Abstract
L-type channel antagonists are of therapeutic benefit in the treatment of hyperlipidaemia and insulin resistance. Our aim was to identify L-type voltage-gated Ca2+ channels in white fat adipocytes, and determine if they affect intracellular Ca2+, lipolysis and lipogenesis. We used a multidisciplinary approach of molecular biology, confocal microscopy, Ca2+ imaging and metabolic assays to explore this problem using adipocytes isolated from adult rat epididymal fat pads. CaV1.2, CaV1.3 and CaV1.1 alpha1, beta and alpha2delta subunits were detected at the gene expression level. The CaV1.2 and CaV1.3 alpha1 subunits were identified in the plasma membrane at the protein level. Confocal microscopy with fluorescent antibodies labelled CaV1.2 in the plasma membrane. Ca2+ imaging revealed that the intracellular Ca2+ concentration, [Ca2 +]i was reversibly decreased by removal of extracellular Ca2+, an effect mimicked by verapamil, nifedipine and Co2+, all blockers of L-type channels, whereas the Ca2+ channel agonist BAY-K8644 increased [Ca2+]i. The finding that the magnitude of these effects correlated with basal [Ca2+]i suggests that adipocyte [Ca2+]i is controlled by L-type Ca2+ channels that are constitutively active at the adipocyte depolarized membrane potential. Pharmacological manipulation of L-type channel activity modulated both basal and catecholamine-stimulated lipolysis but not insulin-induced glucose uptake or lipogenesis. We conclude that white adipocytes have constitutively active L-type Ca2+ channels which explains their sensitivity of lipolysis to Ca2+ channel modulators. Our data suggest CaV1.2 as a potential novel therapeutic target in the treatment of obesity.
Introduction
White fat adipocytes (WFA) are the major energy depot of the body, storing and releasing energy in response to the calorific demands of the body during periods of excess and need respectively (Arner et al. 2011). It is widely accepted that an impairment of WFA triglyceride metabolism in obesity is a major aetiological factor of the metabolic syndrome and type 2 diabetes. Whereby the inability of WFA to appropriately store energy as fat leads to ectopic disposition of lipids in insulin–responsive tissues such as skeletal muscle, liver and pancreas to promote insulin resistance and associated systemic disorders (Sattar & Gill 2014). To address dysfunctional fat storage, it is first necessary to understand the regulation of lipid storage in healthy WFA.
Extracellular Ca2+ influx is implicated in the processes of fat storage (Arruda & Hotamisligil 2015): lipolysis (Schimmel 1978, Izawa et al. 1983, Allen & Beck 1986) and lipogenesis (Avasthy et al. 1988). Indeed, studies in Drosophila mutants indicate that impaired adipocyte cytosolic Ca2+ is associated with increased lipid deposition (Baumbach et al. 2014). However, the identity of the Ca2+ pathway is unclear (Draznin et al. 1988).
Several routes by which Ca2+ enters primary WFA are identified; these include voltage-gated Ca2+ channels, VGCC (Clausen & Martin 1977, Pershadsingh et al. 1989) store-operated Ca2+ channels (El Hachmane & Olofsson 2018), and reverse mode Na+/Ca2+ exchange, NCX (Pershadsingh et al. 1989, Bentley et al. 2014). Although VGCCs play a prominent role in Ca2+ entry and function of electrically excitable cells (Lipscombe et al. 2004), this route of influx is ill-defined in non-excitable adipocytes.
Pharmacological investigations show that verapamil and nifedipine, blockers of L-type VGCCs, can inhibit WFA Ca2+ uptake (Martin et al. 1975, Pershadsingh et al. 1989) and impair glucose transport (Draznin et al. 1987), lipolysis (Izawa et al. 1983) and lipogenesis (Avasthy et al. 1988). However, this inference of an association between Ca2+ influx and lipid turnover has come from independent studies where animal species, adipose depot as well as experimental conditions differed. To date, no comprehensive synthetic study exists where changes in Ca2+ influx and WFA function have been examined in primary adipocytes for the same species under similar experimental conditions of similar age and weight. Furthermore, a detailed molecular and immunohistochemical investigation of L-type VGCC expression in WFA is absent.
The aim of this study was to use a combination of Ca2+ imaging, metabolic, immunocytochemical and molecular biology techniques to identify L-type Ca2+ channels in WAT and demonstrate their functional role in lipid turnover.
Materials and methods
Unless stated otherwise, adipocyte isolation and experiments were performed in Hank’s buffer solution which composed of (in mM): NaCl 138, NaHCO3 4.2, KCl 5.6, MgCl2 1.2, CaCl2 2.6, NaH2PO4 1.2, glucose 5 and HEPES 10 (pH 7.4 with NaOH). For nominally Ca2+ free solutions, CaCl2 was equimolarly replaced with MgCl2. All % values are weight per volume. Unless stated otherwise drugs and chemicals are from Sigma.
Ethical approval
Animal care and experimental procedures were carried out in accordance with either the UK Home Office Animals (Scientific Procedures) Act (1986) or Swedish Ethical Review Board. In both instances the local ethical committees approved the animal procedures. Animals were killed by cervical dislocation or CO2.
Isolation and preparation of adipocytes
Epididymal fat pads were taken from Wistar rats (fed ad libitum, 12 h dark/light cycle, weight 220–420 g; Charles River Laboratory). For some experiments tissue from Sprague–Dawley rats was used, but since no difference could be detected, data were pooled. Adipocytes were isolated as described previously (Bentley et al. 2014).
Polymerase chain reaction
For reverse transcriptase PCR (RT-PCR) total RNA was isolated by cell lysis in TRI Reagent. Genomic DNA was removed with RQ1 RNase-free DNase (Promega) with RNA quantity and purity determined by the A260/A280 ratio (1.98 to 2.02). The RNA integrity numbers (6–8) ascertained samples were suitable for RT-PCR (Schroeder et al. 2006). First-strand cDNA synthesis used 1 µg of total RNA with 200 ng random hexamer primers and Avian Myeloblastosis Virus reverse transcriptase (10 units/µL RNA, Promega).
PCR reactions used Dream Taq PCR Master Mix (Thermo Fisher) and primers shown in Table 1. PCR was performed at 95°C for 10 min, followed by 40 cycles at 95°C for 45 s, 58°C for 45 s, 72°C for 45 s and terminated by a final extension step at 72°C for 10 min. PCR products were separated on 1% agarose gels, DNA stained with 0.5 μg/mL ethidium bromide and imaged with GeneSnap software (Syngene, UK). cDNA sequencing (DeepSeq, Nottingham, UK) checked PCR product identity.
Primer sequences.
| Name | Accession number | Sequence | Product size (bp) |
|---|---|---|---|
| β-Actin | V01217 | FWD: 5′-AGGCCCAGAGCAAGAGAG-3′ REV: 5′-CCTCATAGATGGGCACAGT-3′ |
333 |
| 18s rRNA | V01270 | FWD: 5′-TCTGCCCTATCAACTTTCGATG-3′ REV: 5′-AATTTGCGCGCCTGCTGCCTTCCTT-3′ |
137 |
| Cacna1s (CaV1.1) | U31816.1 | FWD: 5′-CAAGTCCTTCCAGGCCCTG-3′ REV: 5′-CGTAGTCAGACTCCGGGTCG-3′ |
271 |
| Cacna1c (CaV1.2) | M67515.1 | FWD: 5′-CGCATTGTCAATGACACGATC-3′ REV: 5′-CGGCAGAAAGAGCCCTTGT-3′ |
217 |
| Cacna1d (CaV1.3) | M57682.1 | FWD: 5′-TTGGTACGGACGGCTCTCA-3′ REV: 5′-CCCCACGGTTACCTCATCAT-3′ |
156 |
| Cacna1f (CaV1.4) | U31816.1 | FWD: 5′-AGCACAAGACCGTAGTGGTG-3′ REV: 5′-ATACCCCCAATGCCACACAG-3′ |
168 |
| Cacnb1 | NM_017346.1 | FWD: 5′-AGTGCCAACAGAAGCAGAAGT-3′ REV: 5′-GTGTTTGCTGGGGTTGTTGAG-3′ |
237 |
| Cacnb2 | NM_053851.1 | FWD: 5′-CTCTTCTTCCCCTGCACCAA-3′ REV: 5′-GCCTCGGCTAAGAGCAGTTT-3′ |
237 |
| Cacnb3 | NM_012828 | FWD: 5′-CCTACGCCCGGGTTTGA-3′ REV: 5′-CAAATGCCACAGGTTTGTGCT-3′ |
174 |
| Cacnb4 | NM_001105733.1 | FWD: 5′-ATGCCAGGTCTGCATGTCTC-3′ REV: 5′-ACATGGGGGTCTGGTGATCC-3′ |
231 |
| Cacna2d1 | NM_012919.3 | FED: 5′-CCAAATCTCAGGAGCCGGT-3′ REV: 5′-GCAATACCAAGGCCAAACTGT-3′ |
219 |
| Cacna2d2 | NM_175592.2 | FWD: 5′-CTGCAGGTCAAGTTGCCAAT-3′ REV: 5′-AGACGCGTTCCACTAACTGC-3′ |
262 |
| Cacna2d3 | NM_175595 | FWD: 5′-TGGACGAGAGGCTGCTTTTG-3′ REV: 5′-ATGTACGCTTCGGTCCACAC-3′ |
180 |
| Cacna2d4 | NM_001191751.1 | FWD: 5′-ATCGCCTTCGACTGCAGAAA-3′ REV: 5′-CTCTCGGTTGTCTCGATCCG-3′ |
255 |
Quantitative PCR (qPCR) reactions were performed in triplicate with SYBR® Green JumpStart TM Taq ReadyMix TM. The thermal profile was 10 min at 95°C, 40 cycles of denaturation at 95°C for 15 s, annealing for 20 s at the primer-specific temperature, and elongation at 72°C for 35 s. The resultant mean threshold cycle (Ct) values were used for gene normalization and expression analyses. Three stable reference genes with the sequences listed in Table 2 were used: glyceraldehyde-3-phosphate dehydrogenase (GAPDH), phosphoglycerate kinase 1 (PGK1) and hypoxanthine guanine phosphoribosyl transferase (HPRT1) (Gorzelniak et al. 2001, Fink et al. 2008, Silver et al. 2008). Relative mRNA levels were quantified by the method of Pfaffl (Pfaffl 2001).
Primer sequences used for qPCR analysis.
| Name | Sequence | Product size (bp) |
|---|---|---|
| Hprt1 | FWD: 5′-CGAGGAGTCCTGTTGATGTTGC-3′ REV: 5′-CTGGCCTATAGGCTCATAGTGC-3′ |
172 |
| Pgk1 | FWD: 5′-TAGTGGCTGAGATGTGGCACAG-3′ REV: 5′-GCTCACTTCCTTTCTCAGGCAG-3′ |
166 |
| GAPDH | FWD: 5′-GGCAAGTTCAATGGCACAGT-3 REV: 5′-TGGTGAAGACGCCAGTAGACTC-3′ |
183 |
Western blot analysis
Cytosolic and solubilized plasma membrane proteins were prepared by differential centrifugation. Cell lysates were prepared at 4°C with a lysis solution that contained 10 µL of protease inhibitor cocktail per 1 mL of lysis buffer (10 mM Tris, 250 mM sucrose, 1 mM EDTA, pH 7.4). Protein content was determined by Lowry. 15–20 µg of protein was resolved by SDS-PAGE on 4–20% gradient precast minigels TGX (BioRad) at 170 volts for 40 min.
Protein transfer onto nitrocellulose membranes was performed at 100 volts for 60 min in cold transfer buffer; Ponceau S staining confirmed transfer. Membranes were blocked in 5% milk in TBST (25mM Tris, pH 7.6; 125 mM NaCl; 0.1% Tween 20) for 3–4 h, at 20–22°C, followed by overnight incubation at 4°C with primary antibodies: rabbit anti-CaV1.3 at 1:200 (ACC-005, Alomone), anti CaV1.2 at 1:500 dilution (ACC-003, Alomone), anti-β-actin at 1:5000 and anti-Na,K-ATPase at 1:20,000 (Abcam). Membranes were washed with TBST and incubated for 1 h at room temperature with a 1:10,000 dilution of secondary antibodies: IRDye 800 CW goat anti-rabbit IgG (Li-COR™) and IRDye 680 CW goat anti-mouse IgG (Li-COR™). Antibody dilutions were made in 5% milk in TBST. Membranes were then washed with TBST, rinsed in H2O and imaged with Li-COR Odyssey infrared Imaging System.
Band intensity was measured using the Li-COR Odyssey software, version 2.1 (Li-COR Bioscience, USA) and analyzed using Image Studio Lite, version 5.2. For each protein band the signal intensity was normalized to that of β-actin expression, which was then expressed relative to that in brain. In initial experiments GAPDH was used as a control, but since comparison of the ratios obtained with that of β-actin expression showed no difference, the two set of data were pooled. For CaV1.2 in brain, bands spanning 130–250 kDa were combined before normalization to β-actin. Samples probed with the primary antibodies in the absence of primary antiserum indicated absence of non-specific binding.
Immunocytochemistry
Adipocytes were attached to poly-l-lysine (25–100 µg/mL)-coated coverslips and fixed with 4% PFA for 10 min. Cells were permeablised in blocking buffer (PBS with 3% BSA and 0.5% Triton-100x) for 10 min then stained with primary anti-Calcium Channel L-type alpha 1C subunit (Cacna1C) antibody conjugated with Atto 594 (ACC-003-AG, Alomone) at 1:200 dilution for 16 h (Raifman et al. 2017). To visualize nuclei cells were stained with Hoechst 33342 (8 µM for 30 min). Positive controls were initially performed with pancreatic beta-cell that express CaV1.2 (Schulla et al. 2003). Images were captured on a Zeiss LSM880C confocal microscope with excitation wavelengths of 405nm for Hoechst33342 and 633 nm for the anti-CaV1.2-ATTO antibody.
Measurement of [Ca2+]i
Adipocytes attached to coverslips were incubated with the Ca2+ fluorophore Fluo-4 AM (1 μM; Molecular Probes) in Hank’s solution with 0.01% BSA for 30 min at 21–23°C in the dark. Coverslips were mounted in a perifusion chamber on an Axiovert 135 Inverted microscope equipped for epifluorescence (Carl Zeiss Ltd). Cells were focused to maximize equatorial circumference and fluorescence. Adipocytes were identified under Kohler illumination as 50–100 μm diameter spheroids with a nuclear protuberance (Fig. 3). Experiments were performed 1–4 h post isolation.
Fluo-4 was excited at 450–490 nm, the emitted light band-pass filtered at 515–565 nm and the signal detected using a Photonics Science ISIS-3 camera with image intensification (luminous gain 8000:1). Fluorescence emission was integrated for 900 ms and captured at 1 Hz with an 8-bit frame grabber (DT3155, Data Translation, Basingstoke, UK) and Imaging Workbench, ver. 6.0 software (Indec Biosystems, Santa Clara, CA, USA). Cells were perifused at 1 mL/min. Since dye extrusion occurred at temperatures >30°C, experiments were performed at 27–28°C.
For data analysis, a region of interest (ROI) was drawn around each cell, background corrected and the time course of its mean fluorescent intensity was calculated. Fluorescence was calibrated by a two-point method (Ni et al. 1994): the maximum fluorescence value, Fmax; determined by permeabilization of the cells with Triton X-100 (0.0125–0.1%) followed by 10 mM EGTA to determine the minimum, Fmin. [Ca2+]i was calculated with the equation:
where F is the background corrected fluorescence, and K d the dissociation constant of Fluo-4: 345 nM (Bentley et al. 2014). Over 75% of basal [Ca2+]i values measured were below the K d of the dye and thus within its linear region of sensitivity. Data were only used from cells in which [Ca2+]i was stable and had a basal value within the 5–95% percentile range (60–380 nM).
To visualize spatial variation of [Ca2+]i it is necessary to mitigate fluorescent signal heterogeneity due to uneven dye loading or/and differences in columnar volume. In the absence of extracellular Ca2+, Ca2+ influx is abolished and heterogeneity in fluorescence was assumed to reflect variation in only cell volume and dye loading. Since these parameters proportionally affect Fluo-4 fluorescence, a ratio of the fluorescence signal in the presence of extracellular Ca2+ with that in its absence was undertaken using ImageJ2 with floating point arithmetic (Rueden et al. 2017) to normalize these confounders to reveal true differences in spatial [Ca2+]i.
Biochemical assays
To control for differences in adipocyte cell-density, biochemical data were normalized to paired values measured under basal or control conditions. Experiments were performed at 37°C.
Free fatty acid assay
Free fatty acid (FFA) release was used to measure lipolysis. Adipocytes were incubated for 60 min under different experimental conditions in 1 mL of Hank’s. After which, 100 μL of supranatant was removed and stored at −20°C prior to assay. FFA was assayed with a non-esterified FFA assay kit (WAKO chemicals). After subtraction of blank, data were normalized to the absorbance of the FFA standard (17 μM oleic acid) and corrected for dilution.
Glucose uptake
Glucose utilization was measured via 14C glucose uptake. After a 30-min pre-incubation under different experimental conditions, 14C glucose was added and the cells were incubated for a further 30 min, final volume 0.5 mL. For the assay, cells were separated from the medium by centrifugation and their 14C content measured via scintillation counting. The GLUT-dependent glucose uptake was determined by subtraction of non-specific measured in the presence of 10 µM Cytochalasin B. For analysis, 14C uptake was normalized to that measured in the absence of insulin and drug additions under control conditions.
Lipogenesis
Lipogenesis was measured via 3-3H-glucose incorporation. Adipocytes were incubated for 30 min under different experimental conditions with a fixed activity of 3-3H-glucose. For the assay, cells were separated from the medium by centrifugation and the 3H content determined via scintillation counting. Lipogenesis was determined as the difference of cellular counts to that measured in the absence of 3-3H-glucose. For analysis, lipogenesis was normalized to that measured in the absence of insulin and drug additions under control conditions.
Justification of drug concentrations employed
Verapamil and nifedipine were used at concentrations employed by others to block L-type VGCCs in this cell type (Martin et al. 1975, Begum et al. 1992, Ni et al. 1994). Moreover, concentrations were employed to mitigate their adsorption by serum (>90%) (Rumiantsev et al. 1989) and absorption by the adipocytes (Louis et al. 2014).
Statistical analysis
Data were checked with the D’Agostino and Pearson omnibus normality test with the appropriate inferential test given in the text. Graphical data are shown as box and whisker plots as median, interquartile range, 10, and 90% confidence intervals. Unless otherwise stated, experiments were performed as repeated measures. Numerical data are quoted as means ± s.e.m. or median with 5−95% confidence intervals (95% C.I.), where n is the number of determinations. Experimental data were collated from at least four animals. Fitting of equations to data used a least squares algorithm with the parameters given in text. Statistical analysis was performed using Graphpad PRISM, version 8.2. Data were considered statistically significant difference when P < 0.05 and in graphics is flagged as *, ** when P < 0.01, *** when P < 0.001 and **** when P < 0.0001.
Results
Molecular evidence and identification of VGCC in WAT
RT-PCR indicated the presence of mRNA for CaV1.1, CaV1.2 and CaV1.3 but not CaV1.4 L-type alpha1 subunits in epididymal WFA (Fig. 1A); mRNA for L-type VGCCs beta2 subunits: Cacnb2, Cacnb3 and Cacnb4, and alpha2delta subunits, Cacna2d1, Cacna2d2 and Cacna2d3 were also detected. cDNA sequence analysis of the PCR products for CaV1.1, CaV1.2 and CaV1.3 gave 98.7 ± 0.3 (n = 3), 98.5 ± 0.2% (n = 6) and 97.8 ± 0.7% (n = 6) identity to the rat VGCC subunits alpha1S (Cacna1s), alpha1C (Cacna1c) and alpha1D (Cacna1d) respectively. Quantification of mRNA levels by qPCR gave a rank expression order of CaV1.2 > CaV1.3 > CaV1.1 (Fig. 1B).
CaV1.1, 1.2 and 1.3 mRNA are expressed in rat adipocytes. (A) RT-CPR products (Rattus norvegicus) of voltage-dependent L-type calcium channels CaV1.1, CaV1.2, CaV1.3 and CaV1.4 alpha1 (Cacna1s, Cacna1c, Cacna1d and Cacna1f), beta2 (Cacnb1, Cacnb2, Cacnb3 and Cacnb4) and alpha2delta (Cacna2d1, Cacna2d2, Cacna2d3 and Cacna2d4) subunits for white fat adipocytes and for control: skeletal muscle for Cacna1s (CaV1.1) and whole brain for all other genes; L, DNA 50 bp ladder. PCR product sizes are in Table 1. (B) Relative expression of CaV1.1, CaV1.2 and CaV1.3 alpha-1 subunits as determined by qPCR. Data are normalized to mRNA expression of CaV1.2 in rat brain. Each point represents a different animal (n = 10), horizontal line is mean. Statistical significance is by one-way ANOVA, with Tukey’s multiple comparison test.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
CaV1.1, 1.2 and 1.3 mRNA are expressed in rat adipocytes. (A) RT-CPR products (Rattus norvegicus) of voltage-dependent L-type calcium channels CaV1.1, CaV1.2, CaV1.3 and CaV1.4 alpha1 (Cacna1s, Cacna1c, Cacna1d and Cacna1f), beta2 (Cacnb1, Cacnb2, Cacnb3 and Cacnb4) and alpha2delta (Cacna2d1, Cacna2d2, Cacna2d3 and Cacna2d4) subunits for white fat adipocytes and for control: skeletal muscle for Cacna1s (CaV1.1) and whole brain for all other genes; L, DNA 50 bp ladder. PCR product sizes are in Table 1. (B) Relative expression of CaV1.1, CaV1.2 and CaV1.3 alpha-1 subunits as determined by qPCR. Data are normalized to mRNA expression of CaV1.2 in rat brain. Each point represents a different animal (n = 10), horizontal line is mean. Statistical significance is by one-way ANOVA, with Tukey’s multiple comparison test.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
CaV1.1, 1.2 and 1.3 mRNA are expressed in rat adipocytes. (A) RT-CPR products (Rattus norvegicus) of voltage-dependent L-type calcium channels CaV1.1, CaV1.2, CaV1.3 and CaV1.4 alpha1 (Cacna1s, Cacna1c, Cacna1d and Cacna1f), beta2 (Cacnb1, Cacnb2, Cacnb3 and Cacnb4) and alpha2delta (Cacna2d1, Cacna2d2, Cacna2d3 and Cacna2d4) subunits for white fat adipocytes and for control: skeletal muscle for Cacna1s (CaV1.1) and whole brain for all other genes; L, DNA 50 bp ladder. PCR product sizes are in Table 1. (B) Relative expression of CaV1.1, CaV1.2 and CaV1.3 alpha-1 subunits as determined by qPCR. Data are normalized to mRNA expression of CaV1.2 in rat brain. Each point represents a different animal (n = 10), horizontal line is mean. Statistical significance is by one-way ANOVA, with Tukey’s multiple comparison test.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Western blots of the plasma membrane fraction revealed a 260 kDa CaV1.3 alpha-subunit (Fig. 2B) (N’Gouemo et al. 2015) and an extended form of CaV1.2 of >250 kDa (Fig. 2A); the latter bigger than the canonical neuronal isoform of 210 kDa (Raifman et al. 2017). CaV1.2 protein expression was 10-fold greater than that of CaV1.3 (Fig. 2C; P < 0.001, Mann−Whitney); a ratio comparable to the qPCR results (Fig. 1B).
White fat adipocytes express CaV1.2 and CaV1.3 protein in membrane fractions. Representative Western blots of CaV1.2 (A) and CaV1.3 (B) in adipocyte cell lysate fraction (ACF) and membrane fraction (AM) of white fat adipocytes. Note the larger molecular weight (MW) of the CaV1.2 protein in adipocytes: >250 kDa compared to the proteolytic cleavage product of 210 kDa in the whole brain cell lysate (BCL) positive control. (C) Relative protein expression of CaV1.2 (n = 5) to CaV1.3 (n = 9) in membrane fractions from white fat adipocytes. Data normalized to β-actin. Horizontal lines are the medians. Statistical significance is by Mann−Whitney.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
White fat adipocytes express CaV1.2 and CaV1.3 protein in membrane fractions. Representative Western blots of CaV1.2 (A) and CaV1.3 (B) in adipocyte cell lysate fraction (ACF) and membrane fraction (AM) of white fat adipocytes. Note the larger molecular weight (MW) of the CaV1.2 protein in adipocytes: >250 kDa compared to the proteolytic cleavage product of 210 kDa in the whole brain cell lysate (BCL) positive control. (C) Relative protein expression of CaV1.2 (n = 5) to CaV1.3 (n = 9) in membrane fractions from white fat adipocytes. Data normalized to β-actin. Horizontal lines are the medians. Statistical significance is by Mann−Whitney.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
White fat adipocytes express CaV1.2 and CaV1.3 protein in membrane fractions. Representative Western blots of CaV1.2 (A) and CaV1.3 (B) in adipocyte cell lysate fraction (ACF) and membrane fraction (AM) of white fat adipocytes. Note the larger molecular weight (MW) of the CaV1.2 protein in adipocytes: >250 kDa compared to the proteolytic cleavage product of 210 kDa in the whole brain cell lysate (BCL) positive control. (C) Relative protein expression of CaV1.2 (n = 5) to CaV1.3 (n = 9) in membrane fractions from white fat adipocytes. Data normalized to β-actin. Horizontal lines are the medians. Statistical significance is by Mann−Whitney.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Immunohistochemical evidence of VGCC in WAT
Confocal microscopy revealed that fixed adipocytes preserved their morphology (Fig. 3A) as demonstrated by retention of their nucleus and spherical form. Atto 594-labelled antibodies identified CaV1.2 in the plasma membrane; this was granular in appearance and densest near the nucleus (Fig. 3B). Given this finding, the spatial distribution of [Ca2+]i was examined (Fig. 3C). Figure 3C shows six adipocytes with identifiable nuclei, all of which responded with a reversible decrease in [Ca2+]i on removal of bath Ca2+ (Fig. 5A). Although the change in [Ca2+]i was uniformly distributed in the extra-nuclear cell membrane for all six cells, three had a higher [Ca2+]i in the perinuclear region (Fig. 3D); data which support a higher density of VGCCs in this region.
CaV1.2 in the plasma membrane. (A and B) Confocal images of fixed rat epididymal white fat adipocytes. (A) Blue, nucleus stained with Hoechst 33342; Magenta, Atto 594-labelled antibody to CaV1.2. Image captured over 62 s. (B) Magenta only channel to highlight CaV1.2 labelling which appears densest in the nuclear region. Arrows indicate associated nuclei. (C and D) Greyscale epifluorescent Ca2+ images of a field of six rat epididymal white fat adipocytes. (C) Adipocytes under basal conditions, arrows indicate nuclei. Note nuclear protuberances and brighter circumferential fluorescence where the cytoplasm has the deepest volume parallel to the plane of illumination. (D) Image shown in C is ratioed to that observed in the absence of extracellular Ca2+ to normalize dye loading and columnar volume. The brighter fluorescence in the perinuclear region of cells i, ii, and iii indicates a higher Ca2+ level. (C) and (D) are averages of 100 frames.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
CaV1.2 in the plasma membrane. (A and B) Confocal images of fixed rat epididymal white fat adipocytes. (A) Blue, nucleus stained with Hoechst 33342; Magenta, Atto 594-labelled antibody to CaV1.2. Image captured over 62 s. (B) Magenta only channel to highlight CaV1.2 labelling which appears densest in the nuclear region. Arrows indicate associated nuclei. (C and D) Greyscale epifluorescent Ca2+ images of a field of six rat epididymal white fat adipocytes. (C) Adipocytes under basal conditions, arrows indicate nuclei. Note nuclear protuberances and brighter circumferential fluorescence where the cytoplasm has the deepest volume parallel to the plane of illumination. (D) Image shown in C is ratioed to that observed in the absence of extracellular Ca2+ to normalize dye loading and columnar volume. The brighter fluorescence in the perinuclear region of cells i, ii, and iii indicates a higher Ca2+ level. (C) and (D) are averages of 100 frames.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
CaV1.2 in the plasma membrane. (A and B) Confocal images of fixed rat epididymal white fat adipocytes. (A) Blue, nucleus stained with Hoechst 33342; Magenta, Atto 594-labelled antibody to CaV1.2. Image captured over 62 s. (B) Magenta only channel to highlight CaV1.2 labelling which appears densest in the nuclear region. Arrows indicate associated nuclei. (C and D) Greyscale epifluorescent Ca2+ images of a field of six rat epididymal white fat adipocytes. (C) Adipocytes under basal conditions, arrows indicate nuclei. Note nuclear protuberances and brighter circumferential fluorescence where the cytoplasm has the deepest volume parallel to the plane of illumination. (D) Image shown in C is ratioed to that observed in the absence of extracellular Ca2+ to normalize dye loading and columnar volume. The brighter fluorescence in the perinuclear region of cells i, ii, and iii indicates a higher Ca2+ level. (C) and (D) are averages of 100 frames.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Evidence for constitutive Ca2+ influx
Epifluorescent imaging of adherent adipocytes revealed a skewed basal [Ca2+]i and Gaussian cell diameter distributions with median values of 135 nM (129−145 nM, 95% C.I., n = 555) and 80 µm (79−81 µm, 95% C.I., n = 580) respectively (Fig. 4A and B). [Ca2+]i values are similar to those previously reported (Schwartz et al. 1991, Hardy et al. 1992, Ni et al. 1994, Gaur et al. 1998).
Measurement of intracellular [Ca2+]i in isolated adipocytes. (A) Distribution of basal [Ca2+]i (n = 588). (B) Distribution of adipocyte diameters (n = 547). (C) Scatter plot of [Ca2+]i versus cell diameter (n = 495). (D) Scatter plot of cell diameter for 233 adipocytes with each vertical data set taken from a given weighed animal (45 in total). Solid lines in (C) and (D) are drawn by linear regression with slopes of 0.65 ± 0.29 nM/µM (P < 0.03) and 0.058 ± 0.02 µm/g (P < 0.01) respectively. Dotted lines are 95% C.I. for the fits shown. (E) Individual distributions of basal [Ca2+]i for epididymal adipocytes from 13 different rats (n = 7−40). Note the variation in [Ca2+]i within any given animal was greater than between animals (ANOVA).
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Measurement of intracellular [Ca2+]i in isolated adipocytes. (A) Distribution of basal [Ca2+]i (n = 588). (B) Distribution of adipocyte diameters (n = 547). (C) Scatter plot of [Ca2+]i versus cell diameter (n = 495). (D) Scatter plot of cell diameter for 233 adipocytes with each vertical data set taken from a given weighed animal (45 in total). Solid lines in (C) and (D) are drawn by linear regression with slopes of 0.65 ± 0.29 nM/µM (P < 0.03) and 0.058 ± 0.02 µm/g (P < 0.01) respectively. Dotted lines are 95% C.I. for the fits shown. (E) Individual distributions of basal [Ca2+]i for epididymal adipocytes from 13 different rats (n = 7−40). Note the variation in [Ca2+]i within any given animal was greater than between animals (ANOVA).
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Measurement of intracellular [Ca2+]i in isolated adipocytes. (A) Distribution of basal [Ca2+]i (n = 588). (B) Distribution of adipocyte diameters (n = 547). (C) Scatter plot of [Ca2+]i versus cell diameter (n = 495). (D) Scatter plot of cell diameter for 233 adipocytes with each vertical data set taken from a given weighed animal (45 in total). Solid lines in (C) and (D) are drawn by linear regression with slopes of 0.65 ± 0.29 nM/µM (P < 0.03) and 0.058 ± 0.02 µm/g (P < 0.01) respectively. Dotted lines are 95% C.I. for the fits shown. (E) Individual distributions of basal [Ca2+]i for epididymal adipocytes from 13 different rats (n = 7−40). Note the variation in [Ca2+]i within any given animal was greater than between animals (ANOVA).
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Both basal [Ca2+]i (Spearman r = 0.17, 0.084–0.25 95% C.I., P < 0.0001, Fig. 4C) and body weight (Spearman r = 0.13, 0.01–0.26 95% C.I., P < 0.05, Fig. 4D) were positively correlated with adipocyte diameter, although [Ca2+]i did not correlate with individual animal weight (Spearman, Fig. 4E).
To explore Ca2+ influx, the effect of equimolar substitution of extracellular Ca2+ with Mg2+ was investigated. Removal of bath Ca2+ reversibly decreased [Ca2+]i, Δ[Ca2+]i, by 30% (25–34%, 95% C.I.; P < 0.0001, Friedman; Fig. 5A and B). Δ[Ca2+]i was negatively correlated with basal [Ca2+]i: that is cells with the highest basal values underwent the largest percentage decrease on Ca2+ removal (Spearman r = −0.32, P < 0.001; Fig. 5C). Δ[Ca2+]i, was unrelated to cell diameter or body weight (Spearman). Prolonged exposure to Ca2+ removal for over 3 h did not affect cell integrity which suggest that this intervention is not cytotoxic.
Extracellular Ca2+ removal decreases intracellular [Ca2+]i. (A) [Ca2+]i time courses measured for six adipocytes within a single field in response to removal of bath Ca2+ (Ca-free), followed by 0.1% DMSO. (B) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free) and recovery (Wash) (n = 138). Statistical inference by Friedman with Dunn’s multiple comparison tests. (C) Relationship between the decrease in [Ca2+]i, Δ[Ca2+]i, on removal of bath Ca2+and basal [Ca2+]i. Solid line drawn by linear regression with a slope of −0.17 ± 0.01%/nM (P < 0.0001). Dotted lines are the 95% C.I. for the fit shown. (D) Mean time course of [Ca2+]i for five adipocytes in response to 5 mM CaCl2 (HiCa) followed by 2.5 mM CoCl2 (Cobalt) added to the bath. (E) Δ[Ca2+]i responses to HiCa (n = 17), Cobalt (n = 19) and removal of extracellular Ca2+ (Ca-free; n = 140). Dashed line indicates no effect. Data are from seven animals. Statistical inference was by Kruskal–Wallis with Dunn’s multiple comparison tests. A full colour version of this figure is available at https://doi.org/10.1530/JOE-19-0493.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Extracellular Ca2+ removal decreases intracellular [Ca2+]i. (A) [Ca2+]i time courses measured for six adipocytes within a single field in response to removal of bath Ca2+ (Ca-free), followed by 0.1% DMSO. (B) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free) and recovery (Wash) (n = 138). Statistical inference by Friedman with Dunn’s multiple comparison tests. (C) Relationship between the decrease in [Ca2+]i, Δ[Ca2+]i, on removal of bath Ca2+and basal [Ca2+]i. Solid line drawn by linear regression with a slope of −0.17 ± 0.01%/nM (P < 0.0001). Dotted lines are the 95% C.I. for the fit shown. (D) Mean time course of [Ca2+]i for five adipocytes in response to 5 mM CaCl2 (HiCa) followed by 2.5 mM CoCl2 (Cobalt) added to the bath. (E) Δ[Ca2+]i responses to HiCa (n = 17), Cobalt (n = 19) and removal of extracellular Ca2+ (Ca-free; n = 140). Dashed line indicates no effect. Data are from seven animals. Statistical inference was by Kruskal–Wallis with Dunn’s multiple comparison tests. A full colour version of this figure is available at https://doi.org/10.1530/JOE-19-0493.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Extracellular Ca2+ removal decreases intracellular [Ca2+]i. (A) [Ca2+]i time courses measured for six adipocytes within a single field in response to removal of bath Ca2+ (Ca-free), followed by 0.1% DMSO. (B) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free) and recovery (Wash) (n = 138). Statistical inference by Friedman with Dunn’s multiple comparison tests. (C) Relationship between the decrease in [Ca2+]i, Δ[Ca2+]i, on removal of bath Ca2+and basal [Ca2+]i. Solid line drawn by linear regression with a slope of −0.17 ± 0.01%/nM (P < 0.0001). Dotted lines are the 95% C.I. for the fit shown. (D) Mean time course of [Ca2+]i for five adipocytes in response to 5 mM CaCl2 (HiCa) followed by 2.5 mM CoCl2 (Cobalt) added to the bath. (E) Δ[Ca2+]i responses to HiCa (n = 17), Cobalt (n = 19) and removal of extracellular Ca2+ (Ca-free; n = 140). Dashed line indicates no effect. Data are from seven animals. Statistical inference was by Kruskal–Wallis with Dunn’s multiple comparison tests. A full colour version of this figure is available at https://doi.org/10.1530/JOE-19-0493.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Doubling the extracellular Ca2+ concentration produced a 9% increase in [Ca2+]i (4–16%, 95% C.I.; P < 0.0001 Wilcoxon signed-rank test; Fig. 5D and E). Conversely, addition of 2.5 mM Co2+, an inorganic inhibitor of VGCCs, irreversibly decreased [Ca2+]i by 13% (8–17%, 95% C.I.; P < 0.0001 Wilcoxon signed-rank; Fig. 5D); a magnitude comparable to that seen with removal of extracellular Ca2+ (Fig. 5E).
Pharmacological manipulation of Ca2+ influx
We next tested a variety of pharmacological agents that affect VGCC activity. Although both verapamil and nifedipine significantly decreased [Ca2+]i relative to DMSO control (Fig. 6) (P < 0.001 and P < 0.0001 respectively, Wilcoxon signed-rank test), the effect of nifedipine was small (Fig. 6B and F). Verapamil decreased Δ[Ca2+]i to a similar extent to Ca2+ removal (Kruskal–Wallis, Fig. 6F). In the presence of nifedipine Ca2+ removal further decreased [Ca2+]i (Fig. 6B and G). In any given cell, the amount by which verapamil decreased [Ca2+]i positively correlated with that seen with extracellular Ca2+ removal (Pearson r = 0.77, slope 0.66 ± 0.13, P < 0.001; Fig. 6H).
Verapamil and nifedipine decrease the intracellular Ca2+ concentration, [Ca2+]I. (A) Mean time course of [Ca2+]i measured for 11 adipocytes within a single field in response to removal of bath Ca2+ (Caf), followed by 20 µM verapamil (V). (B) Mean time course of [Ca2+]i measured for a field of five adipocytes in response to removal of bath Ca2+ (Caf) in the absence and then presence of 20 µM nifedipine (Nif). Note block by nifedipine is countered by a positive DMSO effect on fluorescence. (C) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free) and recovery (Wash) followed by 20 µM verapamil (Ver) (n = 19 from six animals). (D) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free) and recovery (Wash) followed by the addition of 20 µM nifedipine (Nif) (n = 15 from four animals). (E) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free) and recovery (Wash) followed by 0.1% DMSO (n = 36 from six animals). (F) Comparison of Δ[Ca2+]i produced by removal of bath Ca2+ (Ca-free), 20 µM nifedipine (Nif; n = 15) and 20 µM verapamil (Ver; n = 19). Data are from C and D and are corrected for DMSO effects shown in E (n = 137). (G) [Ca2+]i in the continuous presence of 20 µM nifedipine (Nif1) where Ca2+ is removed from the bath (Nif Ca-free) and added back again (Nif2) (n = 9 from two animals). For C, D, E, F and G, statistical comparison between groups was by Kruskal–Wallis with Dunn’s multiple comparison tests. (H) Relationship between Δ[Ca2+]i produced by 20 µM verapamil and removal of bath Ca2+ (n = 21). Solid line is linear regression of data with a slope of 0.66 ± 0.13. The dotted lines are the 95% C.I. of the fit.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Verapamil and nifedipine decrease the intracellular Ca2+ concentration, [Ca2+]I. (A) Mean time course of [Ca2+]i measured for 11 adipocytes within a single field in response to removal of bath Ca2+ (Caf), followed by 20 µM verapamil (V). (B) Mean time course of [Ca2+]i measured for a field of five adipocytes in response to removal of bath Ca2+ (Caf) in the absence and then presence of 20 µM nifedipine (Nif). Note block by nifedipine is countered by a positive DMSO effect on fluorescence. (C) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free) and recovery (Wash) followed by 20 µM verapamil (Ver) (n = 19 from six animals). (D) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free) and recovery (Wash) followed by the addition of 20 µM nifedipine (Nif) (n = 15 from four animals). (E) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free) and recovery (Wash) followed by 0.1% DMSO (n = 36 from six animals). (F) Comparison of Δ[Ca2+]i produced by removal of bath Ca2+ (Ca-free), 20 µM nifedipine (Nif; n = 15) and 20 µM verapamil (Ver; n = 19). Data are from C and D and are corrected for DMSO effects shown in E (n = 137). (G) [Ca2+]i in the continuous presence of 20 µM nifedipine (Nif1) where Ca2+ is removed from the bath (Nif Ca-free) and added back again (Nif2) (n = 9 from two animals). For C, D, E, F and G, statistical comparison between groups was by Kruskal–Wallis with Dunn’s multiple comparison tests. (H) Relationship between Δ[Ca2+]i produced by 20 µM verapamil and removal of bath Ca2+ (n = 21). Solid line is linear regression of data with a slope of 0.66 ± 0.13. The dotted lines are the 95% C.I. of the fit.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Verapamil and nifedipine decrease the intracellular Ca2+ concentration, [Ca2+]I. (A) Mean time course of [Ca2+]i measured for 11 adipocytes within a single field in response to removal of bath Ca2+ (Caf), followed by 20 µM verapamil (V). (B) Mean time course of [Ca2+]i measured for a field of five adipocytes in response to removal of bath Ca2+ (Caf) in the absence and then presence of 20 µM nifedipine (Nif). Note block by nifedipine is countered by a positive DMSO effect on fluorescence. (C) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free) and recovery (Wash) followed by 20 µM verapamil (Ver) (n = 19 from six animals). (D) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free) and recovery (Wash) followed by the addition of 20 µM nifedipine (Nif) (n = 15 from four animals). (E) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free) and recovery (Wash) followed by 0.1% DMSO (n = 36 from six animals). (F) Comparison of Δ[Ca2+]i produced by removal of bath Ca2+ (Ca-free), 20 µM nifedipine (Nif; n = 15) and 20 µM verapamil (Ver; n = 19). Data are from C and D and are corrected for DMSO effects shown in E (n = 137). (G) [Ca2+]i in the continuous presence of 20 µM nifedipine (Nif1) where Ca2+ is removed from the bath (Nif Ca-free) and added back again (Nif2) (n = 9 from two animals). For C, D, E, F and G, statistical comparison between groups was by Kruskal–Wallis with Dunn’s multiple comparison tests. (H) Relationship between Δ[Ca2+]i produced by 20 µM verapamil and removal of bath Ca2+ (n = 21). Solid line is linear regression of data with a slope of 0.66 ± 0.13. The dotted lines are the 95% C.I. of the fit.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
We next explored the pharmacology of [Ca2+]i recovery following re-addition of Ca2+ to the bath after its removal. Both verapamil (Fig. 7B, F and J) and nifedipine (Fig. 7C, G and J) significantly affected [Ca2+]i recovery (Fig. 7A, E and J). The NCX inhibitor SN-6 neither affected basal [Ca2+]i nor its recovery (Fig. 7H and J). BAY-K8644, an agonist of L-type VGCCs, significantly enhanced recovery (Fig. 7D, I and J).
Pharmacological exploration of Ca2+ recovery. (A, B, C and D) Representative mean time courses of [Ca2+]i in response to removal of bath Ca2+ (Ca-free) followed by the addition of drugs as indicated. (A) 0.1% vovl/vol DMSO; (B) 20 µM verapamil (V); (C) 20 µM nifedipine (Nif); (D) 10 µM BAY-K8644 (BAYK) all in Ca2+ free. (E) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), after addition of 0.1% DMSO (Ca-fDMSO), and then re-addition of bath Ca2+ in the presence of DMSO (n = 19 from 12 animals). (F) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), followed by 20 µM verapamil in Ca2+ free (Ca-fVer), then re-addition of bath Ca2+ in verapamil (Ver) (n = 67 from 12 animals). (G) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), followed by 20 µM nifedipine in Ca2+ free (Ca-fNif), then re-addition of bath Ca2+ in nifedipine (Nif) (n = 33 from six animals). (H) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), followed by 10 µM SN6 in Ca2+ free (Ca-fSN6), then re-addition of bath Ca2+ in SN6 (SN6) (n = 18 from five animals). (I) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), followed by 10 µM BAY-K8644 in Ca2+ free (Ca-fBAYK), and then re-addition of bath Ca2+ in BAY-K8644 (BAYK) (n = 18 from eight animals). All data uncorrected for DMSO effect. (J) Percentage change in basal [Ca2+]i after recovery on re-addition of bath Ca2+ with the various treatments as shown. Statistical comparison was by Friedman with Dunn’s multiple comparison multiple comparison.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Pharmacological exploration of Ca2+ recovery. (A, B, C and D) Representative mean time courses of [Ca2+]i in response to removal of bath Ca2+ (Ca-free) followed by the addition of drugs as indicated. (A) 0.1% vovl/vol DMSO; (B) 20 µM verapamil (V); (C) 20 µM nifedipine (Nif); (D) 10 µM BAY-K8644 (BAYK) all in Ca2+ free. (E) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), after addition of 0.1% DMSO (Ca-fDMSO), and then re-addition of bath Ca2+ in the presence of DMSO (n = 19 from 12 animals). (F) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), followed by 20 µM verapamil in Ca2+ free (Ca-fVer), then re-addition of bath Ca2+ in verapamil (Ver) (n = 67 from 12 animals). (G) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), followed by 20 µM nifedipine in Ca2+ free (Ca-fNif), then re-addition of bath Ca2+ in nifedipine (Nif) (n = 33 from six animals). (H) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), followed by 10 µM SN6 in Ca2+ free (Ca-fSN6), then re-addition of bath Ca2+ in SN6 (SN6) (n = 18 from five animals). (I) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), followed by 10 µM BAY-K8644 in Ca2+ free (Ca-fBAYK), and then re-addition of bath Ca2+ in BAY-K8644 (BAYK) (n = 18 from eight animals). All data uncorrected for DMSO effect. (J) Percentage change in basal [Ca2+]i after recovery on re-addition of bath Ca2+ with the various treatments as shown. Statistical comparison was by Friedman with Dunn’s multiple comparison multiple comparison.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Pharmacological exploration of Ca2+ recovery. (A, B, C and D) Representative mean time courses of [Ca2+]i in response to removal of bath Ca2+ (Ca-free) followed by the addition of drugs as indicated. (A) 0.1% vovl/vol DMSO; (B) 20 µM verapamil (V); (C) 20 µM nifedipine (Nif); (D) 10 µM BAY-K8644 (BAYK) all in Ca2+ free. (E) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), after addition of 0.1% DMSO (Ca-fDMSO), and then re-addition of bath Ca2+ in the presence of DMSO (n = 19 from 12 animals). (F) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), followed by 20 µM verapamil in Ca2+ free (Ca-fVer), then re-addition of bath Ca2+ in verapamil (Ver) (n = 67 from 12 animals). (G) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), followed by 20 µM nifedipine in Ca2+ free (Ca-fNif), then re-addition of bath Ca2+ in nifedipine (Nif) (n = 33 from six animals). (H) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), followed by 10 µM SN6 in Ca2+ free (Ca-fSN6), then re-addition of bath Ca2+ in SN6 (SN6) (n = 18 from five animals). (I) [Ca2+]i in control (Ctrl), after removal of extracellular Ca2+ (Ca-free), followed by 10 µM BAY-K8644 in Ca2+ free (Ca-fBAYK), and then re-addition of bath Ca2+ in BAY-K8644 (BAYK) (n = 18 from eight animals). All data uncorrected for DMSO effect. (J) Percentage change in basal [Ca2+]i after recovery on re-addition of bath Ca2+ with the various treatments as shown. Statistical comparison was by Friedman with Dunn’s multiple comparison multiple comparison.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
L-type Ca2+ influx stimulates basal lipolysis
Since Ca2+ modulates the lipolytic cascade in WFA (Schimmel 1978, Izawa et al. 1983, Allen & Beck 1986), we investigated if L-type VGCCs affected lipolysis (Fig. 9). Isoprenaline stimulated lipolysis with a pEC50 of 7 (6.7–7.3, 95% C.I.) and Hill coefficient of 0.8 (0.43–1.2, 95% C.I.; Fig. 9A). At 10 µM, isoprenaline stimulated lipolysis 9.8 ± 0.63 fold (P < 0.001, one sample t test; n = 39). Consistent with previous reports (Izawa et al. 1983, Allen & Beck 1986), removal of extracellular Ca2+ neither affected the EC50 nor Hill coefficient for isoprenaline, but decreased lipolysis by ~30% (P < 0.0004; Fig. 8A and C). Insulin inhibited the beta-adrenoceptor-stimulated lipolysis with a pEC50 of 9.4 (9.6–9.3, 95% C.I.; Fig. 9B).
Extracellular Ca2+-influx potentiates lipolysis. (A) Lipolysis as a function of isoprenaline in the presence (black circles) and absence (open circles) of bath Ca2+. Solid lines are fits of the data with sigmoidal dose–response curves with parameters given in the text. Data means ± s.e.m. (n = 4–5). (B) Inhibition of lipolysis by insulin. Solid line is a fit of the data to dose–response curve with parameters given in the text. Data are means ± s.e.m. (n = 5–9). (C) Effects of interventions on lipolysis stimulated by 10 µM isoprenaline: Ca free, removal of bath Ca2+; Ver, 5 µM verapamil; Nif, 20 µM nifedipine; Ins, 20 nM insulin; BAYK, 1 µM BAY-K8644; HiK, 50 mM bath [K+]o (n = 6–16). (D) Effects of interventions on basal lipolysis. Key as for C (n = 6–16). (E) Effects of interventions on beta-adrenoceptor-mediated lipolysis inhibited by 20 nM insulin: Key as for C (n = 6–10). (F) Effects of interventions on 10 µM isoprenaline-stimulated lipolysis: OXY, 1 µM oxytocin; THAP, 10 µM thapsigargin; BAPTA, 10 µM BAPTA-AM (n = 9–21). (G) Effects of interventions as shown in F on basal lipolysis (n = 6–10). DMSO the solvent for verapamil, nifedipine, and BAY-K8644 was without effect on lipolysis. Statistics are Wilcoxon sign test relative to 100%.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Extracellular Ca2+-influx potentiates lipolysis. (A) Lipolysis as a function of isoprenaline in the presence (black circles) and absence (open circles) of bath Ca2+. Solid lines are fits of the data with sigmoidal dose–response curves with parameters given in the text. Data means ± s.e.m. (n = 4–5). (B) Inhibition of lipolysis by insulin. Solid line is a fit of the data to dose–response curve with parameters given in the text. Data are means ± s.e.m. (n = 5–9). (C) Effects of interventions on lipolysis stimulated by 10 µM isoprenaline: Ca free, removal of bath Ca2+; Ver, 5 µM verapamil; Nif, 20 µM nifedipine; Ins, 20 nM insulin; BAYK, 1 µM BAY-K8644; HiK, 50 mM bath [K+]o (n = 6–16). (D) Effects of interventions on basal lipolysis. Key as for C (n = 6–16). (E) Effects of interventions on beta-adrenoceptor-mediated lipolysis inhibited by 20 nM insulin: Key as for C (n = 6–10). (F) Effects of interventions on 10 µM isoprenaline-stimulated lipolysis: OXY, 1 µM oxytocin; THAP, 10 µM thapsigargin; BAPTA, 10 µM BAPTA-AM (n = 9–21). (G) Effects of interventions as shown in F on basal lipolysis (n = 6–10). DMSO the solvent for verapamil, nifedipine, and BAY-K8644 was without effect on lipolysis. Statistics are Wilcoxon sign test relative to 100%.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Extracellular Ca2+-influx potentiates lipolysis. (A) Lipolysis as a function of isoprenaline in the presence (black circles) and absence (open circles) of bath Ca2+. Solid lines are fits of the data with sigmoidal dose–response curves with parameters given in the text. Data means ± s.e.m. (n = 4–5). (B) Inhibition of lipolysis by insulin. Solid line is a fit of the data to dose–response curve with parameters given in the text. Data are means ± s.e.m. (n = 5–9). (C) Effects of interventions on lipolysis stimulated by 10 µM isoprenaline: Ca free, removal of bath Ca2+; Ver, 5 µM verapamil; Nif, 20 µM nifedipine; Ins, 20 nM insulin; BAYK, 1 µM BAY-K8644; HiK, 50 mM bath [K+]o (n = 6–16). (D) Effects of interventions on basal lipolysis. Key as for C (n = 6–16). (E) Effects of interventions on beta-adrenoceptor-mediated lipolysis inhibited by 20 nM insulin: Key as for C (n = 6–10). (F) Effects of interventions on 10 µM isoprenaline-stimulated lipolysis: OXY, 1 µM oxytocin; THAP, 10 µM thapsigargin; BAPTA, 10 µM BAPTA-AM (n = 9–21). (G) Effects of interventions as shown in F on basal lipolysis (n = 6–10). DMSO the solvent for verapamil, nifedipine, and BAY-K8644 was without effect on lipolysis. Statistics are Wilcoxon sign test relative to 100%.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Ca2+-influx via L–Type VGCC does not affect glucose uptake and lipogenesis. (A) Effect of 5 µM nifedipine on glucose uptake. (B) Effect of 1 µM BAY-K 8644 on glucose uptake. (C) Effect of 5 µM nifedipine on lipogenesis. (D) Effect of 1 µM BAY-K 8644 on lipogenesis. Drug additions are indicated by filled bars, controls by open bars, conditions are as indicated. Ins, insulin, glu, glucose. Data are all paired with n = 5 for each condition. Statistical inference by Friedman’s test.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Ca2+-influx via L–Type VGCC does not affect glucose uptake and lipogenesis. (A) Effect of 5 µM nifedipine on glucose uptake. (B) Effect of 1 µM BAY-K 8644 on glucose uptake. (C) Effect of 5 µM nifedipine on lipogenesis. (D) Effect of 1 µM BAY-K 8644 on lipogenesis. Drug additions are indicated by filled bars, controls by open bars, conditions are as indicated. Ins, insulin, glu, glucose. Data are all paired with n = 5 for each condition. Statistical inference by Friedman’s test.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Ca2+-influx via L–Type VGCC does not affect glucose uptake and lipogenesis. (A) Effect of 5 µM nifedipine on glucose uptake. (B) Effect of 1 µM BAY-K 8644 on glucose uptake. (C) Effect of 5 µM nifedipine on lipogenesis. (D) Effect of 1 µM BAY-K 8644 on lipogenesis. Drug additions are indicated by filled bars, controls by open bars, conditions are as indicated. Ins, insulin, glu, glucose. Data are all paired with n = 5 for each condition. Statistical inference by Friedman’s test.
Citation: Journal of Endocrinology 244, 2; 10.1530/JOE-19-0493
Interventions that promoted Ca2+ influx: BAY-K8644 or elevation of [K+]o (Bentley et al. 2014), stimulated basal, but not isoprenaline-stimulated, lipolysis (Fig. 8C and D). Interventions that decreased Ca2+ influx: removal of extracellular Ca2+ or verapamil inhibited both isoprenaline-stimulated and basal lipolysis (Fig. 8C and D). Insulin stimulated basal lipolysis (Fig. 8D), an effect consistent with its ability to elevate [Ca2+]i (Clausen & Martin 1977). The capacity of 20 nM insulin to block beta-adrenoceptor-stimulated lipolysis was unaffected by interventions that affected [Ca2+]i (Fig. 8E).
We checked if intracellular Ca2+ handling affected lipolysis. 1 µM oxytocin, which mobilizes intracellular Ca2+ (Kelly et al. 1989), did not affect basal lipolysis (Fig. 8G), but consistent with others (Fain et al. 1997) inhibited isoprenaline-stimulated lipolysis (Fig. 8F). Neither store depletion with 1 µM thapsigargin nor increased cytosolic Ca2+ buffering with 10 µM BAPTA-AM affected isoprenaline-stimulated lipolysis Fig. 8F); however, both treatments impaired basal lipolysis (Fig. 8G); actions consistent with the greater sensitivity of basal lipolysis to [Ca2+]i compared to that stimulated by isoprenaline (Fig. 8C and D).
Although alpha-adrenoceptor activation mobilizes intracellular Ca2+ stores in WFA (Hardy et al. 1992), like others (Blackmore & Augert 1989, Seydoux et al. 1996), we did not see an increase in [Ca2+]I with beta-adrenoceptor activation (n = 7).
L-type Ca2+ influx does not affect glucose uptake or lipogenesis
Insulin at 2 nM maximally stimulated glucose uptake fivefold (P < 0.001, one sample t test) (Fig. 9A). Insulin–stimulated glucose uptake was unaffected by either 5 µM nifedipine or 1 µM BAY-K8644 (Fig. 9A and B); outcomes that were independent of insulin concentration. Lipogenesis was also maximally stimulated by 2 nM insulin (~ninefold, P < 0.001, one sample t test) (Fig. 9C and D). However, neither 5 µM nifedipine nor 1 µM BAY-K8644 significantly affected lipogenesis (Fig. 9C and D).
Discussion
Molecular evidence for L-type Ca2+ channels
Our Western blot, PCR and in situ immunolabelling data demonstrate expression of CaV1.2 and CaV1.3 L-type VGCCs in primary white fat adipocytes. The presence of CaV1.2/CaV1.3 in WFA is consistent with transcriptomic data published for human (Fagerberg et al. 2014) and mouse fat tissue (Yue et al. 2014); however, in contrast to these, we failed to detect CaV1.4. Although the 250 kDa CaV1.2 protein band went undetected in our brain tissue control we did detect its 210 kD proteolytic cleavage product (Buonarati et al. 2017, Shi et al. 2017). Using the same antibody, others have also detected just a 210 kDa CaV1.2 band (N’Gouemo et al. 2015). The 140 kDa band may relate to the 130 kDa or 150 kDa non-CaV1.2 epitope that this antibody recognized in CaV1.2-knockout mice (Bavley et al. 2017, Buonarati et al. 2017). Though adipocytes possessed a CaV1.2 immuno-positive band, they did not show any further proteolytic cleavage products, data that suggest that this protein has post-translational modification with a polymorphic proteolytic cleavage site. For CaV1.3, we only obtained positive immunoblots with early batches of antibody, a recognized problem with commercial antibodies to CaV1.3 (Buonarati et al. 2017). In WFA, CaV1.3 was comparable in molecular weight to that observed in our brain controls and heart (268 kDa) (Le Scouarnec et al. 2008). The presence of mRNA for L-type VGCC alpha1, beta2 and alpha2delta subunits suggests that adipocytes have the capacity to traffic and assemble functional VGCCs (Dolphin 2016).
Constitutive Ca2+ influx through L-type Ca2+ channels
Our finding that both Co2+ and verapamil decreased [Ca2+]i by similar amounts to Ca2+ removal is indicative of a Ca2+-influx pathway mediated by L-type VGCCs. This notion is reinforced by enhancement of Ca2+ influx by the L-type VGCCs dihydropyridine agonist BAY-K8644. Our data contrast to studies where neither nitrendipine nor verapamil affected [Ca2+]i under basal conditions (Gaur et al. 1996a ,b ). One possible reason for this discrepancy is that we corrected for the effect of DMSO on fluorescence to reveal a block, whereas previous studies had not.
The fact that removal of bath Ca2+ had a larger effect in adipocytes with higher basal [Ca2+]i suggests that basal [Ca2+]i is set by the prevalent VGCC activity. These findings, combined with the ability of verapamil and nifedipine to prevent, and BAY-K8644 to enhance Ca2+ recovery, suggest that basal [Ca2+]i in WFA is maintained by a constitutive Ca2+ influx via L-type VGCCs – an idea supported by impairment of 45Ca2+ uptake in WFA by L-type VGCCs antagonists (Martin et al. 1975).
The resting membrane potential of primary white adipocytes, measured by ourselves (Bentley et al. 2014) and others (Ramírez-Ponce et al. 1990, Lee & Pappone 1997), is around −30 mV. As this voltage is within the activation range of L-type VGCCs (Xu & Lipscombe 2001) a ‘window Ca2+ current’ (Fleischmann et al. 1994) is expected. Constitutive L-type VGCC activity in electrically non-excitable cells is not unique; for example, it has been recorded at a similar Vm (−30 mV) in osteoclasts (Miyauchi et al. 1990). To be constitutively active, inactivation of these VGCCs must be incomplete.
Although whole-cell voltage-clamp would have been ideal for electrophysiological characterization of the VGCCs we did not attempt this for technical reasons. First, the adipocyte cytoplasm is a relatively thin ~0.3 µm layer wrapped around a lipid droplet of ~80 µm diameter (Bentley et al. 2014); this creates a membrane time constant of 100’s of ms (Bentley et al. 2014), compared to ~2 ms for a neuron (Coombs et al. 1956) of similar diameter (Henneman & Mendell 2010). This difference precludes voltage-clamping of VGCCs under physiological conditions due to space clamp considerations and ‘voltage escape’ (Armstrong & Gilly 1992). Secondly, although Ca2+ influx can affect [Ca2+]i, we have previously shown that it is too small to affect Vm (Bentley et al. 2014), a result indicative of low channel density. Indeed, 45Ca2+ tracer studies (Martin et al. 1975) have measured the DHP-sensitive Ca2+ influx in WFA at ~0.024 pmol/s/cm2: ~1.5 amol/s/cell or ~0.3 pA of whole-cell inward Ca2+ current, a value too small to measure with whole-cell voltage clamp.
Constitutive Ca2+ influx modulates lipid metabolism
Our data confirm that both basal and stimulated lipolysis requires extracellular Ca2+ (Bleicher et al. 1966, Ziegler et al. 1980, Allen & Beck 1986). However, we now show that lipolysis is sensitive to agents that modulate L-type VGCC activity. Although, isoprenaline-stimulated lipolysis could not be enhanced by interventions that increase [Ca2+]i, presumably because it was already maximal, like basal lipolysis, it was impaired by verapamil and Ca2+ removal. The idea that Ca2+-influx promotes lipolysis is also supported by the action of high K+, where substitution of bath Na+ with K+ elevates [Ca2+]i by reverse Na+-Ca2+ exchange (Bentley et al. 2014).
Multiple targets exist for extracellular Ca2+ during beta-adrenoceptor-stimulated lipolysis. Extracellular Ca2+ enhances beta-adrenoceptor-stimulated cAMP production (Ziegler et al. 1980) and is also required for undefined lipolytic processes downstream of cytosolic cAMP (Allen & Beck 1986). Indeed, lipolysis can be potentiated by increasing [Ca2+]i with Ca2+ ionophores in the absence of cAMP elevation (Gaion & Krishna 1982) or conversely, as shown here, depressed with Ca2+ chelators (Efendić et al. 1970). However, the exact mechanisms by which Ca2+, and indeed Ca2+ influx modulates basal lipolysis remains unknown; it is unlikely to involve the Ca2+-sensitive, isoform III, of adenylyl cyclase found in WFA (Wang et al. 2009) since changes in Ca2+ influx do not affect adipocyte cAMP levels under basal conditions (Ziegler et al. 1980). However, adipocyte lipoprotein lipase is positively modulated by Ca2+ (Efendić et al. 1970, Soma et al. 1989, Carmen & Víctor 2006).
Translational context
The idea that Ca2+ influx via CaV1.2/CaV1.3 regulates basal lipolysis agrees with translational data. Rats with chronic renal failure are hyperlipidaemic and possess WFA with elevated intracellular Ca2+ levels (Ni et al. 1995). The observation that verapamil reversed these phenomena suggests that exacerbated Ca2-influx via L-type VGCCs may be responsible for the elevated basal lipolysis. This notion is also supported by the ability of nicardipine to decrease plasma levels of FFA in spontaneously hypertensive rats (Cignarella 1994). Moreover, in humans, verapamil can also decrease basal plasma FFA levels, again data suggestive of VGCC-mediated Ca2-influx-dependent lipolysis (Hvarfner et al. 1988). Such effects do not arise through impairment of beta-adrenoceptor activation since antihypertensives such as nicardipine and verapamil actually promote catecholamine levels through activation of baroreflex-mediated sympathetic output. The widespread failure of the DHP L-type channel antagonists to affect plasma FFA in man can be reconciled by the binding of these drugs to CaV1.2/1.3 being compromised by FFA (Pepe et al. 1994).
Conclusion
Our study provides direct, corroborative evidence for the existence of CaV1.2/CaV1.3 L-type voltage-gated Ca2+ channels in white adipocytes. The basal concentration of intracellular Ca2+ appears to reflect the ambient level of VGCC activity on an individual cell basis, apparently unrelated to adipocyte size or animal weight. Importantly, CaV1.x channels contribute to a persistent state of Ca2+ influx in WFA, without the need for cell excitability, and appear to have a key role in lipolysis. Consequently, dysregulation of CaV1.x in WFA may contribute to lipid storage disorders that may contribute to, or indeed precipitate, the metabolic syndrome. As such, these cation channels may be a potential therapeutic target for the treatment of hyperlipidaemia, peripheral insulin resistance and obesity.
Declaration of interest
The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.
Funding
This work was supported by Diabetes UK (Grant ID: RB03CJ), Leverhulme Trust (Grant ID: RPG-2017-162), Swedish Diabetes Foundation (DIA2015-062) and Swedish Medical Research Council (Grant ID: 2013-7107).
Acknowledgements
P P was in receipt of the Royal Thai Government scholarship. D C B was in receipt of a BBSRC studentship. N A is in receipt of a Schlumberger Foundation PhD fellowship.
References
Allen DO & Beck RRR 1986 Role of calcium ion in hormone-stimulated lipolysis. Biochemical Pharmacology 767–772. (https://doi.org/10.1016/0006-2952(86)90244-3)
Armstrong CM & Gilly WF 1992 Access resistance and space clamp problems associated with whole-cell patch clamping. Methods in Enzymology 100–122. (https://doi.org/10.1016/0076-6879(92)07007-B)
Arner P, Bernard S, Salehpour M, Possnert G, Liebl J, Steier P, Buchholz BA, Eriksson M, Arner E, Hauner H, et al. 2011 Dynamics of human adipose lipid turnover in health and metabolic disease. Nature 110–113. (https://doi.org/10.1038/nature10426)
Arruda AP & Hotamisligil GS 2015 Calcium homeostasis and organelle function in the pathogenesis of obesity and diabetes. Cell Metabolism 381–397. (https://doi.org/10.1016/J.CMET.2015.06.010)
Avasthy N, Jeremy JY & Dandona P 1988 The role of calcium in mediating phorbol ester- and insulin-stimulated adipocyte lipogenesis. Diabetes Research 91–95.
Baumbach J, Hummel P, Bickmeyer I, Kowalczyk KM, Frank M, Knorr K, Hildebrandt A, Riedel D, Jäckle H & Kühnlein RP 2014 A Drosophila in vivo screen identifies store-operated calcium entry as a key regulator of adiposity. Cell Metabolism 331–343. (https://doi.org/10.1016/j.cmet.2013.12.004)
Bavley CC, Fischer DK, Rizzo BK & Rajadhyaksha AM 2017 Cav1.2 channels mediate persistent chronic stress-induced behavioral deficits that are associated with prefrontal cortex activation of the p25/Cdk5-glucocorticoid receptor pathway. Neurobiology of Stress 27–37. (https://doi.org/10.1016/J.YNSTR.2017.02.004)
Begum N, Sussman KE & Draznin B 1992 Calcium-induced inhibition of phosphoserine phosphatase in insulin target cells is mediated by the phosphorylation and activation of inhibitor 1. Journal of Biological Chemistry 5959–5963.
Bentley DC, Pulbutr P, Chan S & Smith PA 2014 Etiology of the membrane potential of rat white fat adipocytes. American Journal of Physiology: Endocrinology and Metabolism E161–E175. (https://doi.org/10.1152/ajpendo.00446.2013)
Blackmore PF & Augert G 1989 Effect of hormones on cytosolic free calcium in adipocytes. Cell Calcium 561–567. (https://doi.org/10.1016/0143-4160(89)90018-3)
Bleicher SJ, Farber L, Lewis A & Goldner MG 1966 Electrolyte-activated lipolysis in vitro: modifying effect of calcium. Metabolism: Clinical and Experimental 742–748. (https://doi.org/10.1016/s0026-0495(66)80010-0)
Buonarati OR, Henderson PB, Murphy GG, Horne MC & Hell JW 2017 Proteolytic processing of the L-type Ca2+ channel alpha 11.2 subunit in neurons. F1000Research 1166. (https://doi.org/10.12688/f1000research.11808.2)
Carmen GY & Víctor SM 2006 Signalling mechanisms regulating lipolysis. Cellular Signalling 401–408. (https://doi.org/10.1016/j.cellsig.2005.08.009)
Cignarella A 1994 Antithrombotic activity of nicardipine in spontaneously hypertensive rats. Pharmacological Research 273–280. (https://doi.org/10.1016/1043-6618(94)80109-6)
Clausen T & Martin BR 1977 The effect of insulin on the washout of [45Ca]calcium from adipocytes and soleus muscle of the rat. Biochemical Journal 251–255. (https://doi.org/10.1042/bj1640251)
Coombs JS, Curtis DR & Eccles JC 1956 Time courses of motoneuronal responses. Nature 1168–1169. (https://doi.org/10.1038/1781168a0)
Dolphin AC 2016 Voltage-gated calcium channels and their auxiliary subunits: physiology and pathophysiology and pharmacology. Journal of Physiology 5369–5390. (https://doi.org/10.1113/JP272262)
Draznin B, Sussman K, Kao M, Lewis D & Sherman N 1987 The existence of an optimal range of cytosolic free calcium for insulin-stimulated glucose transport in rat adipocytes. Journal of Biological Chemistry 14385–14388.
Draznin B, Sussman KE, Eckel RH, Kao M, Yost T & Sherman NA 1988 Possible role of cytosolic free calcium concentrations in mediating insulin resistance of obesity and hyperinsulinemia. Journal of Clinical Investigation 1848–1852. (https://doi.org/10.1172/JCI113801)
Efendić S, Alm B & Löw H 1970 Effects of Ca++ on lipolysis in human omental adipose tissue in vitro. Hormone and Metabolic Research 287–291. (https://doi.org/10.1055/s-0028-1095061)
El Hachmane MF & Olofsson CS 2018 A mechanically activated TRPC1-like current in white adipocytes. Biochemical and Biophysical Research Communications 736–742. (https://doi.org/10.1016/j.bbrc.2018.03.050)
Fagerberg L, Hallström BM, Oksvold P, Kampf C, Djureinovic D, Odeberg J, Habuka M, Tahmasebpoor S, Danielsson A, Edlund K, et al. 2014 Analysis of the human tissue-specific expression by genome-wide integration of transcriptomics and antibody-based proteomics. Molecular and Cellular Proteomics 397–406. (https://doi.org/10.1074/mcp.M113.035600)
Fain JN, Gokmen-Polar Y & Bahouth SW 1997 Wortmannin converts insulin but not oxytocin from an antilipolytic to a lipolytic agent in the presence of forskolin. Metabolism 62–66. (https://doi.org/10.1016/S0026-0495(97)90169-4)
Fink T, Lund P, Pilgaard L, Rasmussen JG, Duroux M & Zachar V 2008 Instability of standard PCR reference genes in adipose-derived stem cells during propagation, differentiation and hypoxic exposure. BMC Molecular Biology 98. (https://doi.org/10.1186/1471-2199-9-98)
Fleischmann BK, Murray RK & Kotlikoff MI 1994 Voltage window for sustained elevation of cytosolic calcium in smooth muscle cells. PNAS 11914–11918. (https://doi.org/10.1073/pnas.91.25.11914)
Gaion R & Krishna G 1983 Cyclic nucleotides and lipolysis in rat fat cells. Interaction between calcium ionophore A23187 and FCCP, uncoupler of oxidative phosphorylation. Life Sciences 571–576. (https://doi.org/10.1016/0024-3205(83)90201-1)
Gaur S, Yamaguchi H & Goodman HM 1996a Growth hormone regulates cytosolic free calcium in rat fat cells by maintaining L-type calcium channels. American Journal of Physiology C1478–C1484. (https://doi.org/10.1152/ajpcell.1996.270.5.C1478)
Gaur S, Yamaguchi H & Goodman HM 1996b Growth hormone increases calcium uptake in rat fat cells by a mechanism dependent on protein kinase C. American Journal of Physiology C1485–C1492. (https://doi.org/10.1152/ajpcell.1996.270.5.C1485)
Gaur S, Schwartz Y, Tai LR, Frick GP & Goodman HM 1998 Insulin produces a growth hormone-like increase in intracellular free calcium concentration in okadaic acid-treated adipocytes. Endocrinology 4953–4961. (https://doi.org/10.1210/endo.139.12.6387)
Gorzelniak K, Janke J, Engeli S & Sharma AM 2001 Validation of endogenous controls for gene expression studies in human adipocytes and preadipocytes. Hormone and Metabolic Research 625–627. (https://doi.org/10.1055/s-2001-17911)
Hardy RW, Ladenson JH, Hruska KA, Jiwa AH & McDonald JM 1992 The effects of extracellular calcium and epinephrine on cytosolic-free calcium in single rat adipocytes. Endocrinology 3694–3702. (https://doi.org/10.1210/endo.130.6.1597165)
Henneman E & Mendell LM 2010 Functional organization of motoneuron pool and its inputs. In Comprehensive Physiology. Hoboken, NJ, USA: John Wiley & Sons. (https://doi.org/10.1002/cphy.cp010211)
Hvarfner A, Bergström R, Lithell H, Mörlin C, Wide L & Ljunghall S 1988 Changes in calcium metabolic indices during long-term treatment of patients with essential hypertension. Clinical Science 543–549. (https://doi.org/10.1042/cs0750543)
Izawa T, Koshimizu E, Komabayashi T & Tsuboi M 1983 Effects of Ca2+ and calmodulin inhibitors on lipolysis induced by epinephrine, norepinephrine, caffeine and ACTH in rat epididymal adipose tissue. Nihon Seirigaku Zasshi: Journal of the Physiological Society of Japan 36–44.
Kelly KL, Jude T & Corkey BE 1989 Cytosolic free calcium in adipocytes. Journal of Biological Chemistry 26412754–12757.
Le Scouarnec S, Bhasin N, Vieyres C, Hund TJ, Cunha SR, Koval O, Marionneau C, Chen B, Wu Y, Demolombe S, et al. 2008 Dysfunction in ankyrin-B-dependent ion channel and transporter targeting causes human sinus node disease. PNAS 15617–15622. (https://doi.org/10.1073/pnas.0805500105)
Lee SC & Pappone PA 1997 Membrane responses to extracellular ATP in rat isolated white adipocytes. Pflugers Archiv 422–428. (https://doi.org/10.1007/s004240050416)
Lipscombe D, Helton TD & Xu W 2004 L-type calcium channels: the low down. Journal of Neurophysiology 2633–2641. (https://doi.org/10.1152/jn.00486.2004)
Louis C, Van Den Daelen C, Tinant G, Bourez S, Thomé JP, Donnay I, Larondelle Y & Debier C 2014 Efficient in vitro adipocyte model of long-term lipolysis: a tool to study the behavior of lipophilic compounds. In Vitro Cellular and Developmental Biology: Animal 507–518. (https://doi.org/10.1007/s11626-014-9733-6)
Martin BR, Clausen T & Gliemannt J 1975 Relationships between the exchange of calcium and phosphate in isolated fat-cells. Biochemical Journal 121–129. (https://doi.org/10.1042/bj1520121)
Miyauchi A, Hruska KA, Greenfield EM, Duncan R, Alvarez J, Barattolo R, Colucci S, Zambonin-Zallone A, Teitelbaum SL & Teti A 1990 Osteoclast cytosolic calcium, regulated by voltage-gated calcium channels and extracellular calcium, controls podosome assembly and bone resorption. Journal of Cell Biology 2543–2552. (https://doi.org/10.1083/JCB.111.6.2543)
N’Gouemo P, Akinfiresoye LR, Allard JS & Lovinger DM 2015 Alcohol withdrawal-induced seizure susceptibility is associated with an upregulation of CaV1.3 channels in the rat inferior colliculus. International Journal of Neuropsychopharmacology pyu123. (https://doi.org/10.1093/ijnp/pyu123)
Ni Z, Smogorzewski M & Massry SG 1994 Effects of parathyroid hormone on cytosolic calcium of rat adipocytes. Endocrinology 1837–1844. (https://doi.org/10.1210/endo.135.5.7525254)
Ni Z, Smogorzewski M & Massry SG 1995 Elevated cytosolic calcium of adipocytes in chronic renal failure. Kidney International 1624–1629. (https://doi.org/10.1038/ki.1995.226)
Pepe S, Bogdanov K, Hallaq H, Spurgeon H, Leaf A & Lakatta E 1994 Omega 3 polyunsaturated fatty acid modulates dihydropyridine effects on L-type Ca2+ channels, cytosolic Ca2+, and contraction in adult rat cardiac myocytes. PNAS 8832–8836. (https://doi.org/10.1073/pnas.91.19.8832)
Pershadsingh HA, Lee LY & Snowdowne KW 1989 Evidence for a sodium/calcium exchanger and voltage-dependent calcium channels in adipocytes. FEBS Letters 89–92. (https://doi.org/10.1016/0014-5793(89)81169-X)
Pfaffl MW 2001 A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Research e45. (https://doi.org/10.1093/nar/29.9.e45)
Raifman TK, Kumar P, Haase H, Klussmann E, Dascal N & Weiss S 2017 Protein kinase C enhances plasma membrane expression of cardiac L-type calcium channel, CaV1.2. Channels 604–615. (https://doi.org/10.1080/19336950.2017.1369636)
Ramírez-Ponce MP, Acosta J & Bellido JA 1990 Electrical activity in white adipose tissue of rat. Revista Espanola de Fisiologia 133–138.
Rueden CT, Schindelin J, Hiner MC, DeZonia BE, Walter AE, Arena ET & Eliceiri KW 2017 ImageJ2: ImageJ for the next generation of scientific image data. BMC Bioinformatics 529. (https://doi.org/10.1186/s12859-017-1934-z)
Rumiantsev DO, Piotrovskii VK, Metelitsa VI, Slastnikova ID, Martsevich Yu S & Kokurina EV 1989 Serum binding of nifedipine and verapamil in patients with ischaemic heart disease on monotherapy. British Journal of Clinical Pharmacology 357–361. (https://doi.org/10.1111/j.1365-2125.1989.tb05438.x)
Sattar N & Gill JM 2014 Type 2 diabetes as a disease of ectopic fat? BMC Medicine 123. (https://doi.org/10.1186/s12916-014-0123-4)
Schimmel RJ 1978 Calcium antagonists and lipolysis in isolated rat epididymal adipocytes: effects of tetracaine, manganese, cobaltous and lanthanum ions and D600. Hormone and Metabolic Research 128–134. (https://doi.org/10.1055/s-0028-1093458)
Schroeder A, Mueller O, Stocker S, Salowsky R, Leiber M, Gassmann M, Lightfoot S, Menzel W, Granzow M & Ragg T 2006 The RIN: an RNA integrity number for assigning integrity values to RNA measurements. BMC Molecular Biology 3. (https://doi.org/10.1186/1471-2199-7-3)
Schulla V, Renström E, Feil R, Feil S, Franklin I, Gjinovci A, Jing XJ, Laux D, Lundquist I, Magnuson MA, et al. 2003 Impaired insulin secretion and glucose tolerance in beta cell-selective Ca(v)1.2 Ca2+ channel null mice. EMBO Journal 3844–3854. (https://doi.org/10.1093/emboj/cdg389)
Schwartz Y, Goodman HM & Yamaguchi H 1991 Refractoriness to growth hormone is associated with increased intracellular calcium in rat adipocytes. PNAS 6790–6794. (https://doi.org/10.1073/PNAS.88.15.6790)
Seydoux J, Muzzin P, Moinat M, Pralong W, Girardier L & Giacobino J-PP 1996 Adrenoceptor heterogeneity in human white adipocytes differentiated in culture as assessed by cytosolic free calcium measurements. Cellular Signalling 117–122. (https://doi.org/10.1016/0898-6568(95)02035-7)
Shi L, Ko ML & Ko GY-P 2017 Retinoschisin facilitates the function of L-type voltage-gated calcium channels. Frontiers in Cellular Neuroscience 232. (https://doi.org/10.3389/fncel.2017.00232)
Silver N, Cotroneo E, Proctor G, Osailan S, Paterson KL & Carpenter GH 2008 Selection of housekeeping genes for gene expression studies in the adult rat submandibular gland under normal, inflamed, atrophic and regenerative states. BMC Molecular Biology 64. (https://doi.org/10.1186/1471-2199-9-64)
Soma MR, Gotto AM & Ghiselli G 1989 Rapid modulation of rat adipocyte lipoprotein lipase: effect of calcium, A23187 ionophore, and thrombin. Biochimica et Biophysica Acta 307–314. (https://doi.org/10.1016/0005-2760(89)90237-3)
Wang Z, Li V, Chan GCK, Phan T, Nudelman AS, Xia Z & Storm DR 2009 Adult type 3 adenylyl cyclase-deficient mice are obese. PLoS ONE e6979. (https://doi.org/10.1371/journal.pone.0006979)
Xu W & Lipscombe D 2001 Neuronal Ca(V)1.3alpha(1) L-type channels activate at relatively hyperpolarized membrane potentials and are incompletely inhibited by dihydropyridines. Journal of Neuroscience 5944–5951. (https://doi.org/21/16/5944)
Yue F, Cheng Y, Breschi A, Vierstra J, Wu W, Ryba T, Sandstrom R, Ma Z, Davis C, Pope BD, et al. 2014 A comparative encyclopedia of DNA elements in the mouse genome. Nature 355–364. (https://doi.org/10.1038/nature13992)
Ziegler R, Jobst W, Minne H & Faulhaber JD 1980 Calciotropic hormones and lipolysis of human adipose tissue: role of extracellular calcium as conditioning but not regulating factor. Endokrinologie 77–88.
