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Cardiovascular Research 2006 69(4):855-864; doi:10.1016/j.cardiores.2005.11.019
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Copyright © 2005, European Society of Cardiology

OxLDL enhances L-type Ca2+ currents via lysophosphatidylcholine-induced mitochondrial reactive oxygen species (ROS) production

Ian M. Fearon*

Faculty of Life Sciences, The University of Manchester, G.38 Stopford Building, Oxford Road, Manchester, M13 9PT, UK

* Tel.: +44 161 275 5496; fax: +44 161 275 5600. Email address: ian.fearon{at}manchester.ac.uk

Received 1 June 2005; revised 10 November 2005; accepted 11 November 2005


   Abstract
Top
 Abstract
1. Introduction
 2. Materials and methods
 3. Results
 4. Discussion
 References
 
Objective: To examine the mechanisms underlying oxidised LDL- (oxLDL)-induced alterations in Ca2+ currents, an effect which underlies altered vascular contractility and cardiac myocyte function.

Methods: Ca2+ currents (ICa) were recorded by whole-cell patch-clamp in HEK293 cells expressing L-type Ca2+ channel {alpha}1C subunits or isolated rat ventricular myocytes. oxLDL (but not native LDL) significantly enhanced recombinant ICa, an effect mimicked by 1 µM lysophosphatidylcholine (LPC). LPC failed to enhance ICa either in mitochondrial electron transport chain-depleted {rho}0 cells, or in the presence of rotenone (1 µM), or MPP+ (10 µM). The LPC response was similarly ablated by ascorbate (200 µM) or TROLOX (500 µM) and by the mitochondria-targeted antioxidant, MitoQ (250 nM). In myocytes, enhancement of ICa due to LPC was similarly abrogated with rotenone and MitoQ. These data suggest that LPC enhanced recombinant Ca2+ currents due to increased mitochondrial ROS production. In support with this, LPC enhanced fluorescence in HEK293 cells and cardiac myocytes loaded with a ROS-sensitive mitochondrial dye, reduced mitotracker red.

Conclusion: LPC up-regulates L-type Ca2+ currents due to altered mitochondrial ROS production, an effect which mediates the response of the native ICa in cardiac myocytes to oxLDL.

KEYWORDS Ca-channel; Mitochondria; Lipoproteins; Oxygen radicals


   1. Introduction
Top
 Abstract
 1. Introduction
2. Materials and methods
 3. Results
 4. Discussion
 References
 
Elevated blood levels of oxidatively modified low density lipoproteins (oxLDL) are a major factor in atherosclerosis [37], and there is abundant evidence of a role for LDL oxidation in atherogenesis [4]. OxLDLs accumulate in atherosclerotic lesions and trigger endothelial dysfunction [35], involving numerous changes in endothelial cells, including adhesion molecule and nitric oxide synthase (NOS) up-regulation [13,39], proliferation [12], impairment of endothelium-dependent vasodilation [21] and altered electrophysiological properties [23]. OxLDL also elicits atherogenic processes in vascular smooth muscle cells by stimulating proliferation [24] and regulating migration [1].

Lysophosphatidylcholine (LPC) is the major biologically active component of oxLDL, and is produced when phosphatidylcholine is degraded during oxidative lipoprotein modification [33,38]. LPC mediates atherogenic and contractile effects of oxLDL on vascular endothelial and muscle cells. In the endothelium, LPC mediates pro-inflammatory and cNOS-enhancing properties [3,13], and inhibition of migration and agonist-induced endothelium-dependent relaxation [8,32] due to oxLDL. In vascular myocytes, LPC elicits vasoconstriction [32], contributes to the synergy between oxLDL and endothelin-1 on proliferation [45], and stimulates expression of monocyte chemoattractant proteins [34].

OxLDL also directly regulates the electrophysiological properties of cardiac [48] and vascular [31] myocytes, contributing to altered electrical and contractile activity [26,31,48]. In both cell types, the underlying electrophysiological change is altered transmembrane Ca2+ flux through voltage-gated channels. Furthermore in cardiac myocytes, altered transmembrane Na+ and K+ fluxes contribute to altered electrophysiological properties in response to LPC [43,44]. In the present study, the mechanisms underlying the effects of oxLDL and LPC on Ca2+ currents were examined in recombinant and native systems. Similar to myocytes, oxLDL enhanced Ca2+ channel currents, an effect mimicked by LPC. The mechanism underlying this effect was LPC-induced enhancement of the functional expression of {alpha}1C subunits at the plasma membrane and involved mitochondrial ROS production. These data define a sub-cellular mechanism by which oxLDL affects both vascular contractility and electrophysiological and contractile function of cardiac myocytes.


   2. Materials and methods
Top
 Abstract
 1. Introduction
 2. Materials and methods
3. Results
 4. Discussion
 References
 
2.1 Culture of HEK293 cells
Experiments were conducted in HEK293 cells stably expressing human cardiac L-type Ca2+ channel {alpha}1C subunits [15,36]. Cells were grown in minimum essential medium (MEM; Invitrogen, Paisley, UK), containing 9% (v/v) foetal calf serum (Globepharm, Esher, Surrey), 1% non-essential amino acids, gentamicin (50 mg l–1), 10,000 µl–1 penicillin G, 10 mg l–1 streptomycin, 0.25 mg l–1 amphotericin and 400 mg l–1 G418 (all Invitrogen) at 37 °C in a humidified atmosphere of air/CO2 (95:5%).

2.2 Isolation of ventricular myocytes
Myocytes were isolated from rat ventricular muscle as previously described [9]. Rats were killed by stunning and cervical dislocation. Care and use of animals were in accordance with the Animals (Scientific Procedures) Act 1986, which conforms to NIH guidelines.

2.3 Electrophysiology
Dishes with attached cells were continually perfused and whole-cell patch-clamp recordings [11] were made using pipettes of resistance 4–6 M{Omega}. Recordings were made at room temperature (22 ± 2 °C). Current traces were filtered at 5 kHz and digitised at 10 kHz. Capacitative transients were minimised by analogue means and corrections for leak current were made off-line by appropriate scaling and subtraction of the average leak current evoked by small hyperpolarising and depolarising steps (<5 mV). Analyses and voltage protocols were performed using a Multiclamp 700B amplifier, with a Digidata 1322A interface and pCLAMP 9.2 software (Axon Instruments). Current densities (pA/pF) were calculated by dividing evoked currents by cell capacitance. Cells were voltage-clamped at either –80 mV (HEK293 cells) or –40 mV (ventricular myocytes). Currents were evoked by step depolarising the membrane to various test potentials for 100 ms at a frequency of 0.1 Hz. To minimise variation in Ca2+ currents, amplitudes following treatments were compared to those obtained in control cells on the same day. Time constants for channel activation and inactivation were obtained by fitting relevant portions of current records with a single exponential function. Steady-state activation was calculated as the fraction of current evoked by a 100 ms depolarisation to varying test potentials compared with the maximal evoked current. Steady-state inactivation was calculated as a function of the prepulse potential; cells were prepulsed to varying potentials for 5 s, followed by a 100 ms test pulse to +10 mV. Boltzmann fits to steady-state activation and inactivation curves and graphing of the data were performed using Microcal Origin.

HEK293 cells were perfused with (mM): NaCl, 95; CsCl, 5; MgC12, 0.6; BaCl2 20; Hepes, 5; D-glucose, 10; TEA-Cl, 20 (pH 7.4 with NaOH). Myocytes were perfused with (mM): NaCl, 120; CaCl2, 2; KCl, 5; MgCl2, 2, TEA, 20; glucose, 10 and HEPES 10 (pH 7.4 with NaOH). Electrodes were filled with (mM): CsCl, 120; TEA-Cl, 20; MgC12, 2; EGTA, 10; Hepes, 10; ATP, 2 (pH 7.2 with CsOH).

2.4 Western blotting
Cells were removed from culture dishes by incubating in Ca2+-free PBS with 2 mM EDTA for 5 min at 37 °C. To obtain a membrane-enriched fraction, cell pellets were resuspended in 20 mM HEPES, 250 mM sucrose, 2 mM EDTA and protease inhibitors and centrifuged at 1200 x g for 10 min at 4 °C. The supernatant was retained and the pellet re-suspended and re-centrifuged. Supernatants from both centrifugations were pooled and centrifuged at 9000 x g for 10 min. The resultant supernatant was centrifuged at 20,000 g for 80 min at 4 °C. The pellet was resuspended in 20 mM Tris and 2% SDS. Protein concentration was calculated using the BCA protein assay kit (Pierce, Rockford, IL, USA). 40 µg of control and LPC-treated membrane fractions were resolved by SDS-PAGE gel electrophoresis and probed with an anti-Cav 1.2 antibody (1:200; Alomone Labs, Jerusalem, Israel). Membranes were further probed using a horseradish peroxidase-conjugated IgG (Jackson ImmunoResearch, West Grove, PA, USA) as a secondary antibody, and proteins visualised using enhanced chemiluminescent substrate (Amersham Biosciences, Piscataway, NJ).

2.5 Creation of mitochondria-depleted ({rho}0) HEK293 cells
The mitochondrial DNA-depleted ({rho}0; [18]) cell line was generated from HEK293 cells stably expressing {alpha}1C subunits, as described previously [5]. Briefly, cells were grown in MEM supplemented with 2 µg/ml ethidium bromide, 1 mM sodium pyruvate and 50 µg/ml uridine. Cells exhibited a 98% reduction in mtDNA levels, determined by PCR examination utilising mtDNA-specific primer sets. [5], and failed up to take up mitotracker red (Fig. 3A).


Figure 3
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  		Fig. 3 Enhancement of Ca2+ currents by LPC requires ROS production in the mitochondrial ETC. (A) Epifluorescence images of wild-type ({rho}+; left) and mitochondria-depleted ({rho}0; right) HEK293 cells stained with the mitochondria-selective dye, mitotracker red. The lack of fluorescence in the {rho}0 cells indicates their lack of a functional ETC. (B) Mean (± S.E.M.) current density–voltage (I–V) relationships demonstrating the lack of enhancing effect of LPC (1 µM for 3–4 h) on Ca2+ currents in {rho}0 cells. Currents were evoked by step depolarising cells for 100 ms to the indicated potential (holding potential –80 mV). Data were averaged from 8 cells (control) and 8 cells (LPC). (C) Enhancement of Ca2+ currents by LPC was inhibited by the mitochondrial complex I inhibitors rotenone (1 µM) and MPP+ (10 µM), and the antioxidants ascorbic acid (200 µM), TROLOX (500 µM) and MitoQ (250 nM). 10 µM TTFA, 500 nM myxothiazol, 1 µg/ml antimycin A, 500 µM sodium cyanide and the triphenylphosphonium (TPP) cation were all without effect on the response to LPC. Shown are mean (± S.E.M.) current densities evoked by step depolarising cells for 100 ms to +10 mV (holding potential, –80 mV). Data were averaged from between 6 and 16 cells in each case. *P>0.05 compared to control.

 
2.6 Measurement of LPC-induced mitochondrial ROS production
Cells were seeded onto glass coverslips. Following LPC treatment, media was removed and replaced with one containing reduced mitotracker red (500 nM). After incubation for 15 min at 37 °C, slips were rinsed three times with PBS and fixed at room temperature for 20 min in 4% paraformaldehyde in PBS. Slips were further rinsed three times with PBS, and mounted onto glass slides using Vectashield (Vector Laboratories, Peterborough, UK). Cells were visualised using a Zeiss (Welwyn Garden City, UK) Axioplan 2 upright microscope equipped with epifluorescence and a rhodamine filter set. Digital images were taken with a QImaging (Burnaby, BC, Canada) QICAM 12 bit monochrome digital camera and QCapture Pro 5.0 software. Mitochondrial ROS production was quantified using a protocol similar to that described previously [2,20]. To achieve this, line-scan pixel intensity analysis was carried out and the peak intensity of each cell used for statistical analysis. In HEK293 cells, background fluorescence was normalised by subtracting the pixel intensity at the cell centre.

2.7 Isolation and oxidation of lipoproteins
Human LDL ({partial}=0.95 g/ml) was isolated from plasma, by sequential ultracentrifugation as described [28]. LDL was oxidised by the addition of 5 µM CuSO4 in PBS at 37 °C for 4 h in the dark. Oxidation was halted by the addition of 250 µM EDTA and placing the sample at 4 °C. Lipid-peroxide concentration was determined by a spectrophotometric method, as previously described [29].

2.8 Chemicals and statistical analyses
LPC, nifedipine, rotenone, antimycin A, myxothiazol, TTFA, MPP, sodium cyanide, ascorbic acid and TROLOX were from Sigma (Poole, Dorset, UK). MitoQ and triphenylphosphonium (TPP) were generously provided by Dr. Michael Murphy (Medical Research Council Dunn Human Nutrition Unit, Cambridge). Mitochondrial inhibitors and antioxidants were applied for an hour prior to, and during, incubation in LPC. Mitotracker probes were from Invitrogen. Results are expressed as means ± S.E.M. Statistical comparisons of discrete data points were made using paired or unpaired Student's t-tests. Differences in current density–voltage (I–V) relationships were compared using two-way ANOVA, over voltage ranges at which channels were activated.


   3. Results
Top
 Abstract
 1. Introduction
 2. Materials and methods
 3. Results
4. Discussion
 References
 
3.1 OxLDL and LPC enhanced recombinant Ca2+ currents
Initial experiments were carried out in HEK293 cells stably expressing human cardiac L-type Ca2+ channel {alpha}1C subunits [15,36], which are abundantly expressed in cardiac and vascular myocytes [19]. Previous studies on native currents in vascular and cardiac myocytes demonstrated an enhanced L-type Ca2+ current in response to incubation in oxidised LDL (oxLDL) [31,48]. As shown in Fig. 1A, this effect was mimicked in the recombinant system by incubating cells for 6 h in CuSO4-oxidised LDL, with the lipoprotein causing a significant enhancement of Ca2+ currents (P<0.01, ANOVA) For example at a test potential of +10 mV, mean (± S.E.M.) Ca2+ current densities were –6.1 ± 0.8 pA/pF (n=9) in control cells and –8.9 ± 1.7 pA/pF (n=10) in cells incubated in oxLDL. In contrast, native (non-oxidised) LDL was without significant effect on Ca2+ currents (P>0.05, ANOVA; Fig. 1B). Currents following incubation in native LDL were –7.1 ± 1.4 pA/pF (n=6; P>0.05 compared to controls.


Figure 1
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  		Fig. 1 oxLDL and LPC enhance recombinant Ca2+ currents. Recordings were made in HEK293 cells expressing human cardiac L-type Ca2+ channel {alpha}1C subunits [15]. (A) Mean (± S.E.M.) current density–voltage (I–V) relationships demonstrating the increase in Ca2+ currents following a 6-h incubation in oxLDL. (B) Non-oxidised LDL was without significant effect on Ca2+ channel currents. (C) The effect of oxLDL was mimicked by incubating cells for 3–4 h in the major biological component of oxLDL, lysophosphatidylcholine (LPC; 1 µM). (D) LPC was without effect on steady-state activation (circles) and inactivation (squares). LPC was further without effect on kinetics of activation (E), expressed as the time constant ({tau}) obtained by fitting a single exponential function to the activating section of current records. (F) Incubation of cells for 3–4 h in 1 µM C10 had no effect on Ca2+ channel currents. In panels A, B, C and F, currents were evoked by depolarising cells for 100 ms to the indicated test potential (holding potential –80 mV). Insets in these panels show typical individual traces obtained when cells were step depolarised to +10 mV under the indicated conditions. Data were averaged from between 6 and 10 cells in each case. (G) LPC enhanced the membrane expression of L-type {alpha}1C subunits. Membrane preparations were electrophoresed by SDS-PAGE and probed using an {alpha}1C-specific primary antibody. Equal amounts of protein were loaded into each lane. Image shown is typical of results of three identical experiments.

 
To test the hypothesis that the major biologically active component of oxLDL (lysophosphatidylcholine (LPC) [33,38]) mediated these effects of oxLDL, cells expressing {alpha}1C subunits were incubated in 1 µM LPC for 3–4 h. Following this incubation, Ca2+ currents were significantly enhanced (P<0.01, ANOVA; Fig. 1C), such that current densities were increased from –6.7 ± 1.5 pA/pF (n=6) in control cells to 9.7 ± 1.6 pA/pF (n=16) following incubation in LPC. LPC was without effect on steady-state activation or inactivation (Fig. 1D). V0.5 and slope factors for activation were –2.8 ± 0.6 mV and 6.9 ± 0.5 under control conditions, and –3.0 ± 0.3 mV and 6.6 ± 0.2 following LPC treatment (n=6 and P>0.05 in each case). Similarly, V0.5 and slope factors for inactivation were –7.6 ± 2.8 mV and 19.5 ± 2.9 under control conditions, and –7.5 ± 1.0 mV and 18.1 ± 1.0 after exposure to LPC (n=6 and P>0.05 in each case). LPC was also without effect on the kinetics of channel activation (Fig. 1E). For example, at +10 mV the time constant ({tau}) for activation was 1.4 ± 0.2 ms under control conditions and 1.3 ± 0.1 ms after LPC (n=6; P>0.05).

The effect of LPC on Ca2+ currents was not due to a nonspecific effect of the lipid, since the chemically similar but biologically inert C10 lipid (1 µM for 3–4 h) was without effect on Ca2+ currents. At all activating test potentials, current densities in cells exposed to the C10 lipid were not significantly different to those in cells incubated in its absence (P>0.05, ANOVA; Fig. 1F). For example, at +10 mV, current densities following incubation in C10 lipid were –7.1 ± 1.2 pA/pF, compared to –6.7 ± 1.5 pA/pF in control cells (n=6 in each case). Together, these data support the proposal that LPC mediates the enhancement of Ca2+ currents due to oxLDL. This effect of was one of increased membrane expression of {alpha}1C subunits, since immunohistochemical detection of these subunits using an {alpha}1C subunit-specific primary antibody was increased by LPC when probed by Western blotting of a membrane-enriched protein fraction (Fig. 1G).

3.2 LPC selectively enhanced {alpha}1C subunit currents
LPC-enhanced Ca2+ currents were completely abolished by the selective L-type Ca2+ channel blocker, nifedipine (2 µM). Fig. 2A shows a representative time-series recording in which Ca2+ currents in an LPC-treated cell were fully inhibited by nifedipine. When mean data were calculated in 4 such cells, Ca2+ currents at +10 mV in cells incubated in the lipid were –12.5 ± 3.5 pA/pF, and were reduced to –0.0 ± 0.4 pA/pF in the presence of nifedipine (P<0.05). LPC was without significant effect on Ca2+ currents in untransfected cells (P>0.05, ANOVA; Fig. 2B), further suggesting that its effect was one of specific up-regulations of the {alpha}1C subunit and was not due to a nonspecific effect on other transmembrane Ca2+ conductance pathways [41].


Figure 2
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  		Fig. 2 LPC selectively enhanced recombinant L-type Ca2+ currents. (A) Time-series recording showing inhibition of LPC-enhanced whole-cell Ca2+ current by 2 µM nifedipine (horizontal bar). Currents were evoked by step depolarising cells for 100 ms to +10 mV (holding potential –80 mV). Inset, individual current records taken from the time series recording, under control conditions and in the presence of nifedipine as indicated. Typical of 4 such recordings. For 3–4 h prior to recording, cells were incubated in 1 µM LPC. (B) Mean (± S.E.M.) current density–voltage (I–V) relationships obtained in untransfected HEK293 cells, incubated under control conditions and in the presence of 1 µM LPC for 3–4 h, as indicated. Currents were evoked by step depolarising cells for 100 ms to the indicated potential (holding potential –80 mV). Data were averaged from 9 cells (control) and 10 cells (LPC).

 
3.3 Role of ROS production in the mitochondrial ETC in the response to LPC
LPC regulates mitochondrial function [16,30], while surface expression of recombinant {alpha}1C subunits is regulated by altered function of the mitochondrial electron transport chain (ETC) [5]. To test the involvement of the ETC in LPC-induced up-regulation of Ca2+ currents, the effects of LPC were examined in {rho}0 HEK293 cells expressing {alpha}1C subunits [5]. These cells failed to take up the mitochondria-selective dye, mitotracker red (Fig. 3A), demonstrating their ETC deficiency.

In contrast to its effect on currents in wild-type ({rho}+) HEK293 cells (Fig. 1C), 1 µM LPC was without effect in {rho}0 cells (P>0.05, ANOVA; Fig. 3B). At +10 mV, Ca2+ current densities were –5.7 ± 1.3 pA/pF (n=8) in {rho}0 cells under control conditions and –5.0 ± 0.9 pA/pF (n=8) in cells incubated in LPC. While two-way ANOVA indicated no significant difference between control and LPC I–V curves, there was a small systematic reduction in current over the potential range of 0 mV to 50 mV which I interpret as physiologically insignificant, and which is in contrast to the enhancing effect of LPC in the {rho}+ cells.

Effects of LPC were also abrogated by co-incubating with the mitochondrial complex I inhibitors, rotenone (1 µM) and MPP+ (10 µM; Fig. 3C). In contrast, the effects of LPC were unaltered by the complex II inhibitor TTFA (10 µM), the complex III inhibitors myxothiazol (500 nM) and antimycin A (1 µg/ml) and the complex IV inhibitor, sodium cyanide (500 µM; Fig. 3C). Thus, the effects of LPC required a functional mitochondrial ETC and involved regulation of mitochondrial complex I.

The ETC is an abundant source of reactive oxygen species (ROS; [42]), and is a source of superoxide [25] which regulates Ca2+ channel expression [5]. To examine the role of ROS in LPC-mediated Ca2+ current enhancement, we tested its effects in the presence of two antioxidants, ascorbic acid (200 µM) and TROLOX (500 µM). In each case, Ca2+ current enhancement due to LPC was abolished (Fig. 3C). Similarly, the mitochondria-targeted antioxidant, MitoQ (250 nM; [17]), abolished the response to LPC (Fig. 3C). The lipophilic triphenylphosphonium (TPP; 250 nM) cation to which the ubiquinol antioxidant is attached [17] was without effect on the response to LPC (Fig. 3C). Taken together with the lack of effect of LPC in {rho}0 cells and in the presence of rotenone or MPP+, these data suggest that LPC-mediated enhancement of Ca2+ currents involved ROS production in complex I.

To demonstrate that LPC induces mitochondrial ROS production, levels of these oxidants were assayed by determining the intensity of fluorescence following loading of cells with reduced mitotracker red, in a manner similar to that described previously [2,20]. This probe only fluoresces following activation by oxidation and mitochondrial sequestration, producing a ROS-specific mitochondrial staining amenable to quantitative measurement [10]. As shown in Fig. 4A, cells treated with LPC showed an increased fluorescence, suggesting that LPC increased mitochondrial ROS production. To quantify this, line-scan pixel intensity analysis was performed on 40 cells (20 cells in each of two, independent experiments) in each treatment group. Plots representative of those obtained by such analysis are shown in Fig. 4B. Mean (± S.E.M.) background-subtracted peak pixel intensities were 19.8 ± 2.3 in control cells and 43.3 ± 2.2 in cells treated with 1 µM LPC for 4 h (P<0.05; Fig. 4C).


Figure 4
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  		Fig. 4 LPC induces mitochondrial oxidant production. (A) Epifluorescence images of wild-type ({rho}+) HEK293 cells obtained following incubation in reduced mitotracker red. This probe does not fluoresce until it enters an actively respiring cell, where it is oxidised to the corresponding fluorescent mitochondrion-selective probe and sequestered into the mitochondria. Cells were exposed to reduced mitotracker red following a prior incubation under control conditions (left) or for 4 h in the presence of 1 µM LPC (right). In LPC-treated cells the intensity of fluorescence was increased, suggestive of increased ROS production [10]. To quantify this, line scan pixel intensity analysis was performed on 40 cells in each treatment group. (B) Exemplar line scans, obtained from cells indicated in (A). (C) Mean (± S.E.M.) intensity data from line scans. Data are background-subtracted peak pixel intensities, averaged from 40 cells in each treatment group.

 
The above data demonstrate a role for LPC-induced mitochondrial ROS production in mediating Ca2+ current enhancement. In support of this proposal, 1 µM LPC enhanced native Ca2+ currents in ventricular myocytes (P<0.001, ANOVA; Fig. 5A) without affecting activation and inactivation kinetics (Fig. 5B). This effect of LPC was ablated by both 1 µM rotenone and 250 nM MitoQ (Fig. 5C). Finally, LPC caused enhanced mitochondrial ROS production in the native cells, as determined by enhanced fluorescence in cells loaded with reduced mitotracker red (Fig. 5D–F). Thus, similar to the recombinant system, the effect of LPC in enhancing native ICa was mediated via mitochondrial ROS production.


Figure 5
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  		Fig. 5 Experiments were carried out in rat ventricular myocytes. (A) 1 µM LPC enhanced Ca2+ currents. Shown are mean (± S.E.M.) current densities evoked by step depolarising cells for 100 ms to the indicated potential (holding potential, –40 mV). Data were averaged from 9 cells (control) and 6 cells (LPC). (B) LPC was without effect on activation (upper) and inactivation (lower) kinetics of the native Ca2+ current. Data were averaged from 6 cells in each case. (C) LPC-induced current enhancement was ablated by 1 µM rotenone and 250 nM MitoQ. Shown are mean (± S.E.M.) current densities evoked by step depolarising cells for 100 ms to +10 mV (holding potential –80 mV). Data were averaged from between 5 and 11 cells in each case. *P>0.05 compared to control. (D) Epifluorescence images of ventricular myocytes obtained following incubation in reduced mitotracker red. Cells were exposed to the dye following a prior incubation under control conditions (left) or for 4 h in the presence of 1 µM LPC (right). To quantify the increase in fluorescence intensity in the LPC-treated cells, line scan pixel intensity analysis was performed on 45 cells (control) and 51 cells (LPC). (E) Exemplar line scans, obtained from cells in panel D. Mean (± S.E.M.) peak intensity data from these cells are presented in (F).

 

   4. Discussion
Top
 Abstract
 1. Introduction
 2. Materials and methods
 3. Results
 4. Discussion
References
 
Elevation of plasma oxLDL is causative of atherosclerosis [36], and numerous studies have identified a role for oxLDL in mediating atherogenic processes in endothelial cells and vascular myocytes. oxLDL also regulates ion channel expression and function in both cardiac and systemic vascular myocytes, in each case sufficient to cause significant alteration of the cell's contractile and electrical properties [26,31,43,44,48]. Importantly in native cells, up-regulation of L-type Ca2+ currents mediated responses to oxLDL [26,31,48].

In the present study, the mechanisms underlying these responses were examined in a recombinant system. In HEK293 cells expressing L-type Ca2+ channel {alpha}1C subunits in the absence of auxiliary subunits, CuSO4-oxidised LDL up-regulated Ca2+ currents. Thus, the effect of oxLDL was mediated by altered expression and/or function of the pore-forming subunit, and the similarity of the effect in the isolated and cultured cells provided a strong basis for the further examination of the mechanisms underlying oxLDL-mediated enhancement of Ca2+ currents. Comparable effects to those of oxLDL were observed on Ca2+ currents in both native ventricular myocytes and the recombinant system when cells were exposed over a similar time-course to the active component of the lipoprotein, LPC [33,38]. While this effect of LPC contrasted with the moderate inhibition of Ca2+ currents reported previously [27], this highlights the ability of LPC to regulate Ca2+ currents differentially, and possibly by different mechanisms, depending on the length and concentration of exposure. Over the prolonged period examined in these studies, LPC increased the amount of {alpha}1C subunit protein in membrane-enriched fractions, suggesting that the long-term response to LPC was due to increased surface expression of this subunit.

LPC-mediated enhancement of Ca2+ currents was abrogated by both cellular and mitochondria-targeted antioxidants, suggesting that ROS production was involved in the response. In endothelial cells, LPC causes the production of superoxide (O2–) anions via stimulation of NADPH oxidase [22,40], an effect which may play a role in the proliferative and atherogenic effects of oxLDL [12,45,46]. Contrastingly, the source of ROS in the present study was the mitochondrial ETC, since no effects of LPC were observed in HEK293 cells depleted of a functional ETC. This finding is in accordance with the reported alteration of mitochondrial enzyme activity and ROS production induced by LPC [16,30]. Furthermore, oxLDL was recently demonstrated to cause mitochondrial ROS production in isolated aortic endothelial cells [47], an effect which was ablated in mitochondria-depleted {rho}0 cells. Taken together with the data presented here, this suggests that LPC is the component of oxLDL capable of inducing ROS formation. The enhanced fluorescence of the mitochondria-selective and oxidant-sensitive dye, reduced mitotracker red [10], in cells exposed to LPC strongly supports this suggestion.

The specific site involved in LPC-induced mitochondrial ROS production may reside within complex I, since LPC-stimulated enhancement of Ca2+ currents was ablated by the complex I inhibitor, rotenone. In support of this, fluorescence measurements using the ROS-sensitive dye DCF demonstrated that mitochondrial ROS production due to oxLDL in vascular smooth muscle cells was rotenone-sensitive [14]. Moreover, enzyme activity assays showed an induction of complex I activity, but not of the activities of complex IV or citrate synthase, by oxLDL in human umbilical vein endothelial cells [6].

In summary, LPC mediates a cardiovascular response to oxLDL by inducing ROS production within complex I of the mitochondrial ETC to promote enhancement of Ca2+ channel {alpha}1C subunit expression. These data shed new light into the mechanisms underlying physiological and pathological regulation of the cardiovascular system in those exposed long-term to elevated plasma oxLDL.


   Acknowledgements
 
IMF was supported by a Grant-in-Aid (Grant # NA 5230) from the Heart and Stroke Foundation of Ontario. LDL was kindly provided by Drs. Mike Mackness and David Hine (Faculty of Medicine, The University of Manchester). Ventricular myocytes were provided by Dr. David Eisner (Unit of Cardiac Physiology, The University of Manchester). I am grateful to Dr. Mauro Esposti for providing the mitotracker probes and for helpful advice concerning this work. I am further grateful to Dr. I. O'Kelly for assistance with Western blotting. MitoQ and TPP were kindly provided by Dr. Michael Murphy (Medical Research Council Dunn Human Nutrition Unit, Cambridge).


   Notes
 
Time for primary review 15 days


   References
Top
 Abstract
 1. Introduction
 2. Materials and methods
 3. Results
 4. Discussion
 References
 

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