Mitochondrial clearance of calcium facilitated by MICU2 controls insulin secretion

Abstract

Objective

Transport of Ca²⁺ into pancreatic β cell mitochondria facilitates nutrient-mediated insulin secretion. However, the underlying mechanism is unclear. Recent establishment of the molecular identity of the mitochondrial Ca²⁺ uniporter (MCU) and associated proteins allows modification of mitochondrial Ca²⁺ transport in intact cells. We examined the consequences of deficiency of the accessory protein MICU2 in rat and human insulin-secreting cells and mouse islets.

Methods

siRNA silencing of Micu2 in the INS-1 832/13 and EndoC-βH1 cell lines was performed; Micu2^−/− mice were also studied. Insulin secretion and mechanistic analyses utilizing live confocal imaging to assess mitochondrial function and intracellular Ca²⁺ dynamics were performed.

Results

Silencing of Micu2 abrogated GSIS in the INS-1 832/13 and EndoC-βH1 cells. The Micu2^−/− mice also displayed attenuated GSIS. Mitochondrial Ca²⁺ uptake declined in MICU2-deficient INS-1 832/13 and EndoC-βH1 cells in response to high glucose and high K⁺. MICU2 silencing in INS-1 832/13 cells, presumably through its effects on mitochondrial Ca²⁺ uptake, perturbed mitochondrial function illustrated by absent mitochondrial membrane hyperpolarization and lowering of the ATP/ADP ratio in response to elevated glucose. Despite the loss of mitochondrial Ca²⁺ uptake, cytosolic Ca²⁺ was lower in siMICU2-treated INS-1 832/13 cells in response to high K⁺. It was hypothesized that Ca²⁺ accumulated in the submembrane compartment in MICU2-deficient cells, resulting in desensitization of voltage-dependent Ca²⁺ channels, lowering total cytosolic Ca²⁺. Upon high K⁺ stimulation, MICU2-silenced cells showed higher and prolonged increases in submembrane Ca²⁺ levels.

Conclusions

MICU2 plays a critical role in β cell mitochondrial Ca²⁺ uptake. β cell mitochondria sequestered Ca²⁺ from the submembrane compartment, preventing desensitization of voltage-dependent Ca²⁺ channels and facilitating GSIS.

Graphical abstract

Highlights

• •MICU2 deficiency impairs glucose-stimulated insulin secretion.
• •MICU2 deficiency abrogates mitochondrial calcium uptake.
• •Depolarization-evoked cytosolic calcium increases are lower in MICU2-deficient cells.
• •Mitochondria clear calcium from the subplasma membrane region, maintaining calcium influx.

1. Introduction

While the canonical model of glucose-stimulated insulin secretion (GSIS) from pancreatic β cells with a central role of the mitochondrion has been established for more than 30 years, many details remain unclear. Extensive literature links mitochondrial Ca²⁺ uptake to facilitation of insulin secretion [[1], [2], [3], [4], [5], [6]]. However, it is unclear whether the reported effects of elevated matrix-free Ca²⁺ on insulin secretion are due to the bioenergetic consequences of the activation of citric acid cycle enzymes [7], the facilitation of plasma membrane oscillations [8], the generation of so-called coupling factors [5,9], or additional currently unrecognized mechanisms. The molecular identification of the components of the mitochondrial Ca²⁺ uniporter with the ability to selectively ablate one or more components allows the consequences of disrupted mitochondrial Ca²⁺ transport to be investigated in β cells.

The uniporter complex comprises the pore-forming subunit MCU plus up to six regulatory subunits [10]. MCU forms the complex's central pore-forming subunit, while MCU regulatory subunit b (MCUb), essential MCU regulator (EMRE), and mitochondrial uptake 1 and 2 (MICU1 and MICU2) form the accessory components [11]. As part of the Ca²⁺ uniporter holocomplex, MICU1 and MICU2 heterodimerize and act to maintain a threshold for mitochondrial Ca²⁺ uptake, upholding the mitochondrial Ca²⁺-buffering capacity [[12], [13], [14]]. It has been demonstrated in INS-1 832/13 insulinoma cells that MCU and MICU1 play a role in metabolic coupling; reduced expression of either decreases mitochondrial Ca²⁺ uptake and inhibits insulin secretion [8]. Furthermore, β cell-specific Mcu knockout mice display reduced first-phase insulin release in vivo and impaired mitochondrial Ca²⁺ uptake, glucose-induced ATP production, and insulin secretion in vitro [15]. However, the function of MICU2 in stimulus-secretion coupling in pancreatic β cells has not been examined. The situation in other cell types is somewhat unclear. In HeLa cells, silencing of Micu2 leads to increased mitochondrial Ca²⁺ uptake [16,17], but only at cytosolic levels lower than 5 μM [17]. A similar phenotype is also observed in HEK-293T cells [13,18], while silencing of Micu2 in mouse liver leads to impaired Ca²⁺ uptake kinetics [19]. To resolve the role of Ca²⁺ uptake, we examined the impact of MICU2 deficiency on insulin-secreting cells and mouse islets as well as in HEK-293T cells.

2. Materials and methods

2.1. Cell culture and model systems

INS-1 832/13 cells were cultured as previously described [20]. EndoC-βH1 cells [21] were grown on Matrigel-fibronectin-coated (100 μg/mL and 2 μg/mL, respectively, Sigma–Aldrich) culture vessels in DMEM containing 5.6 mM of glucose, 2% BSA fraction V (Roche Diagnostics), 10 mM of nicotinamide (Calbiochem), 50 μM of 2-mercaptoethanol, 5.5 μg/mL of transferrin, 6.7 ng/mL of sodium selenite (Sigma–Aldrich), 100 U/mL of penicillin, and 100 μg/mL of streptomycin (PAA). Both cell lines were cultured at 37 °C with 5% CO₂. Unless otherwise stated, EndoC-βH1 cells were seeded at 2.3 × 10⁵ cells/cm² and INS-1 832/13 cells at 1.5 × 10⁵ cells/cm² in 24-well plates (Matrigel-fibronectin coated or uncoated) and transfected either with siRNA specific to MICU2 (50 nM) or a scrambled siRNA (50 nM). Forty-eight h after knockdown, the growth media were changed to an overnight starvation medium containing 5.6 mM of glucose for INS-1 832/13 cells and 1 mM of glucose for EndoC-βH1 cells. HEK-293T cells were cultured in DMEM containing 1 g/L of d-glucose and supplemented with 10% heat-inactivated FBS, 100 U/mL of penicillin, and 100 μg/mL of streptomycin (PAA) [22]. The cells were plated at 1.5 × 10⁵ cells/cm² in 24-well plates and transfected with either siRNA specific to MICU2 or a scrambled siRNA. All of the assays were performed 48–72 h after knockdown.

2.2. Micu2 knockout mice

We purchased a Micu2 gene trap from the Texas A&M Institute of Genomic Medicine (College Station, TX, USA), bred the mice, and validated the disruption of the Micu2 gene using standard procedures. The Micu2 knockout mice had a B57BL/6J background and were extensively backcrossed. This mouse was previously characterized [23]. A combination of male and female mice was used with no significant differences between the two sexes.

2.3. Insulin secretion assay

INS-1 832/13 cells were incubated in HEPES-balanced salt solution (HBSS; 115 mM of NaCl, 5 mM of KCl, 1 mM of MgSO₄, 1 mM of CaCl₂, 25.5 mM of NaHCO₃, 20 mM of HEPES, and 0.2% BSA at a pH of 7.2) containing 2.8 mM of glucose for 2 h followed by 1 h of stimulation with HBSS containing 2.8 or 16.7 mM of glucose or 15 min of stimulation with HBSS containing 36 mM of KCl and 2.8 mM of glucose [20]. Insulin secretion was measured with a rat insulin ELISA kit (Mercodia A/B, Sweden) according to the manufacturer's instructions. EndoC-βH1 cells were starved overnight in 1 mM of glucose-containing growth medium 1 day before the experiment. On the day of the experiment, the growth medium was removed and replaced with 1 mM of glucose-containing HBSS for 2 h followed by 1 h of stimulation with HBSS containing 1 or 20 mM of glucose [24]. Insulin secretion was measured with a human insulin ELISA (Mercodia A/B, Sweden) according to the manufacturer's instructions.

2.4. RNA isolation and quantitative real-time PCR

Total RNA was extracted from EndoC-βH1 cells, INS-1 832/13, and HEK-293T cells using an RNeasy Mini kit (Qiagen) according to the manufacturer's protocol. RNA concentrations were determined using a NanoDrop Spectrophotometer (Thermo Fisher Scientific). Then 1000 ng of total RNA were reverse transcribed using a RevertAid First-Strand cDNA synthesis kit (Fermentas, Vilnius, Lithuania). Quantitative real-time PCR (qPCR) was performed using TaqMan gene expression assays (EFHA1/Micu2 rat: s235193; EFHA1/Micu2 human: s47976; Hprt1 rat: Rn01527840; Hprt1 human: Hs99999909; Mcu rat: Rn01433739; Micu1 rat: Rn01644475; Cacna1a rat, Rn00563825; Cacna1c rat, Rn00709287; and INS1 mouse, Mm01950294, Applied Biosystems, Life Technologies, Carlsbad, CA, USA) using a 7900HT Fast Real-Time System (Applied Biosystems). qPCR was carried out as follows. Five ng of cDNA were mixed with 0.5 μL of gene assay and 5 μL of TaqMan Master Mix and ddH₂O to a final volume of 10 μL and qPCR was performed using the following program: 50 °C for 2 min, 95 °C for 10 min, 40 cycles of 95 °C for 15 s, and 60 °C for 1 min. In all of the runs, a 10x dilution series was produced of one of the samples and used to generate a calibration curve as well as quality control for amplification efficiency. Gene expression was relatively quantified using the calibration curve. The amount of mRNA was calculated relative to the amount of hypoxanthine-guanine phosphoribosyl transferase (HPRT1) as a reference gene.

2.5. SDS-PAGE and western blotting

Cells were harvested in RIPA buffer followed by lysis by shaking the samples for 30 min at 950 rpm and 4 °C for 30 min. This was followed by centrifugation at 16,000g and 4 °C for 20 min to remove cell debris. Supernatants/lysates were kept at −80 °C until use. Then 10–20 μg of protein were separated on mini-PROTEAN TGX pre-stained gels (Bio-Rad) followed by activation of pre-stained Bio-Rad MP imager. The proteins were blotted onto Trans-Blot Turbo Transfer Packs (Bio-Rad), using Bio-Rad semidry blotter, and total protein for loading control was determined from the pre-stains. Membranes were blocked with 10% BSA or 5% skim milk (for mitochondrial complexes I–V experiments) in TRIS-buffered saline (TBS) unless otherwise stated and hybridized with antibodies for MICU2/EFHA1 (Abcam, ab101465), β-tubulin (Abcam, ab6046), α-tubulin (Abcam, ab7291), and mitochondrial complexes I–V (Abcam, ab110413).

2.6. Single-cell live cytosolic ATP and ATP/ADP ratio measurements using perceval and perceval HR

Single-cell cytosolic ATP/ADP ratio measurements were carried out in INS-1 832/13 cells using the genetically encoded biosensor Perceval HR [25]. The cells were seeded on poly-d-lysine-coated 8-well-chambered cover glasses (Lab-Tek, Thermo Fisher Scientific) at a density of 70,000 cells/cm². Twenty-four h after seeding, the cells were co-transfected with siRNA and 1 μg of plasmid-encoding Perceval HR (Addgene ID: 49,083) at approximately 50% cell confluency. The cells were further grown for 48–72 h before measurements. On the day of imaging, the cells were pre-incubated at 37 °C in 400 μL of buffer P (135 mM of NaCl, 3.6 mM of KCl, 1.5 mM of CaCl₂, 0.5 mM of MgSO₄, 0.5 mM of Na₂HPO₄, 10 mM of HEPES, and 5 mM of NaHCO₃ at a pH of 7.4) containing 2.8 mM glucose. After 1.5 h of incubation, the cells were imaged at 488 nm excitation and 505–535 nm emission recorded on a Zeiss LSM510 inverted confocal fluorescence microscope. Mouse islets were transduced with adenovirus expressing the similar ATP biosensor Perceval [26,27]. Poly-d-lysine-coated 8-well-chambered cover glasses (Lab-Tek, Thermo Fisher Scientific) containing mouse islets were incubated for 2 h with RPMI with 10% FBS, 100 U of penicillin, and 100 mg/mL of streptomycin containing the Perceval adenovirus. Then this medium was replaced with fresh RPMI medium with 10% FBS, 100 U of penicillin, and 100 mg/mL of streptomycin and incubated overnight. On the day of imaging, the cells were pre-incubated at 37 °C in 400 μL of buffer P (135 mM of NaCl, 3.6 mM of KCl, 1.5 mM of CaCl₂, 0.5 mM of MgSO₄, 0.5 mM of Na₂HPO₄, 10 mM of HEPES, and 5 mM of NaHCO₃ at a pH of 7.4) containing 2.8 mM of glucose. After 1.5 h of incubation, the cells were imaged as described for clonal cell lines at 488 nm excitation and emission recorded at 505–530 nm on a Zeiss LSM510 inverted confocal fluorescence microscope using a 40x/0.75 objective. Images were recorded at a frequency of 0.25 Hz (scan time = 3.93 s). Equimolarity was accounted for in the 36 mM KCl solution.

2.7. Single-cell live cytoplasmic-free Ca²⁺ measurements

Cells were seeded onto poly-d-lysine coated 8-well-chambered cover glasses (Lab-Tek, Naperville, IL, USA) at a density of 70,000 cells/cm² per well. Twenty-four h after seeding, the cells were transfected with siRNA and incubated for further 48–72 h prior to imaging. On the day of imaging, the cells were pre-incubated at 37 °C in 400 μL of buffer P containing 2.8 mM of glucose. After 1.5 h of incubation, based on variable Ca²⁺ affinities, INS-1 832/13 and HEK-293T cells were loaded with 2 μM of Fluo4 AM and Fluo5F AM, respectively. Incubation was continued for an additional 30 min. Immediately prior to imaging, the pre-incubation buffer was exchanged with 400 μL of buffer P. Fluo4 AM and Fluo5F AM were excited at 488 nm and emission recorded using a 505 nm emission filter on a Zeiss LSM510 inverted confocal fluorescence microscope using a 40x/0.75 objective. Images were recorded at a frequency of 0.64 Hz (scan time = 1.57 s) using a pinhole diameter of 463 μm. Single-cell-free cytoplasmic Ca²⁺ traces were displayed in arbitrary fluorescent units.

For experiments performed with the line scan configuration, INS-1 832/13 cells and EndoC-βH1 cells were plated, transfected, and loaded as previously described, except Fluo8 AM (2 μM) used instead in EndoC-βH1 cells. Imaging was again performed using a Zeiss LSM510 inverted confocal fluorescence microscope with a 100x/1.45 objective with Fluo4/Fluo8 excited at 488 nm and emission recorded using a 505 nm emission filter. For each recording, a single line was drawn through the center of 1–3 cells. This was the only part of the field of view that was measured during the recording. The pinhole diameter was reduced to 256 μm to minimize signals from outside the focal plane (1.4 μm section). This allowed the Ca²⁺ signals to be tracked as they propagated from the edge to the center of the cells with minimal interference from Ca²⁺ diffusing from above or below the focal plane. Recordings were performed at a frequency of 333 Hz (scan time = 3 ms) and a duration of 6000 cycles.

2.8. Single-cell live mitochondrial Ca²⁺ measurements

Single-cell [Ca²⁺]_mito measurements were carried out using genetically encoded mitochondrial-targeted Ca²⁺ biosensor mito-case12 (Evrogen cat# FP992.). INS-1 832/13 and HEK-293T cells were seeded onto poly-d-lysine-coated 8-well-chambered cover glasses (Lab-Tek, Thermo Fisher Scientific) at a cell density of 70,000 cells/cm². Twenty-four h after seeding, the cells were co-transfected with siRNA and 1 μg of plasmid encoding mito-case12 at approximately 50% cell confluency. The cells were further grown for 48–72 h before measurements. On the day of imaging, the cells were pre-incubated at 37 °C in 400 μL of buffer P containing 2.8 mM of glucose. After 1.5 h, the cells were imaged using a 488 nm laser and emission recorded at 505–530 nm on a Zeiss LSM510 inverted confocal fluorescence microscope using a 100x/1.45 objective. Images were recorded at a frequency of 0.25 Hz (scan time = 3.93 s).

2.9. Live cell mitochondrial-free Ca²⁺ measurements using Rhod2 AM

EndoC-βH1 cells were seeded onto poly-d-lysine-coated 8-well-chambered cover glasses at a density of 70,000 cells/cm² (Lab-Tek, Naperville, IL, USA). The day after seeding, the cells were transfected with siRNA and incubated for 72 h prior to imaging. On the day of imaging, the cells were pre-incubated at 37 °C in 400 μL of buffer P containing 2.8 mM of glucose. After 1.5 h of incubation, 2 μM of Rhod-2 AM (Abcam) was loaded for 30 min at 37 °C in buffer P. The cells were then washed once with buffer P to remove excess dye prior to imaging. Excitation of Rhod-2 AM was performed at 543 nm and emission recorded at 590–615 nm on a Zeiss LSM510 inverted confocal fluorescence microscope. For single mouse pancreatic islet mitochondrial-free Ca²⁺ measurements, poly-d-lysine-coated 8-well-chambered cover glasses containing single complete or partially digested mouse pancreatic islets pre-incubated at 37 °C in 400 μL of buffer P containing 2.8 mM of glucose. After 1.5 h of incubation, 2 μM of Rhod-2 AM (Abcam) was loaded for 30 min at 37 °C in buffer P. The cells were then washed once with buffer P to remove excess dye prior to imaging. Excitation of Rhod-2 AM was performed at 543 nm and emission recorded at 565–615 nm. The examination of the effect of glucose on [Ca²⁺]_mito was performed on a Zeiss LSM510 inverted confocal fluorescence microscope using a 100x/1.45 objective. Images were recorded at a frequency of 0.13 Hz (scan time = 7.86 s).

2.10. Simultaneous measurement of cell cytoplasmic-free Ca²⁺ and cell mitochondrial-free Ca²⁺

For experiments measuring the effect of 36 mM of KCl on [Ca²⁺]_c and [Ca²⁺]_mito in EndoC- βH1, the protocol was the same except that after 1.5 h of incubation, 2 μM of Rhod-2 AM (Abcam) and 2 μM of Fluo8 AM were loaded for 30 min at 37 °C in buffer P. Imaging was performed on a Zeiss LSM510 inverted confocal fluorescence microscope using a 40x/0.75 objective with images recorded at a frequency of 0.30 Hz (scan time = 3.36 s).

2.11. Single-cell live mitochondrial membrane potential (Δψ_m) measurements

INS-1 832/13 cells were seeded onto poly-d-lysine-coated 8-well-chambered cover glasses (Lab-Tek, Naperville, IL, USA) at a density of 70,000 cells/cm². Twenty-four h after seeding, the cells were transfected with siRNA and incubated for 48–72 h prior to imaging. For Δψ_m measurements, the cells were pre-incubated for 2 h in buffer P with 100 nM of TMRM (Invitrogen) and 2.5 μM of cyclosporin A. After incubation, the cells were washed once with buffer P and then imaging was performed. Under these conditions, Δψ_m measurements were performed in quench mode [28]. TMRM was present in the medium throughout the experiment. TMRM fluorescence measurements were performed using 543 nm excitation and a 560 nm long pass emission filter on a Zeiss LSM510 inverted confocal fluorescence microscope using a 40x/0.75 objective. Images were recorded at a frequency of 0.64 Hz (scan time = 1.57 s).

2.12. Single-cell live Subplasmalemmal Ca²⁺ measurements

Single-cell [Ca²⁺]_mem measurements were carried out using PeriCam-based genetically encoded inner plasma membrane-targeted Ca²⁺ probe mem-case12 (Evrogen cat# FP992.). Specifically, INS-1 832/13 and HEK-293T cells were seeded onto poly-d-lysine-coated 8-well-chambered cover glasses (Lab-Tek, Naperville, IL, USA) at a cell density of 70,000 cells/cm². After 24 h of seeding, the cells were co-transfected with siRNA and 1 μg of plasmid-encoding mem-case12 at approximately 50% cell confluency. The cells were then further grown for 48–72 h before measurements. On the day of imaging, the cells were pre-incubated at 37 °C in 400 μL of buffer P containing 2.8 mM of glucose. After 1.5 h, the cells were imaged at 488 nm excitation (8%) and emission recorded at 505–535 nm on a Zeiss LSM510 inverted confocal fluorescence microscope using a 100x/1.45 objective. Images were recorded at a frequency of 0.64 Hz (scan time = 1.57 s).

2.13. Mouse islet insulin secretion and insulin content

Islets were picked by hand as previously described [29] under a stereomicroscope and incubated in HBSS containing 2.8 mM of glucose for 2 h. This was followed by 1 h of stimulation with HBSS supplemented with 2.8 or 16.7 mM of glucose. Aliquots of incubation medium were collected and insulin secretion was measured with a mouse insulin ELISA kit (Mercodia A/B, Sweden) according to the manufacturer's instructions. To extract insulin from mouse islets for insulin content measurements, 300 μl of acid ethanol (95% ethanol, HCL (37%), and water) were added to tubes containing 10 islets. The tubes were then sonicated 3 times for 5 s. The samples then underwent 5 cycles of freezing/thawing consisting of 5 min of freezing in ethanol/dry ice followed by 5 min of thawing on ice. The samples were the centrifuged for 5 min at 13.4×g with 50 μl of supernatant removed into a new tube, stored at −20 °C, and then measured using mouse insulin ELISA.

2.14. Glucose and insulin tolerance tests

All tolerance tests were performed in anesthetized mice, using 0.25 mg of midazolam (Midazolam Panpharma, Fougères, France) combined with 0.5 mg of fluanisone and 0.02 mg of fentanyl (Hypnorm, VetPharma, Leeds, UK). For IVGTT, mice were starved for 2 h prior to tail vein injection with 1 g/kg of d-glucose. For ITT, unfasted mice were intraperitoneally injected with 0.75 mU/g of human insulin (Novo Nordisk, Clayton, NC, USA). In each case, blood was collected via retro-orbital bleeding at the indicated time points after injection. Upon completion, the samples were centrifuged and plasma collected and stored at −20 °C. Insulin and glucose concentrations were then determined by a mouse insulin ELISA kit (Mercodia, Uppsala, Sweden) and Infinity Glucose (Oxidase) Reagent (Thermo Fisher Scientific, Waltham, MA, USA), respectively. Glucose measurements for ITT were determined by an Amplex Red Glucose/Glucose Oxidase Assay kit (Thermo Fisher Scientific, Waltham, MA, USA).

2.15. Statistical analyses

All of the experiments were repeated on 3 different days with the exception of ITT, which was performed twice. Data in all of the figures are presented as mean ± SEM, and unless specifically stated; statistical significance was determined by unpaired two-tailed Student's t-tests. For imaging experiments, n in the figures represents the number of cells or regions of interest (ROIs). For islet insulin secretion and IVGT assays, n in the figures represents the number of animals. For all of the statistical comparisons, a p value of less than 0.05 was considered significant.

3. Results

3.1. Functional consequences of MICU2 knockdown in insulin-secreting cells

To understand the role of MICU2 in stimulus-secretion coupling in pancreatic β cells, we silenced MICU2 in INS-1 832/13 cells, a rodent-derived insulinoma cell line [20], and in EndoC-βH1 cells, a human embryonic insulin-secreting cell line [21,30]. Our experiments revealed that after 48 or 72 h, small interfering (si) RNA treatment was sufficient to significantly reduce the expression of MICU2 at the mRNA and protein level in both cell lines (Figure 1A–D). Notably, the expression of MCU and MICU1 was unaffected by MICU2 silencing (Supplementary Figs. 1A–B).

Figure 1.

Knockdown of MICU2 reduced mitochondrial Ca²⁺ uptake and decreased insulin secretion.MICU2 mRNA expression in INS-1 832/13 cells (A) and EndoC-βH1 cells (B) after treatment with scrambled or MICU2 siRNA. MICU2 protein expression (C) and quantification of Western blotting (D) in EndoC-βH1 cells. Mitochondrial Ca²⁺ ([Ca²⁺]_mito) determined by mito-case12 and Rhod2, respectively, in MICU2-silenced INS-1 832/13 (E and F) and EndoC-βH1 cells (G and H) after glucose stimulation. E and G show average traces while F and H show differences in maximal mitochondrial Ca²⁺ levels between control and MICU2-silenced cells. [Ca²⁺]_mito by mito-case12 and Rhod2, respectively, in MICU2-silenced INS-1 832/13 (I and J) and EndoC-βH1 cells (K and L) after stimulation with KCl; average traces (I and K) and differences in maximal [Ca²⁺]_mito (J and L) between control and MICU2-silenced cells are shown. Insulin secretion in INS-1 832/13 (M) and EndoC-βH1 cells (N); secretion after 1 h of incubation was normalized to levels in cells treated with scrambled siRNA at 2.8 mM of glucose. Data are from at least three independent experiments 72 h after addition of siRNA unless otherwise stated. Mean ± SEM are given. Comparisons were made with an unpaired two-tailed Student's t-test. ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001, and ∗∗∗∗p < 0.0001. See also Fig. S1.

Given the proposed function of MICU2 in the uniporter holocomplex, we examined the uptake of Ca²⁺ into the mitochondria upon MICU2 knockdown. To this end, we transfected INS-1 832/13 cells with a genetically encoded Ca²⁺ biosensor targeted to the mitochondrial matrix, mito-case12 [31]. Our experiments revealed that 16.7 mM of glucose raised [Ca²⁺]_mito and that this increase was attenuated by 48% after silencing of MICU2 (p < 0.0001; Figure 1E–F). Similar findings were observed in EndoC-βH1 cells using another reporter, Rhod2, as these cells are difficult to transfect with plasmids [32]. In these cells, 20 mM of glucose triggered a [Ca²⁺]_mito increase that was attenuated by 57% upon MICU2 silencing (p = 0.04; Figure 1G–H). MICU2 knockdown reduced not only the magnitude, but also the rate of [Ca²⁺]_mito elevation in EndoC-βH1 cells (113 s vs 65 s to maximal [Ca²⁺]_mito; p = 0.0202; Supplementary Fig. 1C). Both Rhod2 and mito-case12 showed “punctate” loading reflective of mitochondrial localization (Supplementary Figs. 1D–E). In INS-1 832/13 and EndoC-βH1 cells, evident from unnormalized mito-case12 and Rhod2 fluorescence, respectively, we observed that mitochondrial Ca²⁺ levels were also lower at basal glucose after Micu2 silencing (Supplementary Figs. 2A–B).

Increasing extracellular K⁺ to 36 mM depolarized the plasma membrane and activated voltage-dependent Ca²⁺ channels. This resulted in a robust increase in [Ca²⁺]_mito in control cells, while there was only a minimal response in MICU2-silenced INS-1 832/13 cells (p < 0.0001; Figure 1I–J). Thus, the diminished mitochondrial response to elevated glucose (Figure 1E–H) could not simply be ascribed to decreased coupling between glucose metabolism and plasma membrane depolarization. EndoC-βH1 cells behaved similarly to rodent insulinoma cells, in which the maximal [Ca²⁺]_mito elevation following depolarization with 50 mM of KCl was diminished by 90% in MICU2-silenced cells (p < 0.0001; Figure 1K–L). This recording was performed simultaneously with the measurement of [Ca²⁺]_c shown in Supplementary Fig. 3B.

Our data suggest that MICU2 plays a role in [Ca²⁺]_mito homeostasis in insulin-secreting cell lines, which would be predicted to impact hormone secretion. Indeed, we found that GSIS was reduced by 41% and 51% at 2.8 (p = 0.0023) and 16.7 mM of glucose (p < 0.0001), respectively, in MICU2-deficient INS-1 832/13 cells (Figure 1M). In MICU2-deficient EndoC-βH1 cells, basal insulin secretion at 2.8 mM of glucose increased by 46% (p = 0.0245; Figure 1N). However, secretion at 20 mM of glucose did not increase; accordingly, the net fold change (2.8 mM–20 mM of glucose) in insulin secretion was markedly and significantly reduced (1.4-fold vs 3.1-fold; p = 0.0018).

3.2. MICU2 deficiency impaired bioenergetic functions in insulin-secreting cells

The mitochondrial membrane potential Δψ_m is an essential facet of mitochondrial bioenergetics underlying both ATP synthesis and Ca²⁺ uptake into the mitochondrial matrix via MCU [33]. The mitochondrial membrane potential is influenced by uptake of Ca²⁺ into the matrix [34]. Given the potential regulatory role of MICU2 in this process and the observed attenuation of mitochondrial Ca²⁺ uptake, we monitored the relative changes in Δψ_m with tetramethylrhodamine methyl ester (TMRM) in quench mode. Indeed, a rise in glucose from 2.8 to 16.7 mM hyperpolarized Δψ_m in INS-1 832/13 cells and this response decreased by 60% after silencing of MICU2 (p < 0.0001; Figure 2A–B). It therefore appears plausible that decreased TCA cycle activation after silencing was the cause of the decreased hyperpolarization. Under basal conditions, unnormalized TMRM fluorescence was stronger in Micu2-silenced cells, indicating that mitochondria were depolarized to a greater extent than in control cells (Supplementary Fig. 2C).

Figure 2.

Bioenergetic consequences of MICU2 knockdown in INS-1 832/13 cells. Glucose-induced mitochondrial hyperpolarization of the inner mitochondrial membrane upon MICU2 knockdown in INS-1 832/13 cells measured by TMRM (A and B); average traces (A) and difference in maximal hyperpolarization (B) between control and MICU2-silenced cells are shown. The cytosolic ATP/ADP ratio in response to an elevation in glucose from 2.8 to 16.7 mM upon MICU2 knockdown in single INS-1 832/13 cells monitored with Perceval HR (C and D); average traces (C) and difference in the maximal increase in the ATP/ADP ratio (D) between control and MICU2-silenced cells are shown. Data are from at least three independent experiments 72 h after adding siRNA. Mean ± SEM are given. Comparisons were made with an unpaired two-tailed Student's t-test. ∗∗p < 0.01 and ∗∗∗∗p < 0.0001.

In light of this, we examined the effect of MICU2 deficiency on glucose-stimulated changes in the cytosolic ATP/ADP ratio using live cell imaging of INS-1 832/13 cells expressing the fluorescent ATP/ADP reporter Perceval HR [25]. We found that knockdown of MICU2 significantly reduced the increase in the cytosolic ATP/ADP ratio upon stimulation with 16.7 mM of glucose (Figure 2C), with the maximal cytosolic ATP/ADP ratio increase diminished by 36% after stimulation with 16.7 mM of glucose in MICU2-silenced INS-1 832/13 cells (p = 0.0013; Figure 2C–D). However, as previously stated, the decreased [Ca²⁺]_mito response to elevated glucose could not be solely ascribed to decreased bioenergetic coupling, since it was also apparent when KCl was elevated. Basal levels of ATP production also appeared to be diminished as evident from reduced unnormalized Perceval HR fluorescence at 2.8 mM of glucose (Supplementary Fig. 2D).

3.3. MICU2 controlled both mitochondrial and cytosolic Ca²⁺ entry

In the next set of experiments, we examined whether MICU2 also influences bulk [Ca²⁺]_c in insulin-secreting cells. Glucose-induced Ca²⁺ oscillations were reduced in Micu2-silenced INS-1 832/13 cells (Supplementary Fig. 3A). However, as this could be the result of lower ATP production rather than a direct effect of MICU2 on [Ca²⁺]_c dynamics, the cells were then depolarized non-metabolically by adding 36 mM of KCl. We predicted that depolarization-evoked [Ca²⁺]_c increases would be more pronounced in MICU2-silenced cells due to reduced buffering by the mitochondria. Surprisingly, we observed that the maximal [Ca²⁺]_c increase upon stimulation with 36 mM of KCl decreased by 48% in MICU2-silenced INS-1 832/13 cells (p < 0.0001; Figure 3A–B). Similar results were obtained in EndoC-βH1 cells, in which silencing of MICU2 led to a 41% reduction in the maximal [Ca²⁺]_c elevation upon 50 mM of KCl stimulation (p = 0.0019; Supplementary Figs. 3B–C). This implied that the role of MICU2 in this paradoxical regulation of [Ca²⁺]_c was conserved in β cells of both rat and human origin. The measurement of [Ca²⁺]_c was performed simultaneously with the Rhod2 imaging shown in Figure 1K.

Figure 3.

Cytosolic and subplasmalemmal Ca²⁺ levels in response to KCl stimulation were perturbed in MICU2-silenced INS-1 832/13 cells. Cytosolic Ca²⁺ levels ([Ca²⁺]c) in MICU2-silenced INS-1 832/13 cells (and scramble control) upon stimulation with 36 mM of KCl as measured by Fluo4 (A and B). Average traces are shown in A, while B shows differences in maximal Fluo4 signals. (C) Image of mem-Case12 staining, n. b. specific membrane localization. Submembrane Ca²⁺ levels of [Ca²⁺]_mem in MICU2-silenced INS-1 832/13 cells upon stimulation with 36 mM of KCl measured using mem-case12 (D and E); average traces are shown in D, while E shows iAUC quantification of KCl's effect on [Ca²⁺]_mem. In (F and G), similar experiments are shown but using oligomycin and antimycin to block mitochondrial Ca²⁺ uptake prior to KCl addition, with oligomycin applied alone as a control. Mean ± SEM are given. (H) Example of line scan selection: the white arrow through the center of the cell is the only part of the field of view that was imaged; the accompanying trace within the white rectangle shows the fluorescence signal for each corresponding point along the arrow. (I) Mean Ca²⁺ intensity across INS-1 832/13 cells ∼500 ms after initiation of response to 36 mM of KCl. Cells had different sizes, so the data were interpolated to standardize each cell to a set number of points, allowing for comparison. The X axis represents the cell axis moving from one side of the plasma membrane (−50) through the center of the cell (0) to the opposite side of the plasma membrane (50). Points close to the extremities represent the submembrane region. (J) Average trace of cytosolic Ca²⁺ at cell edge and cell center in MICU2-silenced INS-1 832/13 cells (and scramble control). (K) iAUC quantification of [Ca²⁺]_c at the cell edge and cell center in MICU2-silenced INS-1 832/13 cells (and scramble control). Subplasmalemmal Ca²⁺ levels ([Ca²⁺]_mem) in MICU2-silenced INS-1 832/13 cells (and scramble control) upon stimulation with 36 mM of KCl. (L) Insulin secretion in control and Micu2-silenced INS-1 832/13 cells after 15 min of incubation in 36 mM of KCl. Comparisons were made with an unpaired two-tailed Student's t-test. In 3L, statistical comparisons were made by a paired two-tailed Student's t-test since variability in short-term incubations was greater. ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001, and ∗∗∗∗p < 0.0001.

Given our findings, we hypothesized that the reduced depolarization-evoked [Ca²⁺]_c increases after MICU2 knockdown (and mitochondrial depolarization) were due to reduced net Ca²⁺ entry across the plasma membrane (diminished entry or enhanced extrusion). To functionally elucidate this, we imaged INS-1 832/13 cells using a genetically encoded Ca²⁺ biosensor targeted to the inner leaflet of the plasma membrane mem-case12 [[35], [36], [37]]. Figure 3C shows the sensor's plasma membrane localization allowing the subplasmalemmal-free Ca²⁺ concentration ([Ca²⁺]_mem) to be determined. If mitochondrial Ca²⁺ sequestration was indeed facilitating net Ca²⁺ entry into the cell by removing Ca²⁺ from the subplasma membrane compartment, then it would be predicted that cells with defective mitochondrial Ca²⁺ transport would show enhanced subplasmalemmal Ca²⁺ elevation in response to depolarization by KCl. Indeed, the maximal [Ca²⁺]_mem elevation was 20% greater in MICU2-deficient INS-1 832/13 cells than in control cells (p = 0.0118; Figure 3D). The incremental area under the curve (iAUC) was also 43% greater in the MICU2-deficient cells (p = 0.015 vs control; Figure 3E).

To establish whether this effect on [Ca²⁺]_mem could be accounted for by restricting MCU activity, we used an alternative strategy to inhibit mitochondrial Ca²⁺ transport [38]. In the presence of the ATP synthase inhibitor oligomycin, ATP is maintained by glycolysis and Δψ_m is retained, or even elevated, allowing mitochondrial Ca²⁺ transport to occur normally. Further adding an electron transport inhibitor such as antimycin rapidly collapses Δψ_m with no a priori effect on (glycolysis-maintained) cytosolic ATP production. Mitochondrial Ca²⁺ transport, however, being dependent upon Δψ_m, is abolished [38]. We found that although treating control INS-1 832/13 cells with oligomycin/antimycin did not increase the maximal [Ca²⁺]_mem elevation, the Ca²⁺ concentration remained elevated longer at the membrane, with a delayed return to the basal state (Figure 3F). This was reflected by the iAUC of the effect increasing by 58% compared to oligomycin control (p = 0.0026; Figure 3G). Thus, this treatment had a comparable effect on [Ca²⁺]_mem as MICU2 deficiency. An important proviso is that the capacity of INS-1 832/13 cells to accelerate glycolysis in the presence of oligomycin is limited by their low expression of lactate dehydrogenase [39]; therefore, experiments were performed during the time line before which ATP depletion became significant as indicated by the unperturbed mitochondrial membrane potential during this time (data not shown). Furthermore, using oligomycin alone as a control implied that any effect upon ATP production could not explain the reduced extrusion of [Ca²⁺]_mem in oligomycin/antimycin-treated INS-1 832/13 cells and must therefore have been due to a decrease in mitochondrial Ca²⁺ uptake.

To further examine whether subplasmalemmal Ca²⁺ elevations occur when mitochondrial Ca²⁺ uptake is abrogated, we imaged INS-1 832/13 cells loaded with Fluo4 using the line scan configuration. This allows repeated imaging across a line through a cell axis as opposed to the entire field of view (Figure 3H). Crucially, this method allows the higher temporal resolution required for tracking Ca²⁺ signal propagation following membrane depolarization. To this end, cell(s) were selected, after which the starvation medium was rapidly replaced with medium containing 36 mM of KCl and the recording started simultaneously. It was expected that MICU2-deficient cells would show enhanced subplasmalemmal Ca²⁺ elevation in response to depolarization by KCl, reflected by a stronger fluorescence signal intensity at the edge of the line scan. We found that, upon KCl-induced depolarization, Ca²⁺ accumulated at the subplasma membrane compartment in siMICU2-treated INS-1 832/13 cells (Figure 3I–K) with a 1.85-fold higher Ca²⁺ signal (measured as iAUC) at the edge of the cell compared to its center (p = 0.0266). Furthermore, there was a 2.5-fold higher Ca²⁺ response at the submembrane compartment in MICU2-deficient INS-1 832/13 cells compared to controls (p = 0.0006 vs controls; Figure 3K). In contrast, there was no significant difference in the Ca²⁺ signal between the edge and center in the control cells (p = 0.8027; Figure 3K). Note that when analyzing the line scan data, the nucleus was avoided when selecting the “center” of the cell.

To ensure that the effects on [Ca²⁺]_mem were not the result of off-target effects of Micu2 siRNA on voltage-gated Ca²⁺ channel expression, expression of L-type and P/Q-type Ca²⁺ channels in Micu2-silenced INS-1 832/13 cells were determined by qPCR: no difference in expression was observed (Supplementary Fig. 4A). We repeated the line scan experiments in EndoC-βH1 cells but were unable to see any difference between control and Micu2-silenced cells (Supplementary Fig. 4B). Using mem-case12 in Endo C-βH1 cells was not possible due to the cytotoxic effects of the transfection protocols.

Collectively, these observations support a model in which MICU2-deficient mitochondria in the vicinity of the plasma membrane are less capable of removing Ca²⁺ from this compartment. Our results thus indicate not only that MICU2 plays a critical role in controlling MCU and Ca²⁺ homeostasis, but also that mitochondria play a critical role in buffering local subplasmalemmal Ca²⁺ as previously observed [40], facilitating net Ca²⁺ entry into the cell. This may occur either by decreasing cytosolic Ca²⁺-mediated desensitization of voltage-dependent Ca²⁺ channels in the plasma membrane [41,42] or alternatively by restricting Ca²⁺ efflux by plasma membrane Ca²⁺-ATPases [43,44]. The latter was unlikely in the present experimental situation since MICU2 deficiency inhibited an increase in the ATP/ADP ratio, which will limit access to ATP for the ATPases. To assess how this change in Ca²⁺ dynamics ultimately affects insulin secretion, INS-1 832/13 cells were exposed to 36 mM of KCl for 15 min: insulin secretion was reduced in MICU2-silenced cells (Figure 3L).

3.4. Micu2-deficient mice showed defective stimulus-secretion coupling and insulin secretion

Our experiments suggested that perturbation of MICU2 led to reduced mitochondrial Ca² uptake, resulting in perturbed Ca²⁺ entry into the cell and, consequently, defective insulin secretion in both rodent-derived INS-1 832/13 and human EndoC-βH1 cell lines. To understand the consequences of MICU2 deficiency in a more physiological setting, we used a global Micu2 knockout mouse [23]. As previously observed in insulin-secreting cell lines, glucose-stimulated (20 mM) maximal [Ca²⁺]_mito elevation was diminished by 57% in Micu2 knockout islets compared to wild-type islets (p < 0.0001; Figure 4A–B). Note that the KCl response was also diminished (Figure 4A), similar to that observed in the cell lines. These results suggested that MICU2 most likely plays a similar role in maintenance of [Ca²⁺]_mito homeostasis in primary cells to that observed in cell lines.

Figure 4.

Micu2 knockout mice exhibited deficiencies in mitochondrial Ca²⁺ homeostasis, ATP/ADP ratio, and insulin secretion. Mitochondrial Ca²⁺ levels [Ca²⁺]_mito in islets isolated from Micu2 knockout (KO) and wild-type mice measured with Rhod2 after stimulation with 20 mM of glucose (A and B). Average traces are shown in (A) while (B) shows differences in maximal Rhod2 signals. A total of 40 islets from 7 wild-type mice and 42 islets from 6 Micu2 KO mice were assayed in response to 20 mM of glucose and 36 mM of KCl. Changes in the cytosolic ATP/ADP ratio in response to 20 mM of glucose in Micu2 KO and wild-type islets were determined by Perceval (C) n = 14 and 27 islets from 4 to 6 wild-type and KO islets, respectively. Average traces are shown in (C) while (D) shows differences in maximal Perceval signals. Insulin secretion from Micu2 KO and wild-type islets at 2.8 and 16.7 mM of glucose after 1 h of incubation (E); (F) shows the fold response of insulin. (G) Insulin content in Micu2 KO and wild-type islets. Plasma glucose (H) and insulin (I) levels following an intravenous glucose tolerance test in Micu2 KO and wild-type mice; n denotes the number of mice. Insets show the area under the curve for glucose (AUC; H) and insulin (J; T = 0–5 min). Data are from at least three independent experiments. (I) Insulin tolerance test (ITT) in Micu2 KO and wild-type mice. Glucose values are expressed as the percentage of glucose in which glucose at 0 min was 100%; n denotes the number of mice. Inset shows the AUC. Mean ± SEM are given. Comparisons of imaging and insulin secretion were made with an unpaired two-tailed Student's t-test (∗p < 0.05 and ∗∗∗∗p < 0.0001) and the Mann–Whitney U-test (∗∗∗p < 0.001) for data from IVGTT.

Furthermore, as [Ca²⁺]_mito homeostasis is critical for cellular bioenergetics, we speculated that the Micu2 knockout might perturb the cytosolic ATP/ADP ratio in primary cells, as was seen in the insulin-secreting cell lines. Therefore, in the next set of experiments, we used an adenovirus expressing the biosensor Perceval, originally described as an ATP/ADP sensor [26], but later shown to report cytoplasmic ATP in mouse islet cells [45]. The maximal Perceval signal was diminished by 50% in Micu2 knockout islets compared with wild-type islets upon stimulation with 20 mM glucose (p < 0.0001; Figure 4C–D). A likely consequence of an abrogated increase in the ATP/ADP ratio in pancreatic β cells is impaired insulin secretion. Batch incubations of Micu2 knockout islets demonstrated that basal (2.8 mM of glucose) insulin secretion was significantly higher compared to that in their wild-type counterparts (p = 0.0427; Figure 4E). However, Micu2 knockout islets failed to release insulin in response to high glucose as efficiently as wild-type islets as illustrated by a 50% reduction in the fold response of insulin secretion (16.7–2.8 mM of glucose; p = 0.0362; Figure 4F). Notably, insulin content was found to be increased in Micu2 knockout islets compared with wild-type islets (p < 0.05; Figure 4G). This effect was most likely due to the accumulation of insulin that was not released since INS1 mRNA expression was similar in wild-type and Micu2 knockout islets (Supplementary Fig. 5A). Supplementary Fig. 5B shows the islet secretion data normalized to the insulin content derived from the analysis shown in Figure 4G: in vitro results shown this way reflect the in vivo observations, in that stimulated insulin secretion was abrogated while basal secretion was unchanged.

To ensure that any effects were not due to changes in the expression of the mitochondrial complexes, we quantified the protein expression of complexes I–V by Western blotting. There were no significant differences between wild-type and Micu2 knockout islets (Supplementary Figs. 5C–D). Our experiments thus suggested that stimulus-secretion coupling in β cells in Micu2 knockout mice was defective and underlied perturbed GSIS, reminiscent of that found in EndoC-βH1 cells (Figure 1N).

In the next set of experiments, we examined the consequences of Micu2 deficiency on whole-body glucose homeostasis. Glucose elimination was not significantly different between the Micu2 knockout and wild-type mice (Figure 4H). However, an intravenous glucose tolerance test (IVGTT) revealed a marked reduction in the initial release of insulin (p = 0.0003; Figure 4I): we observed a 74% and 70% reduction in the plasma insulin levels at 1 and 5 min after glucose administration, respectively, in Micu2 knockout mice compared to wild-type mice. Because insulin secretion was impaired while glucose elimination was unaltered in Micu2 deficiency, we performed an insulin tolerance test (ITT). Both strains of mice responded robustly to insulin, with decreased plasma glucose levels (Figure 4J). These experiments thus demonstrated that the initial release of insulin in Micu2 knockout mice was reduced, likely caused by a stimulus-secretion coupling defect in pancreatic β cells. Hence, Micu2 knockout mice exhibited deficient GSIS both in vitro and in vivo.

3.5. MICU2 regulation of Ca²⁺ in mitochondria, cytosol, and subplasmalemmal space was conserved between cell types

Previous studies in neuronal cells showed that inhibition of ATP generation and dissipation of mitochondrial membrane potential led to reduced [Ca²⁺]_c upon stimulation with glutamate or 50 mM of KCl [46]. Therefore, having shown the role of MICU2 in regulating [Ca²⁺]_mito and [Ca²⁺]_c in insulin-secreting cells, we asked whether this mechanism may also be observed in other cell types. High-throughput gene expression assays previously demonstrated that the human embryonic kidney-derived HEK-293T cell line robustly expresses levels of MICU2 mRNA [47]. It has been shown that this cell line also expresses voltage-dependent Ca²⁺ channels and exhibits endogenous Ca²⁺ currents [48,49]. We therefore used this cell line to address the role of MICU2 in cellular Ca²⁺ homeostasis.

MICU2 was effectively silenced on the mRNA and protein levels in HEK-293T cells (Supplementary Figs. 6A–C). Measurements of [Ca²⁺]_mito with mito-case12 demonstrated that stimulation with 36 mM of KCl increased [Ca²⁺]_mito in HEK293T cells and that this response was attenuated by 46% after silencing of MICU2 (p < 0.0001; Figure 5A–B). In addition, time to maximal [Ca²⁺]_mito peak in MICU2-silenced cells was 126 s compared to 107 s in controls after 36 mM of KCl stimulation (p = 0.0087; Supplementary Fig. 6D). [Ca²⁺]_c measurements using Fluo5F showed that membrane depolarization by 50 mM KCl triggered a [Ca²⁺]_c increase and the maximal [Ca²⁺]_c elevation was reduced by 74% in MICU2-silenced HEK-293T cells compared to control cells (p < 0.0001; Figure 5C–D). For experiments measuring [Ca²⁺]_mem using mem-case12, oligomycin was added to ensure that any effects of MICU2 silencing were independent from the resultant bioenergetic effects of increased [Ca²⁺]_mito. The same was not done in the experiments with INS-1 832/13 cells due to ATP production in the β cell being finely tuned to the extracellular glucose concentration as a consequence of the low-affinity enzymes GLUT2 and glucokinase and downstream dissipative pathways (high leak current) [50]. This means that at low glucose, metabolic flux and ATP production in INS-1 832/13 cells are low compared to HEK293T cells. Therefore, any effect of increased [Ca²⁺]_mito on ATP production would be unlikely to reach the threshold to affect K_ATP channel activity, meaning using oligomycin was not necessary. Reminiscent of the findings in INS-1 832/13 cells, [Ca²⁺]_mem elevation upon 50 mM of KCl stimulation increased by 35% in MICU2-silenced HEK293T cells compared to the control cells (p = 0.0103; Figure 5E–F). Thus, these data demonstrate a conservation of the regulation of [Ca²⁺]_mito and [Ca²⁺]_c by MICU2 in different cell types.

Figure 5.

MICU2 played a similar role in Ca²⁺ homeostasis in HEK-293T cells as observed in insulin-secreting cells. Mitochondrial Ca²⁺ ([Ca²⁺]_mito) determined by mito-case12 in MICU2-silenced HEK-293T cells stimulated with 36 mM of KCl (A and B); (A) shows average traces while (B) shows differences in the maximal mitochondrial Ca²⁺ response. Cytosolic Ca²⁺ levels ([Ca²⁺]_c) determined by Fluo5F in MICU2-silenced HEK-293T cells upon stimulation with 50 mM of KCl (C and D); average traces are shown in (C) while (D) shows differences in maximal Fluo5F signals. Subplasmalemmal Ca²⁺ levels ([Ca²⁺]_mem) in MICU2-silenced HEK-293T cells upon stimulation with 50 mM of KCl (E and F); average traces are shown in (E) while (F) shows differences in maximal mem-case12 signals. Data are from at least three independent experiments 72 h after adding siRNA. Mean ± SEM are given. Comparisons were made with an unpaired two-tailed Student's t-test. ∗p < 0.05, ∗∗p < 0.01, and ∗∗∗∗p < 0.0001. See also Fig. S3.

4. Discussion

It is undisputed that an increase in [Ca²⁺]_c triggers insulin secretion as well as secretion of most other hormones, but it has also been recognized that such a rise in itself does not sustain insulin secretion [51,52]. Cytosolic Ca²⁺ is further transported into the mitochondrial matrix, and an elevation in free matrix Ca²⁺ has been considered to trigger additional regulatory factors [53]. Prior to the molecular identification of the components of the uniporter holocomplex [19,54,55], indirect approaches were required to investigate the effect of modulating [Ca²⁺]_mito, none of which were entirely satisfactory. Removal of external Ca²⁺ [56] affects multiple non-mitochondrial processes, not least exocytosis itself. Moreover, attempts to dampen changes in [Ca²⁺]_mito by increasing the matrix Ca²⁺-buffering capacity by targeting exogenous Ca²⁺-binding proteins to the matrix [5] or loading cells with Ca²⁺-chelator 1,2-bis(o-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid (BAPTA) [57] each succeeded in reducing the bioenergetic response to elevated glucose, but concerns remain about toxicity associated with this treatment [50].

The molecular characterization of the uniporter [19,54,55,58] as well as a number of accessory proteins such as MICU1, 2, 3, and EMRE [19,55,59] opened the possibility of genetic manipulation of the uniporter's components [60]. It was shown that silencing of Mcu abrogates the increase in [Ca²⁺]_mito and the cytosolic ATP/ADP ratio in response to glucose in mouse β cells [61]. Another study demonstrated that silencing of MCU in clonal INS-1-E cells resulted in reduced nutrient-induced hyperpolarization of the inner mitochondrial membrane, ATP production, and GSIS [9]. Moreover, knockdown of MICU1 leads to reductions in [Ca²⁺]_mito, cytosolic ATP levels, and GSIS in clonal insulin-secreting cells [8]. However, the role of MICU2 remains more enigmatic. MICU2 is a peripheral membrane protein associated with the mitochondrial inner membrane and facing the intermembrane space. The absence of MICU2 has been associated with reduced Ca²⁺ uptake in vivo in mouse liver, though silencing in this system also leads to a decrease in the abundance of the pore-forming subunit MCU and the complex [19]. Observations in other cell systems suggested that the accessory protein may have a regulatory role in Ca²⁺ uptake into mitochondria [13,16]. MICU1 and MICU2 are thought to heterodimerize via disulfide bridges and thus regulate Ca²⁺ entry into the mitochondrial matrix via the MCU [62]. Indeed, both MICU1 and MICU2 have been shown to play an important role in setting the threshold for mitochondrial Ca²⁺ uptake in HeLa cells at variable Ca²⁺ concentrations [16]. Furthermore, loss of MICU1 or MICU2 in permeabilized HEK-293T cells has also been shown to disrupt this critical threshold for mitochondrial Ca²⁺ uptake [13,14,63,64].

Herein, using MICU2-deficient rodent and human insulin-producing cell lines as well as Micu2 knockout mice, we were able to show that reduced Ca²⁺ uptake into mitochondria resulted in inhibition of GSIS. This confirmed observations in previous studies [8,9,61] showing that [Ca²⁺]_mito affects mitochondrial metabolism and insulin secretion. However, our experiments could not resolve how Micu2 knockout mice maintained euglycemia despite reduced early insulin secretion. Insulin tolerance, albeit a rough estimate of insulin action in vivo, was unchanged. In fact, we previously reported that ITT is not sensitive enough to reflect changes in insulin sensitivity in the face of reduced early insulin secretion in vivo [65]. This phenotype is also somewhat similar to what was observed in β cell-specific Mcu knockout mice that exhibit impaired insulin secretion 5 min after intraperitoneal glucose injection with improved glucose tolerance despite normal insulin sensitivity [15]. A hyperinsulinemic euglycemic clamp, which is beyond the scope of the present study, would be required to accurately determine insulin sensitivity. Importantly, MCU and MICU1 expression were unaffected by MICU2 silencing in INS-1 832/13 cells, indicating that MICU2 plays a fundamental non-redundant role in regulating [Ca²⁺]_mito in β cells. However, the mechanisms by which MICU2 and [Ca²⁺]_mito might exert control over GSIS were unexpected and add a new dimension to how mitochondrial Ca²⁺ transport might regulate β cell function and insulin secretion.

The key observation novel in the context of β cells is that mitochondrial Ca²⁺ uptake exerts a profound stimulatory effect on the net flux of cations across the plasma membrane, either by enhancing entry or depressing efflux. Insulin-secreting cells have a high concentration of subplasma membrane mitochondria, with roughly 20% of the membrane inner surface associated with mitochondria exerting a strong Ca²⁺-buffering effect [40]. Mitochondria are dynamic organelles and submembrane Ca²⁺ buffering is reduced upon plasma membrane depolarization because of Ca²⁺-dependent remodeling of the cortical F-actin network causing translocation of subplasmalemmal mitochondria to the cell interior [40]. In line with these findings, we demonstrated that restricted mitochondrial Ca²⁺ uptake resulted in more pronounced depolarization-induced increases in Ca²⁺ in the submembrane compartment. However, under the same conditions, the global cytoplasmic Ca²⁺ increases were attenuated.

To rationalize a decreased Ca²⁺ response to plasma membrane depolarization in both the matrix and cytosolic compartments, the simplest assumption is that net Ca²⁺ influx across the plasma membrane is inhibited. Increased extrusion of Ca²⁺ is less likely given the observed decrease in the ATP/ADP ratio as a consequence of MICU2 deficiency and that extrusion is an ATP-dependent process. While the endoplasmic reticulum does take up Ca²⁺ in response to the elevated ATP/ADP ratio at high glucose, the response is too slow to account for a Ca²⁺ deficit [66]. In any case, the effect of restricted mitochondrial Ca²⁺ uptake on bulk [Ca²⁺]_c is seen with elevated K⁺ as well as with high glucose and is thus not directly dependent on bioenergetics. Had this effect only been observed in response to glucose, then the previously postulated positive effect of mitochondrial Ca²⁺ uptake on mitochondrial dehydrogenases would have been the most plausible explanation [67].

A paradoxical decrease in [Ca²⁺]_c elevation following inhibited transport of mitochondrial Ca²⁺ was previously observed in cultured neurons [38,46]. Since these studies predated the molecular identification of the MCU, mitochondrial Ca²⁺ transport was inhibited by mitochondrial depolarization in the presence of the ATP synthase inhibitor oligomycin together with a rotenone or antimycin electron transport inhibitor. The control was oligomycin alone: both control and mitochondrial depolarized cells maintained similar ATP/ADP ratios supported by glycolysis, but the [Ca²⁺]_c transients induced by Ca²⁺ entry through NMDA-selective glutamate receptors or voltage-dependent Ca²⁺ channels drastically reduced. In the present study, treatment with oligomycin + antimycin further confirmed a role of mitochondrial Ca²⁺ uptake in clearing subplasmalemmal Ca²⁺ in insulin-secreting cells, and the observations were similar to those in neurons [38,46]. Importantly, the net neuronal accumulation of ⁴⁵Ca²⁺ is strongly inhibited by mitochondrial depolarization [38,46], confirming the organelle's ability to control net flux across the plasma membrane. Notably, mitochondrial Ca²⁺ uptake has also been implicated in store-operated Ca²⁺ entry, with MCU knockdown in RBL mast cells shown to reduce Ca²⁺ influx through Ca²⁺ release-activated channels [68].

It was hypothesized that mitochondria are able to deplete a subplasma membrane layer of elevated Ca²⁺ [38,46]. Voltage-dependent Ca²⁺ channels become desensitized when local Ca²⁺ accumulations are not promptly removed [44,69,70]. In addition, it has been shown in chromaffin cells, neurons, and cardiac muscle that mitochondrial sequestration of subplasmalemmal Ca²⁺ prevents desensitization of Ca²⁺ channels and thus might uphold hormone secretion [[71], [72], [73]]. Furthermore, mitochondria play an essential role in the maintenance of [Ca²⁺]_c amplitude wave propagation via feedback regulation of plasma membrane channel activity in diverse cellular systems, including rat cortical astrocytes and Xenopus laevis oocytes [74,75]. The response of the subplasmalemmal-targeted Ca²⁺ probe mem-case12 and line scan imaging in the present study provided support for the existence of such a layer of elevated Ca²⁺ [38,46] during β cell depolarization and its depletion by mitochondrial Ca²⁺ uptake. Notably, in the mem-case12 experiments, there was a difference between the effects on [Ca²⁺]_mem between Micu2 silencing and treatment with oligomycin/antimycin, whereby the former caused an increase in the duration of the subplasmalemmal Ca²⁺ increase while the latter resulted in an increased magnitude of the response. This may be explained by the different mechanisms by which mitochondrial Ca²⁺ uptake is inhibited. When MICU2 is deficient, uptake at low Ca²⁺ is likely to increase as the protein plays an important role in suppressing MCU activity under these conditions [11,13,16,17]. Subsequently, when [Ca²⁺]_c is elevated, the mitochondrial buffering capacity decreases. In contrast, oligomycin and antimycin prevent proton transport from the mitochondrial matrix to the mitochondrial intermembrane space, increasing the matrix pH, subsequently collapsing Δψ_m and inhibiting Ca²⁺ entry. These two methods affect the Ca²⁺ electrochemical gradient differently and as a result the Ca²⁺ uptake kinetics, potentially explaining the differential effects on [Ca²⁺]_mem.

Regarding line scan imaging, one could argue that due to its relatively low K_d, Fluo4 is unsuitable for measuring [Ca²⁺]_mem, which can reach concentrations of 10–100 μM [76]. This means there is a risk of Fluo4 saturation at the submembrane compartment. This did not seem to occur in the control, where there was little difference in the Fluo4 signal between the edge and center of the cell. In MICU2-silenced cells, however, there was a clear difference between the edge and center of the cell. If saturation was indeed occurring, this would imply that the observed increase in [Ca²⁺]_mem was in fact an underestimation. Moreover, the saturation problems were mitigated against the complementary findings using the lower-affinity mem-case12 probe (K_d = 1 μM) [36].

The lack of a [Ca²⁺]_mem effect on MICU2-silenced Endo C-βH1 cells may lie in differences in the composition of voltage-gated Ca²⁺ channels between the two cell lines. Notably, the L-type Ca²⁺ channel current, which is largely responsible for Ca²⁺ entry upon high K⁺ stimulation due to the rapid inactivation of other Ca²⁺ channels [77], only accounts for 20–30% of the Ca²⁺ current in Endo C-βH1 cells [78] compared to at least 60% of the Ca²⁺ current in INS-1 832/13 cells [79]. This means the [Ca²⁺]_mem levels attained in EndoC-βH1 cells in the line scan experiment may still be able to be cleared by MICU2-deficient mitochondria. Thus, any buildup of Ca²⁺ in the subplasmalemmal compartment may occur at a slower rate, evading detection in our experiments. This would also explain the smaller effect of MICU2 silencing on depolarization-evoked [Ca²⁺]_c increases in EndoC-βH1.

This study shows that mitochondrial Ca²⁺ transport has additional consequences for β cells beyond facilitation of the TCA cycle, which may have varying effects on insulin secretion. On the one hand, attenuation of subplasmalemmal Ca²⁺ elevations could reduce the triggering of exocytosis in pre-docked vesicles responsible for first-phase insulin release. On the other hand, there is evidence that increased Ca²⁺ in the cytosol is required to transport reserve vesicles to the release site, at least in neuronal systems [80]. Based on the KCl-induced insulin secretion experiments, the attenuation of mitochondrial Ca²⁺ transport leads to a reduction in insulin secretion. First-phase insulin release occurs in the first 5–10 min [81]. Therefore, while there may be an increase in first-phase insulin secretion, an attenuation of the second phase likely caused by decreased transport of reserve vesicles means that insulin secretion decreases over the 15 min time course.

[Ca²⁺]_c increases may also occur as a consequence of endoplasmic reticulum Ca²⁺ release. Indeed, there is evidence in β cells of a basal endoplasmic reticulum Ca²⁺ leak channeled toward the mitochondria, which stimulates mitochondrial bioenergetics, priming the mitochondria for an increase in glucose [82,83]. This process has been implicated in the generation of cytosolic Ca²⁺ oscillations and GSIS [83]. Although ablating Micu2 would also abrogate this process, it appears to be distinct from the mechanism observed herein. The reduction in [Ca²⁺]_mito and the associated increase in [Ca²⁺]_mem were more or less instantaneous upon exposure to KCl and plasma membrane depolarization as opposed to the endoplasmic reticulum Ca²⁺ leak, which seems to occur at a constant rate [82,83].

It was previously demonstrated that dynamic changes in the subplasmalemmal mitochondrial population participate in the regulation of local Ca²⁺ levels and consequently exocytosis in MIN6 cells [40]. A limitation of our study is that, at this point, we have not obtained sufficient data on whether MICU2 deficiency impacts the localization, morphology, or number of mitochondria in the submembrane compartment. Changes in any of these parameters in MICU2-deficient cells would likely also affect subplasmalemmal and cytosolic Ca²⁺ dynamics.

Our results may help resolve some of the contradictions in the concept of amplification pathways and K_ATP-independent mechanisms of insulin secretion. The search for coupling factors responsible for K_ATP-independent insulin secretion began after the discovery that insulin secretion could be provoked by increasing glucose even when the K_ATP channel was kept open by diazoxide and the plasma membrane was depolarized by high concentrations of extracellular KCl, thereby opening voltage-dependent Ca²⁺ channels [51]. Metabolic coupling factors were postulated to account for the insulinotropic effects of glucose under K_ATP-independent conditions and explain why insulin secretion could not be sustained by an elevation of [Ca²⁺]_c alone [52]. However, at least the latter circumstance can be accounted for by the consequences of mitochondrial Ca²⁺ uptake that we observed herein: if Ca²⁺ is not removed from the subplasmalemmal space by mitochondria, voltage-dependent Ca²⁺ channels will be desensitized. A number of metabolites and metabolic pathways have been convincingly implicated in amplification of GSIS [84]. However, most share a similar feature: it has not been resolved how these factors stimulate exocytosis. Moreover, a number of concerns have also been raised about the bioenergetics of the formation of metabolic coupling factors [50].

5. Conclusions

In summary, we uncovered a crucial role of mitochondria in maintaining β cell excitability involving sequestering of Ca²⁺ into mitochondria located at the vicinity of the voltage-dependent Ca²⁺ channels in the plasma membrane that coupled with the well-established bioenergetic effects accounts for the regulatory effect of mitochondrial Ca²⁺ uptake on insulin secretion. The recent molecular characterization of the mitochondrial Ca²⁺ uniporter holocomplex holds promise for further elucidating the complexities of β cell stimulus-secretion coupling in which the mitochondria are at center stage.

Author contributions

N.V., A.H., D.G.N., and H.M. designed the study. N.V., A.H., A.B., A.M.B., H.B., Y.S., K.K., P.S., E.C., and M.F. performed the experiments. N.V., A.H., A.M.B., and M.F. analyzed the data. V.M. provided the Micu2 knockout mice. A.T., V.M., D.G.N., and H.M. provided critical insights to the project. N.V., A.H., A.T., V.M., D.G.N., and H.M. wrote the manuscript.

Study approval

All of the experimental animal protocols were approved by the local animal ethical committee of Lund/Malmö and complied with the guidelines of the Federation of European Laboratory Animal Science Association (FELASA).

Conflict of interest

None declared.

Appendix A. Supplementary data

The following is the Supplementary data to this article: