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Departments of
1 Physiology
2 Pharmacology, University of Saskatchewan, Saskatoon, SK, Canada S7N 5E5
3 Department of Biology, Lakehead University, Thunder Bay, Ontario, Canada P7B 5E1
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Abstract |
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(Received 28 August 2005;
accepted after revision 21 September 2005;
first published online 22 September 2005)
Corresponding author R. Wang: Office of Vice President (Research), Lakehead University, 955 Oliver Road, Thunder Bay, Ontario, Canada P7B 5E1. Email: rwang{at}lakeheadu.ca
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Introduction |
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KATP channels are sensitive to changes in intracellular ATP concentrations. Elevation of intracellular ATP level leads to closure of KATP channels in many metabolically active cells. In this way, the KATP channel is a coupling factor to link metabolic activity and membrane excitability. This feature is especially important for pancreatic ß-cells. When circulating glucose level is elevated, glucose influx into pancreatic ß-cells increases and so does ATP production. Consequential closure of KATP channels on plasma membrane depolarizes membrane and opens voltage-dependent calcium channels. The final eventuality of this chain reaction is increased insulin release due to increased intracellular free calcium. There is no doubt that KATP channels in ß-cells are critical in regulation of glucose-induced insulin secretion (Cook et al. 1988; Ashcroft et al. 1989; Ashcroft & Gribble, 1998). However, beyond the regulatory role of glucose, via alteration of intracellular ATP level, on KATP channels, little is known about the existence of other endogenous regulators for KATP channels in pancreatic ß-cells. By analogy to the stimulatory effect of H2S on KATP channels in vascular SMCs, it is reasonable to believe that H2S may be a novel KATP channel opener in pancreatic ß-cells. To date, endogenous production of H2S in the pancreas, effect of H2S on insulin secretion, and interaction of H2S with pancreatic KATP channels have not been determined.
In the present study, effects of H2S on KATP channels in an insulin-secreting insulinoma cell line, INS-1E cells, were examined using the whole-cell and single-channel recording configurations of the patch-clamp technique. Endogenous levels of H2S were either decreased by transfecting INS-1E cells with CSE-siRNA vector or dialysing the cells with a specific inhibitor for H2S-generating enzyme, or increased by overexpressing CSE gene in INS-1E cells. Actual production of endogenous H2S, expression level of H2S-generating enzymes, and insulin secretion in INS-1E cells were determined. Our study characterized KATP channels in INS-1E cells, demonstrated the important regulatory role of H2S on insulin secretion and pancreatic KATP channel activation for the first time, and revealed endogenous enzymatic production and metabolism of H2S in insulin-secreting cells.
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Methods |
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INS-1E cells derived from a rat insulinoma (kindly provided by Dr C. B. Wollheim, Geneva, Switzerland) were grown in a humidified (5% CO2, 95% O2) atmosphere for up to 2 days in Hepes-buffered RPMI-1640 medium (Sigma) supplemented with 10% fetal bovine serum, 1 mM sodium pyruvate, 50 µM 2-mercaptoethanol (Sigma), 100 units ml–1 penicillin, and 100 µg ml–1 streptomycin (Sigma). For patch-clamp study, cultured INS-1E cells were placed in a Petri dish mounted on the stage of an inverted phase contrast microscope (Olympus IX70). For other biochemical and molecular biology assays, INS-1E cells were harvested and centrifuged at 500 g for 10 min after being rinsed twice with PBS solution.
Electrophysiological recording and analysis
Both whole-cell and single-channel recordings of KATP channel currents were performed as previously described (Cook & Hales, 1984; Zhao et al. 2001). Patch pipettes were pulled from borosilicate glass capillaries using a Flaming/Brown micropipette puller (Model P-87, Sutter Instruments). Pipette resistance was 8–12 M
for single-channel recordings and 1–5 M for whole-cell experiments when filled with electrolyte solution. All electrophysiological recordings were performed at room temperature (20–24°C).
Whole-cell recordings were carried out with an Axopatch 200A patch clamp amplifier via a Digidata 1200 (Axon Instruments Inc.) interface, and analysed off-line using pCLAMP software (version 6.02; Axon Instruments Inc.). Test pulses of 600 ms were made with 10 mV increments from –150 to +50 mV. The holding potential was set at –20 mV. I–V relationships were constructed using stable current amplitude at the end of 600 ms test pulses. Pipettes were filled with a solution containing (mM): KCl 105, MgCl2 1.0, CaCl2 1.0, EGTA 10, and Hepes 10 (pH adjusted to 7.3 with KOH). Unless otherwise specified, ATP concentration of the pipette solution was 0.3 mM. The bath solution contained (mM): NaCl 102, KCl 40, CaCl2 1.0, MgCl2 1.2, glucose 4.5, and Hepes 10 (pH adjusted to 7.4 with NaOH). When glucose concentration was increased in some experiments, equimolar NaCl was removed to maintain osmolality of the bath solution. No leakage subtraction was performed to the original recordings and all cells with visible changes in leakage currents during the course of the study were excluded from further analysis.
For single-channel recording, inside-out configuration of the patch-clamp technique (Hamill et al. 1981) was used. Seal resistance was 10 G. Single-channel currents were filtered at 1 kHz (eight-pole Bessel, –3 db) and recorded with 100 µs sampling interval in a gap-free model. Single-channel current records were displayed and analysed using pCLAMP 7.0 software (Axon Instruments Inc.). For each concentration of tested agents, at least 30 s of channel activity was directly recorded on computer hard disk. Open probability (NPo), i.e. the fraction of time when the channels stay open within the total observation period with N representing the number of single channels in one patch (Wang & Wu, 1997) and single-channel conductance were determined from an all-point amplitude histogram using Fetchan and Pstat programs (Axon Instruments Inc.). The pipette solution contained (mM): KCl 140, MgCl2 1.2, EGTA 10 and Hepes 5 (pH adjusted to 7.2 with KOH). Inside-out patches were bathed in a solution containing (mM): KCl 140, MgCl2 0.53, glucose 4.5, ATP 0.3, ADP 0.3 and Hepes 5 (pH adjusted to 7.4 with KOH).
Measurement of endogenous H2S production
H2S production rate was measured as previously described (Stipanuk & Beck, 1982) with modifications, which has been routinely used in our laboratory (Zhao et al. 2001, 2003; Cheng et al. 2004). Briefly, INS-1E cells cultured for 3–7 days were collected and homogenized in 50 mM ice-cold potassium phosphate buffer pH 6.8. The reaction mixture contained (mM): 100 potassium phosphate buffer pH 7.4, 10 L-cysteine, 2 pyridoxal 5'-phosphate, and 10% (w/v) homogenate. Cryovial test tubes (2 ml) were used as the centre wells, each containing 0.5 ml 1% zinc acetate as trapping solution and a filter paper 2 cm x 2.5 cm to increase air: liquid contacting surface. Reaction was performed in a 25 ml Erlenmeyer flask (Pyrex, USA). The flasks containing the reaction mixture and centre wells were flushed with N2 before being sealed with a double layer of Parafilm. Reaction was initiated by transferring the flasks from ice to a 37°C shaking water bath. After incubating at 37°C for 90 min, 0.5 ml of 50% trichloroacetic acid was added to stop the reaction. The flasks were sealed again and incubated at 37°C for another 60 min to ensure a complete trapping of H2S released from the mixture. Contents of the centre wells were then transferred to test tubes, each containing 3.5 ml of water. Subsequently, 0.5 ml of 20 mMN,N-dimethyl-p-phenylenediamine sulphate in 7.2 M HCl was added immediately followed by addition of 0.5 ml 30 mM FeCl3 in 1.2 M HCl. Absorbance of the resulting solution at 670 nm was measured 20 min later with a spectrophotometer (Siegel, 1965). H2S content was calculated against the calibration curve of standard H2S solutions.
Measurement of insulin secretion from INS-1E cells
Native or transfected INS-1E cells were plated into 24-well plates at a density of 5 x 104 cells per well and tested 24–48 h later when cells reached about 80% confluence. Cells were maintained at 37°C for 2 h in glucose-free RPMI 1640 medium, washed and pre-incubated in glucose-free (0 mM) Krebs-Ringer-bicarbonate medium (pH 7.4) containing (mM): 135 NaCl, 3.6 KCl, 5 NaHCO3, 0.5 NaH2PO4, 0.5 MgCl2, 1.5 CaCl2, 10 Hepes and 0.1% BSA. After 30 min pre-incubation, cells were incubated for 30 min at 37°C in the presence of different glucose concentrations. At the end of each incubation period, the medium was collected and centrifuged for 10 min at 2500 g to remove cell debris. The supernatant was immediately stored at –20°C until insulin determination using the rat insulin ELISA kit (Mercodia AB, Sylveniusgatan, Uppsala, Sweden).
Western immunoblotting
Cultured cells were harvested and lysed in a lysis buffer (EDTA 0.5 M; Tris-Cl 1 M, pH 7.4; sucrose 0.3 M; antipain hydrochloride 1 µg ml–1; benzamidine hydrochloride hydrate 1 M; leupeptin hemisulphate 1 µg ml–1; 1,10-phenanthroline monohydrate 1 M; pepstatin A 1 µM; plenylmethylsulphonyl fluoride 0.1 mM, and iodoacetamide 1 mM). Extracts were separated by centrifugation at 14 000 g for 15 min at 4°C. SDS-PAGE and Western blot analysis were performed as previously described (Yang et al. 2004a). Briefly, equal amount of proteins were boiled in 1 x SDS sample buffer (62.5 mM Tris-Cl, pH 6.8, 2% SDS, 10% glycerol, 50 mM DTT, and 0.01% bromophenol blue) and resolved on a 10% SDS-PAGE gel, and transferred onto polyvinylidene chloride (PVDC) nitrocellulose membranes. Dilutions for the primary antibodies were 1: 1000 for CSE, and 1: 5000 for ß-actin. HRP-conjugated secondary antibody was used at 1: 5000. Immunoreactions were visualized by enhanced chemiluminescence (ECL) and exposed to X-ray film (Kodak Scientific Imaging film). Membranes were stripped by incubating in a buffer containing 100 mMß-mercaptoethanol, 2% SDS and 62.5 mM Tris-HCl (pH 6.8).
CSE-siRNA transfection of INS-1E cells
CSE-targeted 21 nucleotide siRNA was designed using a web based siRNA design program (http://www.ambion.com/techlib/misc/siRNA_finder_html) according to the AA-N19 rule (Brummelkamp et al. 2002; Lake & Castellot, 2003). The targeted sequence was localized at a position 192 bases downstream of the start codon of CSE (GenBank Accession No. NM001902). Forward (ggu uau uua ucc ugg gcu g dtdt) and reverse (cag ccc agg cua aau aac c dtdt) RNA strands with two 5' deoxy-thymidine overhangs were commercially synthesized by Ambion (Austin, TX, USA). GADPH-targeted siRNA was also produced for optimizing transfection conditions. Negative control siRNA, a 21 nucleotide RNA duplex with no sequence homology with all known genes, was also purchased from Ambion. Transfection of siRNA into INS-1E cells was achieved using the siPORT lipid transfection agent from Ambion. Briefly, cells were plated overnight to form 60–70% confluent monolayers. CSE siRNA at 30 nM and the transfection reagent complex were added to cells in serum-free medium for 4 h. Fresh normal growth medium was then added and cells were incubated for another 20 h. As a control, negative siRNA was used to transfect INS-1E cells.
Construction of recombinant CSE adenovirus vector and infection of INS-1E cells
A PCR was performed to amplify opening read frame (ORF) of CSE (GenBank accession number AB052882) from reverse-transcribed rat vascular tissue using a set of primers:
5'-CGTCCCAGCATGCAGAAGAA-3' and
5'-CAGTTATTCAGAAGGTCTGGCCC-3'. The amplified ORF of CSE was ligated into PCR2.1 vector (Invitrogen), and the KpnI–XhoI restriction fragment of CSE was subcloned into the KpnI–XhoI sites of the shuttle vector pAdShuttle-CMV (Qbiogene, Inc.), which contains cytomegalovirus promoter/enhancer element and simian virus 40 polyadenylation signals. Positive clone containing CSE ORF insert was sequenced to confirm the accuracy of the inserted CSE sequence. The resultant plasmid was linearized with PmeI and cotransformed with the adenovirus backbone vector pAdeasy-1 into E. coli BJ5183 cells by electroporation. Homologous recombinants containing CSE cDNA were detected by restriction endonuclease digestion and agarose gel electrophoresis. Recombinant CSE adenovirus vector (Ad-CSE) was then transformed into E. coli DH5
cells for large-scale amplification. The PacI-digested E1-deleted replication-deficient Ad-CSE vector was then transfected into mammalian HEK-293 cells using calcium phosphate–DNA precipitates. The recombinant Ad-CSE was expanded, purified and titrated (He et al. 1998). The recombinant adenovirus encoding bacterial ß-galactosidase (Ad-lacZ) derived from the same vector was used as a control. For adenoviral infection, subconfluent INS-1E cells were incubated with Ad-CSE or Ad-lacZ in serum-free media. After 4 h of incubation, media was removed, and cells were incubated in appropriate media for 48 h. The transfection efficiency of adenoviral vector in INS-1E cells was first determined by infecting cells with Ad-lacZ at various multiplicities of infection (MOI). The cells infected with Ad-lacZ were assayed for ß-galactosidase expression by the in situ X-gal staining method (Hirooka & Sakai, 2004). At MOI 
50, > 90% of cells showed nuclear staining for ß-galactosidase. Subsequent experiments were performed at MOI of 50.
Real-time RT-PCR determination of the transcriptional level of CSE
Native untransfected cells, negative siRNA- or CSE siRNA-transfected cells were harvested from 100 mm culture dishes 24 h after transfection. Monolayers were rinsed twice with PBS, and total RNA was collected using Tri Reagent (Molecular Research Center, Cincinnati, OH, USA). Contaminating DNA was avoided using the DNA-free kit (Ambion), and total RNA (2 µg) was reverse-transcribed into cDNA with AMV reverse transcriptase using random hexamer primers according to the manufacturer's protocol (Roche Applied Science, IN, USA). Controls containing no reverse transcriptase were used to safeguard for genomic DNA contamination in each sample.
Real-time PCR was performed in an iCycler iQ apparatus (Bio-Rad, Harcules, CA, USA) associated with the iCycler optical system software (version 3.1) using the SYBR Green PCR Master Mix. All PCRs were performed in a 20 µl volume using 96-well optical grade PCR plates and optical sealing tape. Negative controls for this experiment were samples without a template. Cycling conditions were 95°C for 90 s followed by 38 cycles of 95°C for 10 s and 60°C for 20 s. For quantification, the target gene was normalized to the internal standard gene ß-actin. A standard curve was constructed with a series of dilutions of total RNA (Ambion) transcribed to cDNA using the protocol outlined above to confirm the same amplifying efficiency in PCR. A standard melting curve analysis was performed using a thermal cycling profile that began at 95°C for 1 min, decreased to 55°C for 1 min, and then ramped to 95°C in 1°C increments to confirm the absence of primer dimers. Product size was determined by running PCR products on a 1.8% agarose gel. Relative mRNA quantification was calculated by using the arithmatic formula ‘2–
CT’, where CT is the difference between the threshold cycle of a given target cDNA and an endogenous reference cDNA (Yang et al. 2004a). Thus, this value yields the amount of the target normalized to an endogenous reference.
Chemicals and statistical analysis
H2S-saturated solution (0.09 M) was freshly made by bubbling pure H2S gas (Praxair; Mississauga, Canada) into Krebs' solution at 30°C for 40 min as previously described (Zhao et al. 2001, 2003; Cheng et al. 2004). At 37°C and pH 7.4, the concentration of H2S in solution was relatively stable (Zhao et al. 2003). Data are expressed as mean ±S.E.M. Multiple comparisons were made with one-way ANOVA followed by a post hoc analysis (Tukey test). Statistical significance was set at P < 0.05.
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To investigate whether cultured INS-1E cells produced H2S under different in vivo conditions, INS-1E cells were incubated with either 5 or 20 mM glucose for 24 h. Increase in extracellular glucose concentration from 5 to 20 mM significantly decreased endogenous production of H2S in INS-1E cells by about 46% (Fig. 1A). Similar inhibition of H2S production in INS-1E cells was also observed with 16 mM glucose (not shown). To further examine whether this glucose-mediated endogenous H2S production was regulated by specific enzymatic process, DL-propargylglycine (PPG), a selective inhibitor of CSE, was used. Lysed INS-1E cells were mixed with PPG to facilitate interaction of this inhibitor with cytosolically located CSE, and then H2S production was assayed. It was found that PPG significantly inhibited H2S production in INS-1E cells (Fig. 1A).
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Characterization of KATP channel in INS-1E cells
Gliclazide is a specific blocker of KATP channels in pancreatic ß-cells (Trube et al. 1986; Ashcroft, 2000). Gliclazide (1 µM) decreased significantly whole-cell KATP currents in INS-1E cells from –1049.9 ± 115 to 522.8 ± 88 pA at –100 mV (n= 5, P < 0.05). Representative results of the effect of gliclazide on whole-cell KATP channels are shown in Fig. 2A. Diazoxide is a potent opener of KATP channels in pancreatic ß-cells (Sturgess et al. 1988; D'hahan et al. 1999). Whole-cell KATP channels in INS-1E cells were significantly stimulated by diazoxide (Fig. 2B). Whole-cell KATP channels in INS-1E cells were also characterized by their sensitivity to intracellular ATP. KATP channel currents were 82.9 ± 8.6 pA pF–1 (–120 mV) with 0.3 mM ATP in the pipette solution (n= 6). When the ATP concentration was increased to 3 mM, KATP channel currents were reduced to 29.7 ± 5.6 pA pF–1 (–120 mV) (n= 5, P < 0.05 versus 0.3 mM ATP in the pipette solution).
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KATP channels in INS-1E cells were also sensitive to glucose stimulation. After glucose concentration of the bath solution was changed from 5 mM to 16 mM, whole-cell KATP currents were significantly reduced (Fig. 4A). With 5 mM glucose in the bath solution, H2S at 100 µM had no effect on whole-cell KATP currents in INS-1E cells (n= 5, P > 0.05). In the presence of 16 mM glucose, application of H2S (100 µM) to INS-1E cells significantly increased KATP currents (Fig. 4B). Application of DL-dithothreitol (DTT) (3 mM) to INS-1E cells for 5 min did not significantly change KATP channel currents in INS-1E cells (79 ± 2.69 versus 86 ± 2.68 pA pF–1 at –100 mV, n= 5, P > 0.05). A lack of effect of DTT on KATP channels has also been reported previously in pancreatic ß-cells (Islam et al. 1993; Krippeit-Drews et al. 1994). Since DTT is a reducing reagent, our result suggests that the stimulatory effect of H2S on KATP channels in INS-1E cells is unlikely to be mediated by a general reducing effect.
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Discussion |
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We recorded a 78 pS KATP channel in INS-1E cells. This single-channel conductance is typical of KATP channels reported in pancreatic ß-cells (Ashcroft & Gribble, 1998). Other characteristics of KATP channels, including burst opening, ATP sensitivity, glucose sensitivity, voltage insensitivity and gliclazide sensitivity are all similar to those described in native pancreatic ß-cells (Mukai et al. 1998). Moreover, 0.3 mM Mg-ATP was required for sustaining KATP channel currents in our study, a property shared by SUR1/KIR6.2 type of KATP channels (Inagaki et al. 1995; Gribble et al. 1997). Sulphonylureas such as gliclazide stimulate insulin secretion by closing KATP channels (Sturgess et al. 1985; Trube et al. 1986; Ashcroft, 2000). Potassium channel openers such as diazoxide inhibit insulin secretion by opening KATP channels (Trube et al. 1986; Dunne et al. 1987; Sturgess et al. 1988; Minami et al. 2003). In our study, diazoxide significantly increased, and gliclazide inhibited, KATP channel activity in INS-1E cells. KATP channels in INS-1E cells were also inhibited by high glucose concentration in the bath solution (16 mM) or ATP (3 mM) in the pipette solution. These features are hallmarks of KATP channels in insulin-secreting cells (Cook & Hales, 1984; Ashcroft & Rorseman, 1989; Ashcroft & Gribble, 1998). Insulin secretion and KATP channel functionality in this way respond to changes in glucose levels. These pharmacological and biophysical properties indicate that KATP channels in INS-1E cells share the same characteristics with those in pancreatic ß-cells (Ashcroft & Gribble, 1998; Mukai et al. 1998).
INS-1E cells are derived from an insulinoma pancreatic ß-cell line (Janjic et al. 1999; Merglen et al. 2004). These cells exhibit stable differentiated ß-cell phenotype, and secrete insulin in response to glucose and non-nutrient secretagogues via the stimulation of KATP channels and a minor amplifying pathway (Merglen et al. 2004). Significant amounts of H2S were produced by INS-1E cells. We have achieved a partial knockdown of the CSE gene in INS-1E cells using the CSE-siRNA technique. Since this partial knockdown significantly reduced the production of H2S, it is believed that CSE is the main enzyme for the production of H2S in INS-1E cells. More importantly, our study demonstrated that this endogenous H2S production in INS-1E cells was mediated by a variance in glucose concentrations, thus providing physiological regulatory mechanisms for H2S production. Functional correlation of H2S levels in INS-1E cells is realized by insulin secretion from these cells. By reducing endogenous H2S production in INS-1E cells, high glucose also stimulated insulin secretion. Furthermore, over-expression of the CSE gene in INS-1E cells via Ad-CSE infection significantly increased endogenous H2S production, thus inhibiting high glucose-stimulated insulin secretion.
In the present study, we demonstrated for the first time that H2S activated KATP channels in INS-1E cells. Without manipulating the endogenous H2S level, exogenous H2S at concentrations equal to or lower than 100 µM had no effect on whole-cell KATP channels in INS-1E cells. This phenomenon might be explained as KATP channels in INS-1E cells were desensitized to endogenous H2S at resting conditions. When INS-1E cells were incubated with a high concentration of glucose (16–20 mM), these cells became sensitive to exogenous H2S that significantly increased KATP channel activity at 100 µM. This sensitizing effect of high glucose concentration on KATP channels could be linked to a high glucose-induced decrease in endogenous H2S production. This hypothesis was further verified by directly inhibiting CSE activity in INS-1E cells. When PPG (1–5 mM) was used to dialyse INS-1E cells to inhibit CSE and endogenous production of H2S, whole-cell KATP channel currents were greatly increased by exogenously applied H2S in a concentration-dependent manner. Another line of evidence supporting conditioning of KATP channels in INS-1E cells by endogenous H2S was derived from single-channel recording studies. Exogenous H2S significantly increased open probability of single KATP channels in inside-out membrane patches of INS-1E cells. With this cell-free recording configuration, substrates and enzymes for endogenous H2S production are eliminated. Therefore, it is highly possible that KATP channels in insulin-secreting cells have been desensitized by a high level of endogenous H2S at resting conditions. Removal or reduction of endogenous H2S would re-sensitize KATP channels so that these channels regain their sensitive response to H2S. We propose that the endogenous H2S level has one switch for turning on or off KATP channels in insulin-secreting cells, whereas the glucose level regulates endogenous H2S production. Under physiological conditions with low extracellular glucose (
5 mM), endogenous H2S level is high, which would keep KATP channels mostly in their open state, thus hyperpolarizing the membrane of insulin-secreting cells. This will result in low activity of voltage-dependent calcium channels and low secretion of insulin from insulin-secreting cells. When the glucose concentration of plasma is elevated, endogenous H2S production in insulin-secreting cells is decreased. Consequent closure of KATP channels leads to increased insulin secretion. In our previous studies, it has been shown that the vasorelaxant effect of H2S was not mediated by any known second messengers, including cGMP, cAMP and PKC pathways (Zhao et al. 2001, 2003; Zhao & Wang, 2002). In the present study, we showed that H2S directly activates KATP channels in cell-free inside-out membrane patches. We also showed that ATP sensitivity of KATP channels was not changed by H2S. Taken together, these observations suggest that the interaction of H2S and KATP channels is not mediated by cytosolic second messengers. As a reducing agent, H2S may reduce selective cysteine residues of KATP channel protein, altering its functional status. However, application of a classical reducing agent (DTT) to INS-1E cells did not replicate the excitatory effect of H2S on KATP channels, suggesting that other mechanisms should be sought to explain the interaction of H2S with the KATP channel complex in a tissue-/cell type-specific manner.
In summary, KATP channels in insulin-secreting INS-1E cells share many common features with their counterparts in native pancreatic ß-cells. H2S increased the activity of these KATP channels by increasing single-channel open probability, but not single-channel conductance. An endogenous high level of H2S in INS-1E cells sets up the tune for KATP channel activity and thus insulin-secreting level at resting conditions. An increase in extracellular glucose concentration lowers endogenous H2S level. This will have two effects. Firstly, activity of KATP channels in INS-1E cells will be significantly reduced so that insulin secretion will be increased. Secondly, KATP channels in INS-1E cells will be partially closed and re-sensitized to H2S. Subsequent changes in the endogenous H2S level would exert a much greater effect on the functional status of KATP channels under these conditions. Interaction among H2S, glucose, and KATP channels in insulin-secreting cells may constitute an important and novel mechanism for the fine control of insulin secretion from pancreatic ß-cells, which is initially triggered by changes in glucose concentration, under physiological and different pathophysiological conditions.
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References |
|---|
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TopAbstractIntroductionMethodsResultsDiscussionReferences |
|---|
Ashcroft FM (1996). Mechanism of the glycaemic effects of sulfonylureas. Horm Metab Res 28, 456–463.[Medline]
Ashcroft SJH (2000). The ß-cell KATP channel. J Membr Bio 176, 187–206.[CrossRef][Medline]
Ashcroft FM & Gribble FM (1998). Correlating structure and function in ATP-sensitive K+ channels. Trends Neurosci 21, 288–294.[CrossRef][Medline]
Ashcroft FM, Rorsman P & Trube G (1989). Single calcium channel activity in mouse pancreatic beta-cells. Ann N Y Acad Sci 560, 410–412.[CrossRef][Medline]
Brummelkamp TR, Bernards R & Agami R (2002). A system for stable expression of short interfering RNAs in mammalian cells. Science 296, 550–553.
Cheng Y, Ndisang JF, Tang G, Cao K & Wang R (2004). Hydrogen sulfide-induced relaxation of resistance mesenteric artery beds of rats. Am J Physiol Heart Circ Physiol 287, H2316–H2323.
Cook DL & Hales CN (1984). Intracellular ATP directly blocks K+ channels in pancreatic ß-cells. Nature 311, 271–273.[CrossRef][Medline]
Cook DL, Satin LS, Ashford ML & Hales CN (1988). ATP-sensitive K+ channels in pancreatic beta-cells. Spare-channel hypothesis. Diabetes 37, 495–498.
D'hahan N, Jacquet H, Moreau C, Catty P & Vivaudou M (1999). A transmembrane domain of the sulfonylurea receptor mediates activation of ATP-sensitive K+ channels by K+ channel openers. Mol Pharmacol 56, 308–315.
Dunne MJ, Illot MC & Peterson OH (1987). Interaction of diazoxide, tolbutamide and ATP4- on nucleotide-dependent K+ channels in an insulin-secreting cell line. J Membr Biol 99, 215–224.[CrossRef][Medline]
Geng B, Yang J, Qi Y, Zhao J, Du Pang Y, J & Tang C (2004). H2S generated by heart in rat and its effects on cardiac function. Biochem Biophys Res Commun 313, 362–368.[CrossRef][Medline]
Gribble FM, Ashfield R, Ammala C & Ashcroft FM (1997). Properties of cloned ATP-sensitive K+ currents expressed in Xenopus oocytes. J Physiol 498, 87–98.[CrossRef][Medline]
Hamill OP, Marty A, Neher E, Sakmann B & Sigworth FJ (1981). Improved patch-clamp techniques for high-resolution current recording from cells and cell-free membrane patches. Pflugers Arch 391, 85–100.[CrossRef][Medline]
He TC, Zhou S, da Costa LT, Yu J, Kinzler KW & Vogelstein B (1998). A simplified system for generating recombinant adenoviruses. Proc Natl Acad Sci U S A 95, 2509–2514.
Hirooka Y & Sakai K (2004). Adenovirus-mediated nitric oxide synthase gene transfer into the nucleus tractus solitarius in conscious rats. Meth Mol Biol 279, 187–200.[Medline]
Inagaki N, Tsuura Y, Namba N, Masuda K, Gonoi T, Horie M et al. (1995). Cloning and functional characterization of a novel ATP-sensitive potassium channel ubiquitously expressed in rat tissues, including pancreatic islets, pituitary, skeletal muscle, and heart. J Biol Chem 270, 5691–5694.
Islam MS, Berggren PO & Larsson O (1993). Sulfhydryl oxidation induces rapid and reversible closure of the ATP-regulated K+ channel in the pancreatic beta-cell. FEBS Lett 319, 128–132.[CrossRef][Medline]
Janjic D, Maechler P, Sekine N, Bartley C, Annen AS & Wollheim CB (1999). Free radical modulation of insulin in INS-1 cells exposed to alloxan. Biochem Pharmacol 57, 639–648.[CrossRef][Medline]
Krippeit-Drews P, Britsch S, Lang F & Drews G (1994). Effects of SH-group reagents on Ca2+ and K+ channel currents of pancreatic ß-cells. Biochem Biophys Res Commun 200, 860–866.[CrossRef][Medline]
Lake AC & Castellot JJ (2003). CCN5 modulates the antiproliferative effect of heparin and regulates cell motility in vascular smooth muscle cells. Cell Commun Signal 1, 5.[CrossRef][Medline]
Lorenz E, Alekseev AE, Krapivinsky GB, Carrasco AJ, Clapham DE & Terzic A (1998). Evidence for direct physical association between a K+ channel (Kir6.2) and an ATP binding cassette protein (SUR1) which affects cellular distribution and kinetic behavior of an ATP-sensitive K+ channel. Mol Cell Bio 18, 1652–1659.
Merglen A, Theander S, Rubi B, Chaffard G, Wollheim CB & Maechler P (2004). Glucose sensitivity and metabolism secretion coupling studied during two-year continuous culture in INS-1E insulinoma cells. Endocrinology 145, 667.
Minami K, Morita M, Saraya A, Yano H, Terauchi Y, Miki T et al. (2003). ATP-sensitive K+ channel-mediated glucose uptake is independent of IRS-1/phosphatidylinositol 3-kinase signaling. Am J Physiol Endocrinol Metab 285, E1289–E1296.
Mukai E, Ishida H, Kato S, Tsuura Y, Fujimoto S, Ishida-Takahashi A et al. (1998). Metabolic inhibition impairs ATP-sensitive K+ channel block by sulfonylurea in pancreatic beta-cells. Am J Physiol 274, E38–E44.[Medline]
Saskura H, Ammala C, Smith PA, Gribble FM & Ashcroft FM (1995). Cloning and functional exoression of the cDNA encoding a novel ATP-sensitive potassium channel subunit expressed in pancreatic beta-cells heart and skeletal muscle. FEBS Lett 377, 33–44.
Siegel LM (1965). A direct microdetermination for sulfide. Anal Biochem 11, 126–132.[CrossRef][Medline]
Stipanuk MH & Beck PW (1982). Characterization of the enzymic capacity for cysteine desulphhydration in liver and kidney of the rat. Biochem J 206, 267–277.[Medline]
Sturgess NC, Ashford ML, Cook DL & Hales CN (1985). The sulfonylurea receptor may be an ATP-sensitive potassium channel. Lancet 2 (8453), 474–475.[Medline]
Sturgess NC, Kozlowski RZ, Carrington CA, Hales CN & Ashford ML (1988). Effects of sulphonylureas and diazoxide on insulin secretion and nucleotide-sensitive channels in an insulin-secreting cell line. Br J Pharmacol 95, 83–94.[Medline]
Trube G, Rorsman P & Ohno-Shosaku T (1986). Opposite effects of tolbutamide and diazoxide on the ATP-dependent K+ channel in mouse pancreatic beta-cells. Pflugers Arch 407, 493–499.[CrossRef][Medline]
Wang R & Wu L (1997). The chemical modification of KCa channels by carbon monoxide in vascular smooth muscle cells. J Biol Chem 272, 8222–8226.
Wang R (2002). Two's company, three's a crowd – can H2S be the third endogenous gaseous transmitter? FASEB J 16, 1792–1798.
William CC & Odle CC (2003). ATP-sensitive K+ channels of vascular smooth muscle cells. J Vascular Eletrophysiol 14, 94–103.
Yang G, Cao K & Wang R (2004a). Cystathionine 
-lyase overexpression inhibits cell proliferation via a H2S-dependent modulation of ERK1/2 phosphorylation and p21Cip/WAK–1. J Biol Chem 279, 49199–49205.
Yang G, Sun X & Wang R (2004b). Hydrogen sulfide-induced apoptosis of human aorta smooth muscle cells via the activation of MAP kinases and caspase-3. FASEB J 18, 1782–1784.
Yokoshiki H, Sunigawa M, Seki T & Sperelakis N (1998). ATP-sensitive K+ in pancreatic, cardiac, and vascular smooth muscle cells. Am J Physiol 274, C25–C37.[Medline]
Zhao W, Ndisang JF & Wang R (2003). Modulation of endogenous production of H2S in rat tissues. Can J Physiol Pharmacol 81, 848–853.[CrossRef][Medline]
Zhao W & Wang R (2002). H2S-induced vasorelaxation and underlying cellular and molecular mechanisms. Am J Physiol 283, H474–H480.
Zhao W, Zhang J, Lu Y & Wang R (2001). The vasorelaxant effect of H2S as a novel endogenous gaseous KATP channel opener. EMBO J 20, 6008–6016.[CrossRef][Medline]
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