria occupy a certain volume beneath the plasma membrane under resting conditions. Voltage-dependent Ca2+ influx results in an increase in cortical F-actin density that displaces mitochondria from the same volume. Sites of exocytosis are devoid of mitochondria, which instead seem to accumulate in proximity to the fusion site. An increase in cortical mitochondria density buffers against changes in Ca2+ concentration and thereby limits insulin secretion.

Materials

Nocodazole, latrunculin A, verapamil and diazoxide were from Sigma-Aldrich (Germany). The following plasmids were used in this study: GFP-CRY2-OCRL (Idevall-Hagren et al., 2012), CIBN-GFP-CAAX (Idevall-Hagren et al., 2012), mApple-Tomm20-N1 (Addgene plasmid #54955), R-GECO [Addgene plasmid #32444 (Zhao et al., 2011)], GCaMP5G [Addgene plasmid #31788 (Akerboom et al., 2012)], mCherry-CHUtr [Addgene plasmid #26740 (Burkel et al., 2007)], mCherry-C1aC1bPKC (Wuttke et al., 2013), iRFP-PH-PLCδ1 (Idevall-Hagren et al., 2012), mito-LAR-GECO [Addgene plasmid #61245 (Wu et al., 2014)] and 5HT6-G-GECO [Addgene plasmid #47499 (Su et al., 2013)].

Blue light-controlled molecular motors

Kif1a-GFP-CIBN was generated by cloning mouse Kif1a into GFP-CIBN using the following primers: Kif1a-NheI-for (5′-TATAGCTAGCGCCACCATGGCTGGGGCCTCTGT-3′), Kif1a-AgeI-rev (5′-TATAACCGGTCCCAGCAGATCTCGCAGCC-3′). Kif1a-iRFP-CIBN was generated by replacing GFP in Kif1a-GFP-CIBN with iRFP using the following primers: AgeI-iRFP-for (5′-CTCACCGGTAGCTGAAGGATCCGTCGCCAG-3′), HindIII-iRFP-rev (5′-CTCAAGCTTGCTCTTCCATCACGCCGATCTG-3′). Tomm20-GFP-C1-CRY2 was generated by amplification of the N-terminal region of mouse Tomm20 using the following primers: Tomm20-Nhe1-for (5′-TATAGCTAGCACCATGGTGGGTCGGA-3′), Tomm20-Nhe1-rev (5′-TATAGCTAGCAGACCACCAGATTGAAAAT-3′). Subsequently, CRY2 was inserted into the multiple cloning site after the GFP using the restriction sites Kpn1 and Mfe1 (CRY2-Kpnl-for 5′-TATAGGTACCATGAAGATGGACAAAA-3′, CRY2-MfeI-rev 5′-TATACAATTGATCTGCTGCTCCGATCATGA-3′). Tomm20-mApple-CRY2 was generated by inserting CRY2 into the Tomm20-mApple plasmid (Addgene plasmid #54955) via the restriction sites Not1 and Xba1 (NotI-Cry2-for 5′-AACGCGGCCGCCATGAAGATGGACAAAAAGACT-3′, XbaI-Cry2-rev 5′-AACTCTAGATGCTGCTCCGATCATGATCT-3′). A stop codon C-terminal to mApple was deleted using a Q5 mutagenesis kit (New England Biolabs), Q5SDM_mCh-Tomm20-for 5′-GCGGCCGCCATGAAGATG-3′ and Q5SDM_mCh-Tomm20-rev 5′-CTTGTACAGCTCGTCCATGC-3′. All plasmids were verified by sequencing.

Cell culture

Two different beta-cell lines were used in this study: the mouse cell line MIN6 (Ishihara et al., 1993) and the rat cell line INS-1E (Merglen et al., 2004). Cell lines were tested for mycoplasma at regular intervals. MIN6 cells were cultured in Dulbecco's modified Eagle medium (4.5 g/l D-glucose), supplemented with 15% fetal calf serum (FCS) and 50 µM β-mercaptoethanol. The INS-1E cells were kept in RPMI-1640 medium supplemented with 50 µM β-mercaptoethanol, 1 mM sodium pyruvate, 10 mM HEPES and 10% FCS (all Life Technologies, The Netherlands). All cells were cultured at 37°C and 5% CO2.

Transfection

Cells were plated on 25-mm glass coverslips and grown for 24–48 h until reaching ∼60% confluence. Expression vectors were subsequently transfected into cells using Lipofectamine 2000 (Life Technologies) according to the manufacturer's instructions. Cells were used for experimentation 24–48 h after interruption of the transfection reaction.

Optogenetics

Blue light-induced recruitment of mitochondria to the subplasma membrane region of cells was accomplished by epifluorescence illumination at the stage of a TIRF microscope using a custom-built illuminator. Briefly, 470-nm light provided by a light-emitting diode (LED) (200 mW, ThorLabs, Newton, NJ) was collimated and focused to a 10-mm (∅) spot that was projected to the cells through the bottom of a quartz prism used to generate evanescent wave excitation (Idevall-Hagren et al., 2010). Illumination (10% output power) was provided as 100-ms pulses every 10 s. In some experiments, cells were instead exposed to blue-light illumination for 10–60 min inside a cell culture incubator using a collimated 470-nm blue-light LED as previously described (Xie et al., 2016). Following illumination, cells were placed at the stage of a TIRF microscope and continuously illuminated with 491-nm blue light derived through the evanescent field.

TIRF microscopy

Selective visualization of the plasma membrane/subplasma membrane region was performed by TIRF microscopy, using either an inverted microscope and a 60×/1.45 NA objective or a prism-based system built around an upright microscope equipped with a 16×/0.8 NA objective, as previously described (Idevall-Hagren et al., 2010; Xie et al., 2016). Before all imaging experiments, cells were pre-incubated for 30 min at 37°C in a buffer containing 125 mM NaCl, 4.9 mM KCl, 1.3 mM CaCl2, 1.2 mM MgCl2 (all Merck, Germany), 10 mM HEPES, 0.1 mg/ml bovine serum albumin (Sigma-Aldrich) and 3 mM glucose (pH 7.40) (Sigma-Aldrich). The cells were subsequently mounted in modified Sykes–Moore chambers, placed at the stage of the microscope and superfused with buffer. Temperature was maintained at 37°C throughout all experiments.

Analysis

Image analysis was performed using Fiji software (Schindelin et al., 2012). For quantifications of subplasma membrane changes in fluorescence, regions of interest were identified on the TIRF micrographs based on the cell footprint as visualized by a general plasma membrane stain (iRFP-PH-PLCδ1). The changes in fluorescence of other markers within this region over the time course of an experiment were subsequently determined, and the data were normalized to the pre-stimulatory level using Excel (Microsoft Corp., Redmond, WA). All experiments were repeated a minimum of three times, unless stated otherwise, and statistical analysis – using one-way ANOVA for multiple comparisons or two-tailed paired or unpaired Student's t-tests – was performed using Prism (version 7.00 for Mac, GraphPad Software, La Jolla, CA). P<0.05 was considered statistically significant.

We thank Antje Thonig for excellent technical assistance and Erik Gylfe for critical reading of the manuscript.

Author contributions

Conceptualization: O.I.-H.; Methodology: N.G.; Software: C.H.; Formal analysis: N.G., G.S., O.I.-H.; Investigation: N.G., O.I.-H.; Resources: O.I.-H.; Data curation: N.G., G.S., O.I.-H.; Writing - original draft: N.G.; Writing - review & editing: N.G., G.S., O.I.-H.; Visualization: N.G.; Supervision: O.I.-H.; Project administration: O.I.-H.; Funding acquisition: O.I.-H.

Funding

This study was supported by grants from Göran Gustafssons Stiftelse för Naturvetenskaplig och Medicinsk Forskning, the Malin and Lennart Philipson Foundation, Vetenskapsrådet [MH2015-03087], Åke Wiberg Stiftelse [M17-0048], Novo Nordisk Fonden [NNF15OC0016100] and EXODIAB (to O.I.-H).

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Competing interests

The authors declare no competing or financial interests.

Supplementary information