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CELLULAR |
Department of
1 Anaesthesiology
2 Physiology
3 Pharmacology and Toxicology, Medical College of Wisconsin, Milwaukee, WI 53226, USA
4 Department of Anaesthesiology and Intensive Care Medicine, Medical University of Graz, Austria
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Abstract |
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(Received 29 July 2006;
accepted after revision 31 August 2006;
first published online 7 September 2006)
Corresponding author M. Bienengraeber: Department of Anesthesiology, Medical College of Wisconsin, 8701 Watertown Plank Road, Milwaukee, WI 53226, USA. Email: mbieneng{at}mcw.edu
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Introduction |
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Therefore, the goal of this investigation was to assess the protective mechanism of mitochondrial K+ flux independently from pharmacological modulators. In the present study, we overexpressed Kir6.2 K+ channel in the mitochondria of HL-1 and HEK293 cell lines, and tested the hypothesis that an increased K+ influx to mitochondria confers protection against metabolic stress.
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Methods |
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The cDNA for Kir6.2 was amplified with Pwo DNA polymerase (Roche, Basel, Switzerland) from mouse Quick-Clone cDNA (Clontech, Mountain View, CA, USA) using 5'-ATAGAATGCGGCCGCACTGTCCCGAAAGGGCATTATCC-3' as forward and 5'-TAAGACTGCGGCCGCATCAGGACAAGGAATCTGGAGAG-3' as reverse primer, and inserted into the NotI site of the pCMV/myc/mito/GFP vector (Invitrogen, Carlsbad, CA, USA), in frame with the mitochondrial targeting sequence and green fluorescence protein (GFP). Kir6.2 cDNA was also inserted into the NotI site of pCMV/myc/mito in order to express Kir6.2 in the absence of GFP (myc/mito–Kir6.2). Point mutations (132GFG134 to AAA) were introduced in the K+-selectivity filter of Kir6.2 to create inactive mito/GFP–Kir6.2AAA (Koster et al. 2002) by performing PCR with complementary primers with the desired amino acid changes (QuickChange, Stratagene, La Jolla, CA, USA). The following forward primer was used (mutations are marked in bold): 5'-GAGGTCCAGGTGACCATTGCTGCCGCCGGACGCATGGTGACAGAG-3'. The reverse primer was complementary. The proper identity and orientation of the constructs were confirmed by DNA sequencing.
HEK293 cells were cultured in DMEM medium containing 10% fetal bovine serum (FBS). HL-1 cells, a murine cardiac muscle cell line (Claycomb et al. 1998), were cultured in Claycomb-Medium (JRH Biosciences, Lenexa, KS, USA) supplemented with 10% FBS, 4 mM glutamine, 10 µM noradrenaline and 1% penicillin/streptomycin on gelatine/fibronectin-coated flasks (White et al. 2004). Cells were maintained in a humidified 5% CO2 incubator at 37°C and passaged the day before transfection at about 50% confluence in 35 mm tissue culture dishes. Transfection was performed in serum-free DMEM medium containing 6 µl Fugene reagent (Roche) and 2 µg plasmid DNA encoding the desired construct. The cells were cultured for an additional 48 h before conducting the experiments. For stable transfection, HL-1 or HEK293 cells were grown in the presence of 600 µg ml–1 neomycin, and colonies were selected to test for expression by GFP fluorescence and Western blot.
Western blotting
The microsomal and mitochondrial fractions were isolated from transfected HEK293 and HL-1 cells by differential centrifugation in 0.3 M mannitol, 0.1% bovine serum albumin, 2 mM EDTA, 10 mM Hepes, pH 7.4. After cell homogenization with a glass homogenizer, the suspension was centrifuged for 10 min at 1000 g at 4°C (the supernatant represents the cellular fraction), followed by supernatant centrifugation at 14 000 g for 15 min at 4°C. The supernatant was centrifuged at 100 000 g for 60 min to pellet the microsomal fraction. The resulting pellets were washed twice with cold isolation buffer, and the protein content was determined using the Lowry method (BIO-RAD, Hercules, CA, USA). Equivalent amounts of protein samples (50 µg) were separated on a 4–20% polyacrylamide gel, and then Western blotting performed as described (Chiari et al. 2005) using a 1: 200 dilution of a rabbit-raised antibody against Kir6.2 (Santa Cruz Biotechnology, Santa Cruz, CA, USA). The blots were stripped and reprobed with an antibody against subunit I of cytochrome c oxidase (Invitrogen) as a marker for mitochondria.
Laser-scanning confocal microscopy
HEK293 and HL-1 cells were visualized using an inverted laser-scanning confocal microscope (Eclipse TE2000-U, Nikon Inc., Japan) with a x40/1.3 oil-immersion objective. Probes were excited at 488 nm with an argon laser, and at 543 nm with a Green HeNe laser. The scanning speed was set to a minimal pixel dwell time of 1.92 µs, and a set of filters (ND4 and ND8) was used in order to minimize dye bleaching. Each 512 x 512-pixel image was averaged twice via software-selected repeated line scan mode to ameliorate signal-to-noise ratio. Data were analysed using Metamorph 6.1 software (Universal Imaging, West Chester, PA, USA).
Mitochondrial staining with MitoFluor Red and colocalization analysis
Cells were loaded with mitochondrial marker MitoFluor Red589 (MFR, Invitrogen) for 10 min (200 nM), and washed three times before confocal analysis. For colocalization analysis, GFP fluorescence was monitored through a 515 nm barrier filter (excitation by argon laser), and MFR fluorescence through a 590 nm filter (excitation by Green HeNe). Each fluorescent wavelength was recorded independently and then combined to create a composite image.
Mitochondrial potassium uptake
Cultured HEK293 cells were loaded with a K+-sensitive fluorescent indicator PBFI AM in a loading protocol that allowed exclusive labelling of the mitochondria (Zoeteweij et al. 1994; Xu et al. 2002). In order to determine the rate of K+ uptake to the mitochondrial matrix, the bath solution was rapidly switched from 0 to 50 mM K+, and net K+ influx was recorded as the change in PBFI fluorescence ratio from two excitation wavelengths (340/380 nm, emission at 510 ± 20 nm). To ensure that the response was evoked through K+-selective channels, the rate of K+ influx was also monitored in the presence of the K+ channel blocker Ba2+ (1 mM). A more detailed description of mitochondrial K+ uptake measurements is provided in the Supplemental material.
Hypoxic stress and assessment of cell damage
To render HL-1 and HEK293 cells hypoxic, serum- and glucose-free DMEM was saturated with 5% CO2 and 95% N2, in the presence of 2-deoxyglucose (10 mM). Culture dishes were put in an airtight chamber in the incubator and flushed with the hypoxic gas mixture. The cells were exposed to hypoxia for 4 h, after which the hypoxic medium was replaced by DMEM containing glucose and 10% FBS, and cells were allowed to reoxygenate for 12 h. The cells that underwent such treatment were compared to control cells that were kept in normal DMEM containing 10% FBS, without hypoxia. In order to assess the degree of cellular damage, LDH release was measured in the supernatant of culture medium, according to the manufacturer's instructions (Diagnostic Chemicals Limited, Oxford, CT, USA) at a wavelength of 340 nm and expressed as percentage change over control (normoxia). In parallel, the cell viability was assessed using the MTT assay. Formazan formation was quantified spectrophotometrically at a wavelength of 570 nm with background subtraction at 650 nm.
Monitoring of mitochondrial Ca2+ during metabolic inhibition
The cells were incubated with Ca2+-sensitive fluorescent indicator rhod-2 AM (10 µM, Invitrogen) in a two-step cold/warm loading protocol (60 min at 4°C and 120 min at 37°C) (Trollinger et al. 2000). After loading, the cells were superfused with a normal external solution (mM: 140 NaCl, 5 KCl, 1.5 CaCl2, 1 MgSO4, 10 Hepes, and 5.5 glucose, pH 7.4). Metabolic inhibition was accomplished by switching to a solution containing (mM): 135 NaCl, 5 KCl, 1.5 CaCl2, 1 MgSO4, 10 Hepes, 10 2-deoxyglucose, and 2.5 NaCN, pH 7.4. Rhod-2 fluorescence was acquired at 590 nm (excitation with Green HeNe). Only the cells that expressed a similar degree of initial rhod-2 loading were analysed. Details on monitoring of mitoCa2+ are provided in the Supplemental material.
Analysis of mitochondrial membrane potential
HEK293 or HL-1 cells were incubated with the mitochondrial membrane potential (
m) indicator tetramethylrhodamine (TMRE, 100 nM, Invitrogen) for 30 min in the culture medium. TMRE fluorescence was obtained through a 590 nm barrier filter upon excitation by Green HeNe. All experiments were performed using identical image settings (gain, pinhole size, objective, filters) and conditions of TMRE incubation. Membrane potential was monitored under control conditions and during cell exposure to various drugs, as well as during metabolic inhibition. Throughout the experiments, TMRE was included in superfusing solutions.
Statistical analysis
Data are presented as mean ± S.E.M. and the number of cells or experiments is shown as n. Statistical comparisons were performed using one-way analysis of variance with Bonferroni's post hoc test. Differences at P < 0.05 were considered significant.
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Results |
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The cDNA encoding for Kir6.2 was amplified by PCR and subcloned into the vector pCMV/mito/GFP in frame and downstream from GFP and the mitochondrial targeting sequence (Fig. 1A). HEK293 cells, lacking expression of endogenous KATP channels (Giblin et al. 1999), were transfected with the generated construct (pCMV/mito/GFP–Kir6.2) and loaded with the mitochondrial marker MFR. GFP fluorescence revealed the intracellular distribution of GFP–Kir6.2 (green, Fig. 1B), and MFR fluorescence displayed the mitochondrial pattern within the cells (red). Colocalization was confirmed by merging the two images (yellow, overlay) and it verified mitochondrial localization of Kir6.2 in transfected cells.
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70 kDa in the cells transfected with pCMV/mito/GFP–Kir6.2 construct (Fig. 1C), which corresponds well with the estimated molecular weight of the GFP–Kir6.2 complex. Mitochondrial enrichment was confirmed by probing against subunit I of cytochrome c oxidase. Lack of GFP–Kir6.2 expression in the plasma membrane was verified by probing the microsomal fraction with Kir6.2 antibody, as well as with patch-clamp recordings (Supplemental material). Mitochondrial Kir6.2 expression was also confirmed in HEK293 cells transfected with inactive mito/GFP–Kir6.2AAA construct, stably transfected HL-1 cardiomyocytes (Fig. 1D) and in cells transfected with myc/mito–Kir6.2 (not shown). The rate of K+ uptake is increased in mitochondria overexpressing Kir6.2
In order to determine whether mitochondrially expressed Kir6.2 acts as a functional K+ channel, we directly measured K+ uptake into mitochondria using the K+-selective fluorescent indicator PBFI AM (Zoeteweij et al. 1994; Xu et al. 2002). A loading protocol that favoured mitochondrial localization of the dye was applied (Fig. 2A). The rate of K+ flux into mitochondria was determined as the change in PBFI fluorescence ratio from two excitation wavelengths (340/380 nm) after increasing the external K+ concentration. As shown in Fig. 2B (representative recordings), and C (data summary) the rate of K+ influx was significantly higher in mitochondria overexpressing Kir6.2, compared to mitochondria expressing only GFP (n = 20 cells). The measured K+ influx was sensitive to the K+ channel blocker Ba2+. These experiments confirmed that mitochondrial K+ uptake is enhanced in mitochondria overexpressing Kir6.2.
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To investigate the influence of mitochondrial K+ channel overexpression on cellular viability, HL-1 cardiomyocytes and HEK293 cells which stably expressed either mito/GFP–Kir6.2 or mito/GFP were subjected to hypoxia/reoxygenation. The relative increase in LDH release after hypoxia/reoxygenation was less in mito/GFP–Kir6.2 than in mito/GFP transfected cells (162.4 ± 7.4% versus 239.6 ± 15.5% for HL-1 cells and 139.6 ± 9.2% versus 184.7 ± 10.1% for HEK293 cells, n = 14 dishes, P < 0.05), as shown in Fig. 3A. Moreover, addition of 5-hydroxydecanoate (5-HD, 200 µM), a reported mitoKATP inhibitor, during hypoxia/reoxygenation did not significantly change the extent of cellular damage in either group (Fig. 3A). These findings were confirmed by the MTT assay (Fig. 3B). Viability was preserved more effectively in the cells with mitochondria overexpressing Kir6.2 (61.4 ± 2.6% for HL-1 cells and 75.3 ± 3.1% for HEK293 cells), when compared to the cells expressing only mito/GFP (45.7 ± 2.5% for HL-1 cells and 59.4 ± 3.4% for HEK293 cells, n = 14 dishes, P < 0.05). Therefore, the overexpression of Kir6.2 K+ channel in mitochondria protects cells against hypoxic stress. However, this protection is not reversed in the presence of 5-HD.
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To monitor the changes in mitoCa2+ during stress, transfected HEK293 cells were loaded with rhod-2 AM, a mitoCa2+-sensitive fluorescent indicator (Hajnoczky et al. 1995). Representative images showing rhod-2 fluorescence in mito/GFP and mito/GFP–Kir6.2-expressing cells recorded at baseline and during exposure to metabolic inhibitors are shown in Fig. 4A (red). GFP fluorescent signal originating from mitochondria of transfected cells (green) verified the mitochondrial localization of rhod-2. As presented in Fig. 4B, an increase in mitochondrial rhod-2 fluorescence observed during metabolic inhibition in mito/GFP transfected cells was substantially blunted in mito/GFP–Kir6.2 transfected cells (241 ± 24% of control level versus 161 ± 12%, respectively, n = 30 cells). MitoCa2+ accumulation was similarly reduced in cells expressing Kir6.2 lacking GFP tag (myc/mito–Kir6.2), while this effect was not observed when cells were transfected with an inactive form of Kir6.2 (mito/GFP–Kir6.2AAA, n = 30 cells). Addition of 5-HD did not affect significantly the extent of rhod-2 fluorescence changes during stress (Fig. 4C).
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Influx of potassium ions through mitochondrial K+-selective channels causes depolarization of the highly negative mitochondrial membrane potential (Debska et al. 2001; Murata et al. 2001). To assess the effect of mitochondrial Kir6.2 expression on m, we loaded transfected HL-1 and HEK293 cells with TMRE, a cationic fluorescent indicator that accumulates in mitochondria proportionally to m. Figure 5A, upper panel shows that in HEK293 cells transfected with mito/GFP (arrows), TMRE accumulated to a similar extent as in untransfected cells (arrowheads). However, the cells that were successfully transfected with mito/GFP–Kir6.2 (lower panel, arrows) exhibited less TMRE fluorescence in mitochondria under identical conditions, indicating their partially depolarized state. The data are summarized in Fig. 5B. The intensity of mitochondrial TMRE fluorescence in mito/GFP–Kir6.2-expressing cells is significantly lower compared to mito/GFP-expressing cells (1034 ± 43 arbitrary units (au) versus 2875 ± 158 a.u. for HL-1 cells and 894 ± 91 a.u. versus 2854 ± 137 a.u. for HEK293 cells, n
= 30 cells, P < 0.05). However, mitochondrial expression of the dominant negative form of Kir6.2 (mito/GFP–Kir6.2AAA) did not result in significantly reduced TMRE accumulation in mitochondria (n
= 30 cells). It should be noted that change in m is proportional to the log of TMRE fluorescence intensity change (O'Reilly et al. 2003). Treatment with reported mitoKATP modulators pinacidil (100 µM, activator) or glibenclamide (50 µM, inhibitor) did not affect the intensity of TMRE fluorescence in mitochondria expressing mito/GFP–Kir6.2. During metabolic inhibition, a decrease in TMRE fluorescence relative to control conditions was detected in both cell groups, indicating mitochondrial depolarization (Fig. 5D). Interestingly, this decrease was more pronounced in mitochondria overexpressing Kir6.2 compared to those expressing only GFP, even though they initially accumulated less TMRE.
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Discussion |
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Since the initial discovery of K+-selective transport in liver mitochondria (Inoue et al. 1991), K+ conductance has been detected in the mitochondria of different tissues (Paucek et al. 1992; Bajgar et al. 2001; Debska et al. 2002; Dahlem et al. 2004). To date, the existence of at least three different K+ channels has been reported in the inner mitochondrial membrane (mitoKATP, mitoBKCa and mitoKv1.3), and their electrophysiological and pharmacological properties have been studied in various models including mitoplasts (Inoue et al. 1991; Xu et al. 2002; Szabo et al. 2005), proteosomes (Paucek et al. 1992), planar lipid bilayers (Zhang et al. 2001; Nakae et al. 2003), and intact cells (Kohro et al. 2001). Evidence has been provided that modulation of activity of these channels could contribute to the cellular protection against hypoxic injury (Garlid et al. 1997; Liu et al. 1998; Sato et al. 2005), and numerous attempts have been made to decipher their exact role in mitochondrial and cellular pathophysiology. However, the molecular structure and the mechanism of mitochondrial K+ channels' protective action remain a topic of discussion.
In order to investigate the impact of alterations in K+ homeostasis on mitochondrial parameters and cellular viability, we selectively expressed the Kir6.2 K+ channel in the mitochondria of cultured cardiomyocytes and HEK293 cells. Mitochondrial expression was accomplished by the addition of cytochrome c oxidase targeting presequence, which was used previously by others to direct various proteins to mitochondria (Rizzuto et al. 1992). Patch-clamp and Western blot experiments confirmed that no additional K+ current was present in the plasma membrane. The orientation and the mechanism whereby Kir6.2, fused to the hydrophilic GFP, inserts into the inner mitochondrial membrane, is unknown. It may be possible that the connecting loop between the two transmembrane helices is translocated through the membrane utilizing the TIM22 complex (Rehling et al. 2003). The finding that Kir6.2 targeted to mitochondria increases K+ uptake and partially depolarizes m suggests its functional incorporation into the inner mitochondrial membrane, as well as some spontaneous channel opening even under normoxic conditions (John et al. 1998). The increase of K+ flux into mitochondria overexpressing Kir6.2, as assessed with the K+-sensitive fluorescent indicator PBFI AM, demonstrated that the detected changes in the membrane potential could be directly correlated with an enhanced K+ conductance of the mitochondrial membrane. Nevertheless, these changes in m had no detrimental effects on the cellular viability in mito/GFP–Kir6.2-expressing cells. Rather, the cellular tolerance to stress was substantially enhanced, as documented by the improved survival of mito/GFP–Kir6.2 cells exposed to hypoxia/reoxygenation. During the metabolic inhibition, mitochondrial depolarization was more pronounced in the cells expressing mito/GFP–Kir6.2 than in control cells, indicating that Kir6.2 over-expressed in mitochondria retains its ability to act as a metabolic sensor. Additional control experiments in which Kir6.2 was expressed without the attached GFP tag (myc/mito–Kir6.2) confirmed that GFP is not altering the properties of Kir6.2 in the mitochondrial membrane. Furthermore, in order to exclude the non-specific effects of the Kir6.2 protein inserted into the inner mitochondrial membrane, we generated a construct for the expression of Kir6.2 that does not exhibit K+ transport activity (mito/GFP–Kir6.2AAA). This was accomplished by introducing 132GFG134 to AAA amino acid mutations in the K+-selectivity filter of the pore (Supplemental material, Koster et al. 2002; Tong et al. 2006). When mito/GFP–Kir6.2AAA was expressed in HEK293 cells, mitochondrial membrane potential and Ca2+ accumulation during stress did not differ from the cells expressing only mito/GFP. This finding indicates that the effects we observed pertaining to mitochondrial function can indeed be ascribed to K+ influx via mitoKir6.2.
Activation of the putative mitochondrial K+ channels (mitoKATP and mitoBKCa) mediates cellular protection against metabolic stress in a variety of tissues (Bajgar et al. 2001; Oldenburg et al. 2002; Hai et al. 2005), and is considered a central event in the mechanism of ischaemic preconditioning (Liu et al. 1998; Gross & Fryer, 1999; Shintani et al. 2004; Sato et al. 2005). However, the mechanism whereby an increased K+ inflow to mitochondria elicits cellular protection remains controversial. Increased activity of the mitochondrial K+ channels depolarizes m and attenuates mitochondrial Ca2+ accumulation during ischaemia, by decreasing a driving force for the Ca2+ uptake (Ishida et al. 2001; Murata et al. 2001). A reduced Ca2+ uptake would prevent the opening of the mitochondrial permeability transition pore (mPTP), cytochrome c release and apoptosis/necrosis (Crompton, 1999; Halestrap, 1999). Others have suggested that the K+ influx is accompanied by the alkalinization of the matrix, an increase in reactive oxygen species (ROS) and a mild swelling of mitochondria, which are all essential for protection (Kowaltowski et al. 2001; Andrukhiv et al. 2006; Costa et al. 2006). Expansion of the mitochondrial matrix under hypoxic stress could protect critical mitochondrial energetic functions such as fatty acid oxidation, respiration and ATP production (Halestrap, 1989; Dos Santos et al. 2002).
Most of the previous studies utilized various pharmacological activators and inhibitors of mitochondrial K+ channels in order to investigate the link between mitochondrial K+ homeostasis and cellular protection. However, recent findings revealed that many of these agents have additional mitochondrial and cellular targets. This led to the assumption that their mechanism of action may not be based on modulation of K+ fluxes, but could involve different metabolic pathways (Hanley et al. 2002; Das et al. 2003). Our results indicate that increased mitochondrial K+ influx indeed has the potential to protect cell from hypoxic stress. The viability of cardiomyocytes and HEK293 cells that underwent hypoxia/reoxygenation was substantially improved by the mitochondrial overexpression of Kir6.2. Monitoring of mitochondrial Ca2+ during simulated ischaemia revealed that Ca2+ accumulation was both diminished and delayed in these mitochondria. In addition, we also observed an increase in ROS production (Supplemental material), which has been shown to participate in the triggering phase of cardioprotection by preconditioning.
Subunits of the sarcolemmal KATP channel have been found in mitochondria in several studies (Lacza et al. 2003a, 2003b; Singh et al. 2003). Recently, immunogold labelling revealed that both Kir6.1 and Kir6.2, but not SUR, are present in heart mitochondria (Lacza et al. 2003b). This study suggested the possibility that an inwardly rectifying Kir6.x may form the mitoKATP channel. Conversely, an earlier study showed that flavoprotein fluorescence, an indicator of mitochondrial K+ channels' activity, was unaffected after disruption of the Kir6.2 gene in a mouse model (Suzuki et al. 2002). Further, non-targeted expression of Kir6.2 did not result in improved cellular tolerance to stress without addition of the KATP channel opener (Jovanovic et al. 1998). In the present study, we explored the effectiveness of the reported mitoKATP channel modulators pinacidil, glibenclamide and 5-HD on mitochondrially expressed Kir6.2. These drugs did not affect mitoKir6.2-evoked changes in m, cell survival or mitochondrial Ca2+. The absence of effect of the mitoKATP channel regulators in our model could indicate that Kir6.2 in the mitochondria does not resemble the mitoKATP channel. However, this could also be attributed to the absence of a regulatory subunit that is required to complement Kir6.2. Both glibenclamide and pinacidil modulate sarcolemmal KATP channel activity via an interaction with the SUR subunit. Although a SUR-like protein with a smaller molecular mass than SUR has been reported in mitochondria (Lacza et al. 2003b), and mitochondrial ATP-binding cassette protein 1 has been recently associated with an increased resistance against oxidative stress (Ardehali et al. 2005), no specific interaction of these proteins with the K+ channel openers or sulphonylureas has been reported until now. Similarly, the site of 5-HD action is also unknown. The specificity of 5-HD for the mitoKATP channel became questionable due to the fact that it can enter the fatty acid ß-oxidation metabolic pathway in mitochondria (Hanley et al. 2005). Similar non-specific targets have been also reported for pinacidil and glibenclamide (Cook, 1987; Tominaga et al. 1995; Hanley et al. 2002). However, at least HL-1 cardiomyocytes potentially express regulatory proteins involved in mitoKATP channel regulation. Possibly the presence of the GFP group affects the proper assembly of Kir6.2 with the eventual subunits, or alters its sensitivity to different modulators.
While our data do not confirm or rule out the existence of mitoKATP or other mitochondrial K+ channels, this study demonstrates that mitochondrially expressed Kir6.2 acts as a functional channel, and provides direct evidence that an increased K+ flux into mitochondria elicits cellular protection against hypoxia/reoxygenation injury. Heterologous expression studies, possibly combined with deletion/silencing, of other candidate proteins for mitochondrial K+ channels might prove a valuable tool in investigating the function and composition of endogenous channels responsible for cytoprotection, independently from partially non-specific pharmacological drugs.
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