Zn++-DEPENDENT REDOX SWITCH IN THE INTRACELLULAR T1-T1 INTERFACE OF A Kv CHANNEL

Guangyu Wang1, Candace Strang2, Paul J. Pfaffinger2 and Manuel Covarrubias1 From the 1Department of Pathology, Anatomy and Cell Biology,

Jefferson Medical College of Thomas Jefferson University, Philadelphia, PA 19107 2Division of Neuroscience, Baylor College of Medicine, Houston, TX 77030

Running Title: Functional role of the Kv4 channel T1 Zn++ site 1Address correspondence to: Manuel Covarrubias, 1020 Locust Street, JAH245, Philadelphia, PA 19107; Tel. (215) 503-4341; Fax (215) 923-2218; E-Mail; Manuel.Covarrubias@jefferson.edu

The thiol-based redox regulation of proteins plays a central role in cellular signaling. Here, we investigated the redox regulation at the Zn++ binding site (HX5CX20CC) in the intracellular T1-T1 inter-subunit interface of a Kv4 channel. This site undergoes conformational changes coupled to voltage-dependent gating, which may be sensitive to oxidative stress. The main results show that internally applied nitric oxide (NO) inhibits channel activity profoundly. This inhibition is reversed by reduced glutathione and suppressed by intracellular Zn++; and at least two Zn++ site cysteines are required to observe the NO-induced inhibition (C110 from one subunit and C132 from the neighboring subunit). Biochemical evidence suggests strongly that NO induces a disulfide bridge between C110 and C132 in intact cells. Finally, further mutational studies suggest that intra-subunit Zn++ coordination involving H104, C131 and C132 protects against the formation of the inhibitory disulfide bond. We propose that the interfacial T1 Zn++ site of Kv4 channels acts as a Zn++-dependent redox switch that may regulate the activity of neuronal and cardiac A-type K+ currents under physiological and pathological conditions.

The highly-conserved inter-subunit Zn++ binding motif (HX5CX20CC) in the intracellular T1 domain differentiates voltage-gated K+ channels such as Kv2, Kv3 and Kv4 from the Shaker Kv1 channels (1,2). Zn++ is coordinated by a cysteine from one subunit and a histidine along with two cysteines from the neighboring subunit (Fig.1). In the absence of β-subunits, Zn++ binding to the T1 site in non-Shaker K+

channels is thought to stabilize the tetrameric structure of the channels (3,4). However, co-expression of Kv4 channels with KChIPs (Kv4-specific β-subunits) overrides the essential role of Zn++ binding in subunit assembly (5,6) while KChIP1 binds to the Kv4-T1 domain without altering the structure of the Zn++ site and the T1-T1 interface (7,8). Furthermore, mutant Kv4 channels lacking the Zn++ site retain nearly normal gating when co-expressed with auxiliary subunits KChIP1 and DPPX-S (5); and our previous studies showed that the cysteines in the Kv4-T1 Zn++ site are unexpectedly accessible to thiol-specific reagents and that the T1-T1 interface at this location is dynamic and functionally coupled to voltage-dependent gating (5,9). Although it is clear that this interface is not the gate that opens the channel, rearrangements in and around the interfacial Zn++ site may regulate the conformational changes required for the opening of the intracellular activation gate (9). This scenario is structurally plausible because just above the Zn++ site, the T1 domain is linked to the voltage sensing domain via the T1-S1 linker; and the C-terminal cytoplasmic region immediately distal to the S6-tail, which may serve as the actual activation gate, may also interact with the T1-T1 interface (10). The intriguing conservation of an apparently non-essential Zn++ binding site at this gating regulatory domain suggests that it may play a signaling function rather than a purely structural role. In support of this hypothesis, reactive cysteines involved in Zn++ binding in proteins are often identified as critical components in redox signaling (11-14). To test this hypothesis, we asked the following questions: 1) Is the T1 Zn++ site a target of nitrosative and oxidative regulation? 2) Are the

http://www.jbc.org/cgi/doi/10.1074/jbc.M609182200The latest version is at JBC Papers in Press. Published on March 1, 2007 as Manuscript M609182200

Copyright 2007 by The American Society for Biochemistry and Molecular Biology, Inc.

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

2

Zn++ coordinating residues in the T1 domain responsible for the redox regulation? 3) What is the role of Zn++? 4) What interactions are involved and what is the mechanism of the regulation? To answer these questions, we investigated the modulation of a heterologously expressed Kv4.1 channel by nitric oxide (NO) and intracellular Zn++ and the underlying molecular mechanism. This modulation is significant because many studies support the physiologically relevant relationship between NO and Zn++ homeostasis in excitable tissues (12,14-17). Particularly, various studies have reported redox modulation of Kv4-related A-type currents in neurons and muscle (15,18-22). However, the underlying molecular mechanisms have remained unsolved. This study strongly suggests that the functionally active T1-T1 intersubunit interface of Kv4 channels is a Zn++-dependent redox switch, which may play a central modulatory role in excitable tissues.

EXPERIMENTAL PROCEDURES Chemicals and Reagents H2O2 (30% w/v), DL-dithiotreitol (DTT), CuSO4, ZnCl2, 1,10 o-phenanthroline, S-nitroso-N-acetylpenicill-amine (SNAP) and glutathione (GSH) were obtained from Sigma-Aldrich Corp. (St. Louis, MO). 1,10 o-phenanthroline was solubilized in ethanol. (NOC-9, MAHMA/NO, (Z)-1 {N-Methyl-N-[6-(N-methylammoniohexyl) amino]} diazen-1-ium-1, 2-diolate] (MAHMA-NONOate) and sodium 2-(N,N-dimethyl-amino)-diazenolate-2-oxide (DEA-NONOate) were purchased from Axxora Bio-sciences, San Diego, CA. Methanethiosul-fonate (MTS) reagents (2-trimethylammonium-ethyl-methanethiosulfonate bromide, MTSET and 2-aminoethyl methanethiosulfonate hydro-bromide, MTSEA and methyl methan-ethiolsulfonate, MMTS) were purchased from Toronto Research Chemicals (North York, Ontario, Canada) and stored in a desiccator at -20oC. Tetrakis-(2-pyridylmethyl ethylene-diamide (TPEN) was purchased from Molecular Devices (Eugene, OR). As reported by the manufacturer, TPEN has an apparent binding affinity for Zn++ of the order of 3×10-16 M. All working solutions of DTT, GSH, 1,10

o-phenanthroline, MAHMA-NONOate, SNAP, DEA-NONOate and MTS reagents were made just before use. Molecular Biology and Heterologous Expression Kv4.1 (mouse) and DPPX-S were maintained in pBluescript II KS and pSG5 (Stratagene), respectively. KChIP1 was maintained in a modified pBluescript vector, pBJ/KSM. KChIP1 and DPPX-S are gifts from M. Bowlby (Wyeth-Ayerst Research, Princeton, NJ) and B. Rudy (New York University, New York, NY), respectively. All mutants were produced by using the QuickChangeTM site-directed mutagenesis kit from Stratagene and confirmed by automated sequencing (Nucleic Acid Facility of the Kimmel Cancer Institute, Thomas Jefferson University). The capped cRNAs for Xenopus oocyte expression were synthesized by using the in vitro transcription kit, Message Machine (Ambion, Inc. Austin TX). For all the experiments, we co-expressed Kv4.1 WT and mutants with KChIP1 and DPPX-S. The expression of the Kv4.1 ternary complex was necessary because mutations in the putative Zn2+ site yielded non-functional channels or inhibited expression profoundly. Previous studies showed that the apparently lethal phenotype of Zn2+ site mutants can be corrected by coexpression of the channels with KChIPs (5,6); and we have found that DPPX-S boosts the expression of the channels even further (5) which made possible the recordings from inside-out macropatches. Electrophysiology Inside-out patch-clamp recordings were made using asymmetrical KCl solution. Patch electrodes contained (mM) 96 NaCl, 2 KCl, 1.8 CaCl2, 1 MgCl2 and 5 Hepes (pH 7.4, adjusted with NaOH) and the bath solution consisted of (mM) 98 KCl, 0.5 MgCl2, 1 EGTA, 10 HEPES (pH 7.2, adjusted with KOH). The internal solution containing ZnCl2 had no EGTA. Passive leak and capacitive transients from macropatch currents were substracted on-line by using a P/4 procedure. The currents were filtered at 1.5-5 kHz. All the experiments were carried out at room temperature (22±1oC) and reagents were applied to the intracellular side of the patches. MAHMA-NONOate and DEA-NONOate were prepared just after forming the inside-out patch

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

3

and establishing a stable current level. The powder was promptly solubilized in 10 mM NaOH at 100 mM. Several doses of stock solution were immediately stored below –70 °C. The final working solution at 0.1 mM was applied to the intracellular side of the channel. At room temperature, these two reagents generate 2NO with a half-life of ~3 min and ~15 min at pH 7.4, respectively (23). For disulfide cross-linking experiments, we used a mild oxidizing solution containing 50 µM CuSO4 and 200 µM 1,10 o-phenanthroline (Cu/P) (24). To promote the formation of the disulfide bond between a cysteine pair in the T1-T1 interface, the cytoplasmic side of the inside-out patches was treated for ~5 min with fresh Cu/P. After Cu/P washout, 20 mM DTT was used to reduce the disulfide bond (5) or 400 µM MTSET was employed to test for the presence of free thiolate groups. Protein Biochemistry The Kv4.2-T1 domain with a poly-His tag at the NH2-terminus was expressed in bacteria, and purified by a standard Ni++ column protocol as described previously (4). The T1 protein was in a Tris buffered saline solution (TBS) of 50 mM Tris - HCl, pH 7.4 and 150 mM NaCl. The protein was incubated with 2 mM GSH for at least ~20 minutes at room temperature. After reaction with GSH, the sample was prepared either for atomic absorption measurements or for FPLC analysis as described previously (3). Atomic absorption spectroscopy was performed on a Perkin-Elmer AAAnalyzer 600 in the Chemistry Department of Rice University (Houston, TX). Zn++ absorption was read at 213.9 nm with a slit width set at 0.7 nm. Commercially available Zn++ standards to quantify the Zn++ content in dilute nitric acid were used to make a standard curve. Spiked Zn++ standards in the phosphate buffered saline background were also prepared to calibrate the standard curve for the protein samples. No difference was found between the diluted nitric acid and the phosphate buffered protein background matrix. Protein concentration was determined by absorbance at 280 nm, and corrected for light scattering, if necessary with an extinction coefficient of 1 A280 unit - ml/ mg protein.

In order to confirm the formation of the disulfide bond across the T1-T1 interface, T293 cells were grown in 6 well plates (35 mm wells) in Opti-MEM plus 10% FBS. Cells at ~70% confluence were transfected with CMV promoter plasmids for Kv4.1 constructs plus KChIP3 and enhanced-green-fluorescent-protein (EGFP) (1 µg: 1 µg: 0.5 µg) using Lipofectamine 2000 (Invitrogen, Carlsbad, CA). Thirty-six hours after transfection, cells were placed in Dulbecco's- phosphate-buffered-saline with divalents (DPBS, Gibco- Invitrogen, Carlsbad, CA) at room temperature. For NO treatment, 1 ml DPBS solution with 1 mM MAHMA-NONOate was added twice to the bath for 10 min each. The solution was removed and cells were solubilized in 100 µl SDS sample buffer. Half of each sample was treated with 200 mM DTT prior to running on a 6% SDS-PAGE gel. Gels were transferred to Immobilon (Millipore, Billerica, MA) and western blotted with anti-Kv4.1 (Alomone Labs, Jerusalem, Israel) at 1:400 and detected with Goat-anti-Rabbit-HRP (Pierce, Rockford, Il) at 1:5000. Exposures were 10 sec in Pico ECL (Pierce) on BIOMAX MR film (Kodak, New Haven, CT). Data Acquisition and Analysis Voltage-clamp protocols and data acquisition were controlled by a Pentium-4 class desktop computer interfaced to a 12 bit A/D converter (Digidata 1200) or a 16 bit A/D converter (Digidata 1322) and driven by Clampex 8.0 or 9.0 (Axon Instruments). Clampfit 8.0 or 9.0 (Axon Instruments) and Origin 7.0 (Origin Lab Inc.) were used for data reduction and analysis. The time courses of peak current inhibition were evaluated quantitatively by assuming exponential decays to estimate the time constants and fractional currents at steady-state. Data from at least three patches for each measurement are presented as mean ± SEM. The one–way ANOVA test was used to evaluate statistically significant differences between two groups of data.

RESULTS

Zn++ site cysteines are major targets of nitrosative modulation in a Kv4 channel

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

4

complex Upon application of the fast NO donor MAHMA-NONOate (100 µM) to the cytoplasmic side of inside-out patches of Xenopus oocyte expressing the ternary wild-type (WT) Kv4 channel complex (Kv4.1+KChIP1+DPPX-S) (Experimental Procedures), the outward K+ currents were inhibited quickly and profoundly (Fig. 2A-C and 6). The time course of this inhibition was approximately exponential with the following best-fit rate constant (ko = 1/τ) and % inhibition at steady-state (I%): 0.040 ± 0.006 s-1 and 63 ± 4 %, respectively. The inhibition was similarly fast (0.042 ± 0.001 s-1) but more profound (85 ± 2%) when all intracellular cysteines but the three in the T1 Zn++ site (C110, C131 and C132) were mutated to alanines in the C11xA mutant (Fig. 2D-F; and 6). DEA-NONOate, a slower NO donor (Experimental Procedures) produced a similar result (data not shown). In sharp contrast, the inhibition was greatly suppressed when the three Zn++ site cysteines were mutated to alanines in the C3xA mutant (Fig. 2G-I and 6) or when all fourteen internal cysteines were mutated to alanines in a C14xA mutant (“Cys-less”; Fig. 3A-B and 6). Both the ko and I% are reduced ~3-fold relative to values from the C11xA mutant. The residual slow response may arise from nonspecific nitrosation or oxidation of non-cysteine residues (e.g., Met, Ser and Tyr) in the Kv4 channel complex (25) because gating of the C14xA mutant co-expressed with KChIP1 and DPPX-S is not affected by thiol-specific reagents (5). In this study, this small and slow cysteine-independent response was not investigated further, and we regard it as background inhibition. The specific inhibition was not caused by a by-product of MAHMA-NONOate or DEA-NONOate decomposition upon NO release because washout did not reverse it and application of the exhausted NO-donor had no effect on the Kv4 current (data not shown). Once released from its donor, NO is highly labile; therefore, internally-applied H2O2, which is relatively stable, was also used in several experiments as a control reagent to verify the functional role of the cysteines in the T1 Zn++ site. Under identical conditions, high concentrations of H2O2 yielded results that were qualitatively similar to those obtained with NO: the rate and

degree of inhibition of the Kv4 channel complex depended sharply on the presence of cysteines in the T1 Zn++ site (Fig. 3C and 6; Supplemental Information, Fig. 1S). Lower concentrations of H2O2 were also attempted; however, the slow rate to the inhibition under these conditions made these experiments generally impractical for quantitative analysis due to the limited lifetime of the inside-out patches. The role of external cysteines in these responses is unlikely because, in agreement with previous studies, the external application of H2O2 had no effect on the Kv4 current (Fig. 3C) (26,27). C110 plays a central role in redox modulation of the Kv4 channel complex To determine which Zn++ site cysteine in the T1-T1 interface may underlie the nitrosative modulation, we probed two additional mutants: C12xA-a and C13xA. The former has two internal cysteines (A110, C131 and C132) from one side of the interface (Fig. 3D), and the latter has one internal cysteine only (C110, A131 and A132) from the other side of the interface (Fig. 3G). Thus, no disulfide bridges can be formed across the interface upon oxidation. Like mutants C13xA and C14xA, the C12xA-a mutant only exhibited a background response when MAHMA-NONOate or H2O2 were applied internally (Fig. 3E-F and 6); but the C13xA mutant, which carries C110 only was inhibited rapidly and more severely by MAHMA-NONOate (ko = 0.025 ± 0.004 s-1 and I% = 70 ± 1 %) or H2O2 (ko = 0.010 ± 0.001 s-1 and I% = 67 ± 4 %) than C12xA-a (p < 0.005; Fig. 3H-I and 6). Any additional sensitivity of the C12xA-a mutant could not be uncovered even by chelating any possible protective Zn++ with 20 µM TPEN, a high affinity Zn++ chelator (Fig. 3F). These results suggest that C110 plays a unique and critical role in redox modulation of the Kv4 channel complex. Supporting this conclusion, C110 was protected by pre-forming a thiol-specific adduct with MMTS (a non-polar methane thiosulfonate reagent) in the C13xA mutant (Experimental Procedures). This treatment reduced the subsequent responses to MAHMA-NONOate and H2O2 to levels that were close to background (Fig. 3H-I).

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

5

The combined presence of C110 and H104 is sufficient to observe nitrosative modulation of the Kv4 channel complex The above findings were surprising because modification of any cysteine in the Kv4.1-T1 Zn++ site by MTSET induces profound inhibition of the channel, which may result from steric and/or electrostatic interactions in the T1-T1 interface (5). The selective inhibition of the C13xA mutant (C110 only) by nitrosation and oxidation suggests that modification of any individual Zn++ site cysteine is not always sufficient to produce the robust inhibition; therefore, it is necessary to consider more specific interactions that require a particular location and orientation of the groups involved. In the C13xA mutant, two Zn++–coordinating residues C110 and H104 are located in vicinal subunits across the T1-T1 interface (Fig. 4A). Although the thiol group (-SH) of C110 cannot form a strong H-bond with H104, C110 that is oxidized by NO or H2O2 could provide a strong proton donor (S-NO or S-OH) or acceptor (S-NO, SO2

- or SO3-) to form an H-bond with the

imidazole group of H104, which is a strong proton donor or acceptor at physiological pH (28). Thus, mutation of either C110 or H104 to alanine should suppress the inhibition because the non-polar side chain of alanine cannot serve as a strong proton donor or acceptor to form the H-bond (28). Accordingly, the mutants C14xA (“Cys-less”; H104 is the sole Zn++ ligand remaining) and C13xA-H104A (C110 only) exhibited responses to NO or H2O2 that were indistinguishable from background (k0=0.007- 0.013 s-1 and I%=27 %; Fig. 3C-D and 6). Similar results were obtained with C13xA-H104Q and C13xA-H104L (data not shown).

The H-bond hypothesis predicts that any modification of C110 that promotes the formation of an H-bond with the H104 should inhibit the channel profoundly. Thus, we carried out additional experiments with two internally-applied thiol-specific MTS reagents: MMTS, which adds a methyl group (-CH3) to the reacting cysteine but cannot serve as a strong proton donor or acceptor; and MTSEA, a strong proton donor, which adds the ammonium group (-NH3) to the reacting cysteine. As expected, MMTS inhibited the C13xA mutant less effectively than MTSEA

(MMTS: ko = 0.010 ± 0.001 s-1 and I% = 37 ± 8%; MTSEA: ko = 0.016 ± 0.006 s-1 and I% = 83 ± 3 %; Fig. 4E-F and 6); and furthermore, the H104A mutation prevented the inhibition by MTSEA, but had no effect on the response induced by MMTS (Fig. 4E-F and 6). H104L and H104Q produced similar results (data not shown). Thus, a putative H-bond between an highly oxidized form of C110 and the imidazole group of H104 may be responsible for the nitrosative regulation of the mutant Kv4 channels that may favor the interaction between those residues (Discussion).

Are C110 and H104 sufficiently close to permit the formation of the putative H-bond? To answer this question, we tested whether a disulfide bond between C110 and C104 (i.e., H104C) can straight-jacket the T1-T1 interface under mild oxidizing conditions and thereby inhibit the channel. Previously, we used this strategy to show that the T1-T1 interface is dynamic (9). Similarly, the current from the C13xA-H104C mutant (Fig. 5A) was inhibited upon exposing the cytoplasmic side of the patch to Cu-phenanthroline (Cu/P, a mild oxidizing agent) (Fig. 5C). Consistent with the formation of a disulfide bond between C110 and C104, this inhibition was not observed when C110 was mutated to alanine (Fig. 5B and C) and was reversed by the application of 20 mM DTT (Experimental Procedures; Fig. 5D). Moreover, the robust inhibition of the C13xA-H104C mutant by MTSET was significantly suppressed upon pre-oxidation of the cysteine pair with Cu/P. This result suggests that pre-forming a disulfide bond with C110 prevents the reaction with MTSET. Supporting this interpretation, the protective effect of a disulfide bond against MTSET was eliminated by the C110A mutation (Fig. 5E-F). The Zn++ site in the Kv4 T1-T1 interface is a redox switch Previous studies showed that NO can induce the formation of a disulfide bond when the thiolate groups of two cysteines are in close proximity (14,29). Once the first cysteine is nitrosylated, the sulfur atom becomes vulnerable to nucleophilic attack by a free thiol group from a nearby cysteine, causing release of the NO group and subsequent disulfide bond formation before the first cysteine becomes highly oxidized (29). In this

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

6

case, the disulfide bond can be reversed by a reducing reagent but the highly oxidized product cannot. As expected, application of 10 mM reduced GSH to the cytosolic side of a patch reversed the rapid NO-induced inhibition of the C11xA mutant quickly and completely. In contrast, the inhibition of the C13xA mutant which retained C110 and H104 at the T1 Zn++ site was irreversible (Fig. 7A). GSH per se had little or no effect on the C11xA current (Fig. 7D). Preferentially, disulfide bonding may occur between C132 and C110 upon nitrosation because the C12xA-b mutant (Table 1) was significantly inhibited by exposure to MAHMA-NONOate (Fig. 6) and this inhibition was fully reversed by GSH (data not shown). Using a mild oxidizing reagent in a previous study, we also inferred the preferential disulfide bonding between C110 and C132 (9). The presence of GSH-reversible NO modulation argues for a functional redox switch at the T1 Zn++ site of the Kv4 channel complex. Consistent with this hypothesis, a redox buffer consisting of 1 mM SNAP (a slow NO donor; Experimental Procedures) and 1 mM GSH inhibited the C11xA mutant but had little or no effect on the “Cys-less” C14xA mutant (Fig. 7B).

The thiol group of GSH can also bind to Zn++ with significant affinity (30). Thus, GSH may compromise the quaternary structure of the isolated Kv4 T1 tetramer, which depends on inter-subunit Zn++ binding (Fig. 1B) (3). As expected, GSH dissociated the purified Kv4.2-T1 tetramer into T1 monomers (Fig. 7C). Interestingly, however, a Zn++ atom remains bound to the T1 monomer (Fig. 7C legend). Conceivably, GSH competes off the weak Zn++-coordinating thiolate group of C110 and thus dissociates the tetramer but fails to release the tightly bound intra-subunit Zn++

coordinated by H104, C131 and C132. Supporting the presence of a weak interaction between C110 and Zn++ in the intact channel, the C11xA mutant induced robust voltage-dependent currents with similar kinetics under the following experimental conditions (consecutively, in the same patch): the cell-attached configuration (the intact oocyte cytoplasm may contain both Zn++ and GSH), inside-out configuration exposed to a Zn2+-free

intracellular solution, 10 µM ZnCl2 and 1 mM internal GSH (Fig. 7D). Thus, the weak inter-subunit Zn2+ bridge, unlike the disulfide or H-bond, would allow the conformational change in T1-T1 interface that is tightly coupled to voltage-dependent gating (9).

To confirm that the formation of a disulfide bond in the interfacial T1 Zn++ site is the mechanism that underlies the modulation of the Kv4.1 channel by NO, we exposed intact T293 cells to MAHMA-NONOate and subsequently examined the oligomeric state of the channel by SDS-PAGE electrophoresis and Western blot analysis (Fig. 8B). Without MAHMA-NONOate, most of the protein from WT, C12xA-b and C14xA (Fig.8A) appeared as a monomer even in the absence of DTT. In contrast, when the cells were treated with MAHMA-NONOate a significant amount of the protein from WT and C12xA-b migrated as high molecular weight oligomeric complexes; but the C14xA protein remained monomeric. Furthermore, treating the WT and C12xA-b proteins with DTT shifted them to their monomeric configurations (Fig. 8B). Even a small amount of high molecular weight Kv4.1 protein, which is seen in both the WT and C12xA-b lanes under basal conditions, was eliminated by DTT treatment. Thus, the Kv4.1 subunits with all cysteines in the T1 Zn++ site or two cysteines across the T1-T1 interface only (C110 and C132 in the Zn++ site of the C12xA-b mutant) undergo disulfide cross-linking, which is significantly enhanced upon nitrosation; and as expected, NO-induced cross-linking does not happen when there are no intracellular cysteines. These results show that disulfide cross-linking between C110 and C132 occurs in intact mammalian cells and can be regulated dynamically by NO. Indirectly, these results also suggest that Zn++ is not protecting C132 in the intact cell; otherwise, C132 would be involved in intra-subunit Zn++ coordination, and therefore, unavailable to react with C110. Zn++ protects the Kv4 channel complex against nitrosative or oxidative inhibition Although Zn++ binding is not required for channel gating (Fig. 7D) (5,6), intra-subunit Zn++-binding could protect against the formation of an inhibitory disulfide bond induced by nitrosation. Accordingly, saturating

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

7

concentrations of internal Zn++ suppressed the inhibition of the C11xA mutant (Fig. 9A) by NO (Fig. 9D) and H2O2 (Fig. 9E), and the inhibition of the C12xA-a mutant (Table 1; Fig. 9B; C131, C132 and H104 are available) by MTSET (Fig. 9F). Even when C110, C132 and H104 were left intact and C131 was mutated to Ala (C12xA-b; Fig. 9C), the inhibition by Cu/P was greatly reduced in the presence of internal Zn++ (Fig. 9G). These observations suggest that Zn++ binding to the Kv4 T1 domain (involving C131, C132 and H104) may protect the channel against nitrosative or oxidative inhibition by preventing the formation of the inhibitory interfacial disulfide bridge. Contrary to the other Zn++ site cysteines, C110 would be more susceptible to NO because it is weakly bound to Zn++. Supporting this hypothesis, the C11xA mutant was first slightly inhibited upon co-application of MAHMA-NONOate and ZnCl2 as seen before (background inhibition, Fig, 9D). During this period the unprotected C110 may become nitrosylated; therefore, subsequent washout of both reagents quickly inhibited the mutant channel because the removal of Zn++ allows the close interaction between nitrosylated C110 and the free thiol group of C132 to form the disulfide bond (Supplemental data, Fig. S2).

DISCUSSION

The intracellular T1 domain of Kv

channels is a multitasking region that is responsible for subfamilily specific assembly, docking of auxiliary subunits and gating (1,2,7,9,10). The role of the T1 domain in gating is puzzling because the membrane spanning regions that constitute the Kv channel core control voltage-dependent gating and K+-selective permeation. Nevertheless, the T1-T1 intersubunit interface appears to undergo a conformational change coupled to voltage-dependent gating (9). In intact Kv4 channels, restriction of this conformational change by modification of specific T1 Zn++ site cysteines hinders activation (5,9). The T1-T1 conformational change may correspond to the cooperative introduction of a permissive closed state that precedes the opening of the pore. Thus, the T1-T1 interface would be a site

where intracellular signaling molecules can modulate Kv4 channel gating effectively. The presence of the reactive cysteines in the T1 Zn++ site of Kv4 channels makes this special location an attractive target for redox modulation. The results showed that the Kv4.1 channel is quickly and profoundly inhibited by intracellular NO in a Zn++–dependent manner. This inhibition may involve two distinct and mutually exclusive interactions: 1) a disulfide bridge across the T1-T1 interface (C110-C132) (Fig. 10A), which may be physiologically significant because a reducing agent (GSH) readily reverses the inhibition by NO; and 2) a putative H-bond between two interfacial Zn++ site determinants, H104 and C110 (Fig, 10B). Given that the interfacial T1 Zn++ site is highly conserved, it may act as a Zn++–dependent redox switch in neuronal and cardiac tissues where Kv4 channels are key regulators of membrane excitability. Molecular interactions responsible for the redox modulation at the T1 Zn++site of a Kv4 channel In the Kv4.1 T1-T1 interface, C131, C132 and H104 form a metal coordination complex with structural Zn++ in a reduced intracellular environment (Fig. 10A). Patch excision may induce the release of Zn++ and C110 may be oxidized by H2O2, MTSEA or nitrosylated by NO. The modified or highly oxidized C110 would then form an H-bond with the imidazole group of H104 when C131 and C132 are not available (Figs. 4C, D, F and 10B). Because the inhibition by NO and H2O2 is not reversed by GSH, we propose that the thiol group of C110 is highly oxidized from (Cys-S-NO) or (Cys-S-OH) to (Cys-S-O2H) or (Cys-S-O3H) (Fig. 10B) (29,31,32). However, when C132 and C131 are available, the nitrosylated C110 may prefer to form a disulfide bridge with the thiol group of C132 (Fig. 10A). This event would, however, depend on the occupancy of the Zn++ site, as discussed below. GSH did not reverse the inhibition by high concentrations of H2O2 even when the Zn2+ site cysteines are available (Supplemental data, Fig. S1D). We propose that a highly oxidized C110 forms a thiolsulfinate with the thiol group of C132 (Fig. 10A) (33).

Supporting the proposed mechanisms (Fig. 10), structural studies have demonstrated

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

8

that either hydrogen-bonding or disulfide bridging at the active sites of enzymes are important interactions associated with modulation by oxidation. The former involves a single cysteine and a nearby non-cysteine residue, but the latter is formed between two adjacent cysteines. In any case, the reaction of NO and H2O2 with reactive thiols results in a local structural rearrangement that promotes enzyme activation or inactivation. For example, the Oδ3 atom of the oxidized C25 forms H-bonds with His159-Nδ1 in an inactive form of papain (34). In the flavoprotein NADH peroxidase with its native C42-sulfenic acid redox center, C42 Oδ

is H-bonded to H10 Nε2 (35,36). H-bonding interactions involving oxidized cysteines are also found in human glutathione reductase (hGR) (37), protein tyrosine phosphatase 1B (PTP1B) (38) and the photosensitive nitrile hydratase (Nhase) (39). In addition to the H-bonding interactions, disulfide bridging is also found in thiol-based regulatory switches with (RsrA, PerR, and Hsp33) or without (OxyR, OhrR, and CrtJ) metal involvement (12,33,40).

Although the proposed H-bond appears to be sufficient to strait-jacket the T1-T1 interface and thus to inhibit channel activity, it may only be formed in the C13xA mutant. More likely, the native Kv4 channel complex employs thiol-disulfide exchange reactions at the Zn++ site to regulate the channel’s activity in response to reactive nitrosative or oxidative species under physiological conditions. However, nitrosative or oxidative stress could also produce irreversible thiolsulfinate or thiolsulfonate at the Zn++ site under pathological conditions (33). The role of Zn++ binding in the interfacial T1 Zn++ site of Kv4 channels Unlike the disulfide bridge or the H-bond, the intersubunit Zn++ bridge involving C110 may be weak and thus fails to inhibit channel activity by straight-jacketing the dynamic T1-T1 interface. Previously, this possibility was considered likely because at least one cysteine at the Kv4-T1 Zn2+ site is free to react with MTSET in a Zn++-independent manner (5). The new results reported here strongly suggest that C110 is the free reactive cysteine. First, GSH appears to

disrupt the interfacial Zn++ bridge without affecting Zn++ content and the Kv4.1 current (Fig. 7C–D). Second, the presence of Zn++ protects C131 and C132 from MTSET modification when H104 is available (Fig. 9F). Third, the presence of Zn++ appears to prevent the inhibitory interfacial disulfide bonds or thiolsulfinate induced by NO, Cu/P or H2O2 (Fig. 9D, E and F). Fourth, the presence of Zn2+ cannot prevent oxidation of C110 by NO (Supplemental data, Fig. S2). Finally, the Kv4.1 current is independent of Zn++, GSH (Fig. 7D), Zn++-binding cysteines or TPEN (5) but tightly coupled to the conformational change in the T1-T1 interface (9). Therefore, C110 is free or coordinates Zn++ weakly to keep the T1-T1 interface functionally active and dynamic. A similar case was also found in latent human fibroblast collagenase whose activation depends on the dissociation of Cys73 from a zinc-binding site (41). In contrast to the weak interfacial binding between C110 and Zn++, the coordination of Zn++ by C131, C132 and H104 is stronger; and therefore, elevated intracellular Zn++ may prevent the nitrosative regulation (see below). Zn++ protection against the effects of oxidation was also found in transcripton factor proteins (33) and dimethylargininase-1, which is inhibited by Cys-S-nitrosylation (42). Physiopathological implications Thiol-based redox cellular regulation in response to nitrosative, oxidative or disulfide signaling has received significant attention (11,12,29,33,40,43). In particular, several studies have suggested that the physiopathology of degenerative brain disorders is linked to intracellular Zn++

homeostasis and oxidative/nitrosative stress (13,17,44-46). Therefore, the modulation of the neuronal Kv4 channel complex by nitrosative/oxidative stress is especially relevant. This channel complex underlies the somatodendritic A-type K+ current (ISA) (47-49), which regulates different aspects of neuronal excitability such as frequency of repetitive firing, dampening of back-propagating action potentials, and compartmentalization of dendritic action potentials. Therefore, ISA ultimately impacts somatodendritic signal integration (47,50).

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

9

Under basal physiological conditions, Zn++ buffering systems keep the concentration of free Zn++ in the subnanomolar range (11,51); and probably, there is an equilibrium between the Zn++–bound and Zn++–free species of the Kv4 channel. Thus, the Zn++ site may undergo nitrosative redox modulation, which would induce a disulfide bond across the T1-T1 interface and thereby inhibit Kv4 channel gating. As a result, distal dendrites would become hyperexcitable. This modulation may depend on the concentrations of intracellular Zn++. For instance, elevated intracellular Zn++ resulting from intracellular Zn++ overload or mobilization induced by enhanced synaptic activity or NO, respectively, may target the Kv4 Zn++ site (11). Zn++ binding may thus protect against nitrosation/oxidation by preventing disulfide bond formation across the T1-T1 interface or excessive cysteine oxidation. A local and rapid Zn++ overload in dendritic spines may occur during excitotoxic

activity and NO signaling may mobilize Zn++ by cysteine nitrosylation of critical Zn++ buffering proteins (metallothioneins) (11,46). Conceivably, protecting the activity of Kv4 channels by Zn++ is an acute compensatory mechanism that helps to regulate electrical excitability under physiological and pathological conditions. This mechanism may account for nitrosative/oxidative modulation of the A-type K+ currents in neurons, heart and smooth muscle (15,18,19,21,22). Furthermore, it may also confer Zn++–dependent redox modulation to Kv2 and Kv3 channels, which share the T1-T1 Zn++ site. Zn++ protection and strong reducing conditions in normal cells may explain the resistance of the Kv4 channel to externally applied reactive oxygen species (26,27,52). This Zn++–dependent redox regulation is distinct from that found in Kv1 channel complexes, where the Kv1-β subunits act as redox enzymes (53).

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

10

REFERENCES

1. Bixby, K. A., Nanao, M. H., Shen, N. V., Kreusch, A., Bellamy, H., Pfaffinger, P. J., and Choe, S. (1999) Nat. Struct. Biol. 6, 38-43

2. Nanao, M. H., Zhou, W., Pfaffinger, P. J., and Choe, S. (2003) Proc. Natl. Acad. Sci. U. S. A 100, 8670-8675

3. Strang, C., Kunjilwar, K., DeRubeis, D., Peterson, D., and Pfaffinger, P. J. (2003) J Biol. Chem. 278, 31361-31371

4. Jahng, A. W., Strang, C., Kaiser, D., Pollard, T., Pfaffinger, P., and Choe, S. (2002) J Biol. Chem. 277, 47885-47890

5. Wang, G., Shahidullah, M., Rocha, C. A., Strang, C., Pfaffinger, P. J., and Covarrubias, M. (2005) J. Gen. Physiol 126, 55-69

6. Kunjilwar, K., Strang, C., DeRubeis, D., and Pfaffinger, P. J. (2004) J. Biol. Chem. 279, 54542-54551

7. Pioletti, M., Findeisen, F., Hura, G. L., and Minor, D. L., Jr. (2006) Nat. Struct. Mol. Biol. 13, 987-995

8. Wang, H., Yan, Y., Liu, Q., Huang, Y., Shen, Y., Chen, L., Chen, Y., Yang, Q., Hao, Q., Wang, K., and Chai, J. (2007) Nat. Neurosci. 10, 32-39

9. Wang, G. and Covarrubias, M. (2006) J. Gen. Physiol 127, 391-400 10. Long, S. B., Campbell, E. B., and MacKinnon, R. (2005) Science 309, 897-902 11. Maret, W. (2006) Antioxid. Redox. Signal. 8, 1419-1441 12. Paget, M. S. and Buttner, M. J. (2003) Annu. Rev. Genet. 37, 91-121 13. Hao, Q. and Maret, W. (2005) J. Alzheimers. Dis. 8, 161-170 14. Lipton, S. A., Choi, Y. B., Takahashi, H., Zhang, D., Li, W., Godzik, A., and Bankston,

L. A. (2002) Trends Neurosci. 25, 474-480 15. Muller, W. and Bittner, K. (2002) J. Neurophysiol. 87, 2990-2995 16. Gow, A. and Ischiropoulos, H. (2002) Am. J. Physiol Lung Cell Mol. Physiol 282, L183-

L184 17. Bossy-Wetzel, E., Talantova, M. V., Lee, W. D., Scholzke, M. N., Harrop, A., Mathews,

E., Gotz, T., Han, J., Ellisman, M. H., Perkins, G. A., and Lipton, S. A. (2004) Neuron 41, 351-365

18. Han, N. L., Ye, J. S., Yu, A. C., and Sheu, F. S. (2006) J. Neurophysiol. 95, 2167-2178 19. Prasad, M. and Goyal, R. K. (2004) Am. J. Physiol Cell Physiol 286, C671-C682 20. Ruschenschmidt, C., Chen, J., Becker, A., Riazanski, V., and Beck, H. (2006) Eur. J.

Neurosci. 23, 675-685 21. Rozanski, G. J. and Xu, Z. (2002) J. Mol. Cell Cardiol. 34, 1623-1632 22. Rozanski, G. J. and Xu, Z. (2002) Am. J. Physiol Heart Circ. Physiol 282, H2346-H2355 23. Fitzhugh, A. L. and Keefer, L. K. (2000) Free Radic. Biol. Med. 28, 1463-1469 24. Liu, Y., Jurman, M. E., and Yellen, G. (1996) Neuron 16, 859-867 25. Hoshi, T. and Heinemann, S. (2001) J. Physiol 531, 1-11 26. Serodio, P., Kentros, C., and Rudy, B. (1994) J. Neurophysiol. 72, 1516-1529 27. Duprat, F., Guillemare, E., Romey, G., Fink, M., Lesage, F., Lazdunski, M., and Honore,

E. (1995) Proc. Natl. Acad. Sci. U. S. A 92, 11796-11800 28. Ippolito, J. A., Alexander, R. S., and Christianson, D. W. (1990) J. Mol. Biol. 215, 457-

471 29. Stamler, J. S. and Hausladen, A. (1998) Nat. Struct. Biol. 5, 247-249

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

11

30. Neculai, A. M., Neculai, D., Griesinger, C., Vorholt, J. A., and Becker, S. (2005) J. Biol. Chem. 280, 2826-2830

31. Poole, L. B., Karplus, P. A., and Claiborne, A. (2004) Annu. Rev. Pharmacol. Toxicol. 44, 325-347

32. Claiborne, A., Yeh, J. I., Mallett, T. C., Luba, J., Crane, E. J., III, Charrier, V., and Parsonage, D. (1999) Biochemistry 38, 15407-15416

33. Wilcox, D. E., Schenk, A. D., Feldman, B. M., and Xu, Y. (2001) Antioxid. Redox. Signal. 3, 549-564

34. Kamphuis, I. G., Kalk, K. H., Swarte, M. B., and Drenth, J. (1984) J. Mol. Biol. 179, 233-256

35. Crane, E. J., III, Vervoort, J., and Claiborne, A. (1997) Biochemistry 36, 8611-8618 36. Yeh, J. I., Claiborne, A., and Hol, W. G. (1996) Biochemistry 35, 9951-9957 37. Becker, K., Savvides, S. N., Keese, M., Schirmer, R. H., and Karplus, P. A. (1998) Nat.

Struct. Biol. 5, 267-271 38. van Montfort, R. L., Congreve, M., Tisi, D., Carr, R., and Jhoti, H. (2003) Nature 423,

773-777 39. Noguchi, T., Nojiri, M., Takei, K., Odaka, M., and Kamiya, N. (2003) Biochemistry 42,

11642-11650 40. Ilbert, M., Graf, P. C., and Jakob, U. (2006) Antioxid. Redox. Signal. 8, 835-846 41. Springman, E. B., Angleton, E. L., Birkedal-Hansen, H., and Van Wart, H. E. (1990)

Proc. Natl. Acad. Sci. U. S. A 87, 364-368 42. Knipp, M., Braun, O., Gehrig, P. M., Sack, R., and Vasak, M. (2003) J. Biol. Chem. 278,

3410-3416 43. Yoshida, T., Inoue, R., Morii, T., Takahashi, N., Yamamoto, S., Hara, Y., Tominaga, M.,

Shimizu, S., Sato, Y., and Mori, Y. (2006) Nat. Chem. Biol. 2, 596-607 44. Frederickson, C. J., Koh, J. Y., and Bush, A. I. (2005) Nat. Rev. Neurosci. 6, 449-462 45. Capasso, M., Jeng, J. M., Malavolta, M., Mocchegiani, E., and Sensi, S. L. (2005) J.

Alzheimers. Dis. 8, 93-108 46. Takeda, A. (2001) Biometals 14, 343-351 47. Jerng, H. H., Pfaffinger, P. J., and Covarrubias, M. (2004) Mol. Cell Neurosci. 27, 343-

369 48. Jerng, H. H., Kunjilwar, K., and Pfaffinger, P. J. (2005) J. Physiol 568, 767-788 49. Nadal, M. S., Ozaita, A., Amarillo, Y., de Miera, E. V., Ma, Y., Mo, W., Goldberg, E.

M., Misumi, Y., Ikehara, Y., Neubert, T. A., and Rudy, B. (2003) Neuron 37, 449-461 50. Birnbaum, S. G., Varga, A. W., Yuan, L. L., Anderson, A. E., Sweatt, J. D., and

Schrader, L. A. (2004) Physiol Rev. 84, 803-833 51. Colvin, R. A., Fontaine, C. P., Laskowski, M., and Thomas, D. (2003) Eur. J. Pharmacol.

479, 171-185 52. Akar, F. G., Wu, R. C., Deschenes, I., Armoundas, A. A., Piacentino, V., III, Houser, S.

R., and Tomaselli, G. F. (2004) Am. J. Physiol Heart Circ. Physiol 286, H602-H609 53. Heinemann, S. H. and Hoshi, T. (2006) Sci. STKE. 2006, e33 FOOTNOTES

We thank Dr. Mark Bowlby (Wyeth Research, Princeton, NJ) for providing KChIP1 cDNA and Dr. Bernardo Rudy (New York University, New York, NY) for providing DPPX-S cDNA. Also, we thank members of the Covarrubias lab (Mr. Aditya Bhattacharji, Mr. Kevin Dougherty and Mr.

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

12

Thanawath Harris) for constructing Kv4.1 mutants for expression in mammalian cells, and verification of functional expression; and Dr. Toshinori Hoshi for suggesting SNAP as an alternative NO donor. This work was supported by NIH research grants R01 NS032337 (MC), P01 NS037444 (PJP) and NIH training grant T32 AA07463 (GW).

#The abbreviations used are: DTT, DL-dithiotreitol; SNAP, S-nitroso-N-acetylpenicillamine; GSH, glutathione; MAHMA-NONOate, (NOC-9, MAHMA/NO, (Z)-1 {N-Methyl-N-[6-(N-methylammoniohexyl) amino]} diazen-1-ium-1, 2-diolate]; DEA-NONOate, sodium 2-(N,N-dimethylamino)-diazenolate-2-oxide; NO, nitric oxide; H2O2, hydrogen peroxide; MTS, methanethiosulfonate; MTSET, 2-trimethylammonium-ethyl-methanethiosulfonate bromide; MMTS, methyl methanethiolsulfonate; MTSEA, 2-aminoethyl methanethiosulfonate hydrobromid; TPEN, tetrakis-(2-pyridylmethyl) ethylendiamide); WT, wild-type; Cu/P, Cu-phenanthroline.

FIGURE LEGENDS

Fig. 1 A structural model of the Kv4 channel. A, left, side view of the Kv4 tetramer consisting transmembrane voltage-sensor domain (VSD) and pore domain (PD) and the intracellular T1 domain. For clarity, the auxiliary subunits KChIP1 and DPPX-S are not shown. Right, top view of the Kv4.2-T1 domain. Four blue spheres represent the location of Zn++ atoms in the T1-T1 inter-subunit interfaces as found in the crystal structure of the isolated Kv4.2-T1 domain (2). B, Stereo close-up of the Zn++ binding site in the T1 domain. H104, C131 and C132 are from the same subunit and C110 is from the neighboring subunit. A standard color scheme is used to represent the relevant atoms (sulfur atoms in yellow). The atomic distances between Zn++ and H104, C110, C131 and C132 are 2.1, 2.4, 2.2 and 2.2 Ǻ, respectively. Fig. 2 Inhibition of the Kv4.1 channel by NO. Left, Simplified views of the T1 Zn++ site residues in WT (A) and C11xA (D) and C3xA (G). The “+” represents the location of the Zn++ atom as found in the crystal structure of the isolated Kv4.2-T1 domain (2). Middle, WT (B), C11xA (E) and C3xA (H) currents from inside-out patches before and after the internal application of 100 µM MAHMA-NONOate. All channels were co-expressed with DPPX-S and KChIP-1 (Experimental Procedures). Currents were evoked by a step depolarization from a holding potential of -100 mV to +80 mV (start-to-start interval was 3-s). Right, the normalized peak currents plotted against time for WT (C), C11xA (F) and C3xA (I). Solid lines are best-fit single exponential decays with the following time constants and fractional current levels at steady-state (in parenthesis): 28 s (0.36), 42 s (0.12), 50 s (0.75) for WT, C11xA and C3xA, respectively. Fig. 3 Inhibition of Kv4.1 Zn++ site mutants by NO and H2O2. Left, Simplified views of the T1 Zn++ site residues in C14xA (A), C12xA-a (D) and C13xA (G). Middle, time-dependent inhibition of C14xA (B), C12xA-a (E) and C13xA (H) by internally-applied MAHMA-NONOate (100 µM). Solid lines are best-fit single exponential decays with the following time constants and fractional current levels at steady-state (in parenthesis): 145 s (0.69), 146 s (0.71), 39 s (0.35), for C14xA, C12xA-a, and C13xA, respectively. When pretreated with MMTS (hollow symbols; H), the inhibition of the C13xA mutant channel was suppressed (best-fit time constant and fractional current were 240 s and 0.74, respectively). Right, time-dependent inhibition of C14xA (C), C12xA-a (F) and C13xA (I) by internally-applied H2O2 (58 mM). Solid lines are best-fit single exponential decays with the following time constants and fractional current levels at steady-state (in parenthesis): 238 s (0.70), 152 s (0.72), 102 s (0.34) for C14xA, C12xA-a, and C13xA, respectively. External application (hollow symbols) had not effect (C). The presence of 20 µM TPEN (hollow symbols) did not affect the inhibition of the C12xA-a mutant channel (F). When pretreated with 1 mM MMTS (hollow symbols; I), the inhibition

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

13

of the C13xA mutant channel was suppressed (best-fit time constant and fractional current are 217 s and 0.68, respectively. All plots were generated as explained in Fig. 2 legend. Fig. 4 The presence of C110 and H104 is sufficient to observe the inhibition of Kv4.1 channels by NO, H2O2 and MTSEA. Upper, Simplified views of the T1 Zn++ site residues in C13xA (A) and C13xA-H104A (B). Below, time-dependent inhibition of C13xA (solid symbols) and C13xA-H104A (hollow symbols) mutants by internally-applied 100 µM MAHMA-NONOate (C), 58 mM H2O2 (D), 1 mM MMTS (E) and 200 µM MTSEA (F). For C13xA, solid lines are best-fit single exponential decays with the following time constants and fractional current levels at steady-state (in parenthesis): 39 s (0.35) (C), 102 s (0.34) (D), 166 s (0.69) (E) and 63 s (0.20) (F). For C13xA-H104A mutant, solid lines are best-fit exponential decays with the following time constants and fractional current levels at steady-state (in parenthesis): 147 s (0.66) (C), 126 s (0.71) (D), 166 s (0.69) (E) and 110 s (0.62) (F). All plots were generated as explained in Fig. 2 legend. Fig. 5 Disulfide cross-linking across the T1-T1 inter-subunit interface inhibits the Kv4.1 channel. Upper, Simplified views of the T1 Zn++ site residues in C13xA-H104C (A) and C14xA-H104C (B). Below, C, Inhibition of a Kv4.1 mutant (C13xA-H104C) by internally applied Cu/P. Solid line is the best-fit single exponential decay with the following time constant and fractional current level at steady-state (in parenthesis): 115 s (0.50). The mutant C14xA-H104C exhibited no response. D, Reversibility of the Cu/P-induced inhibition of the C13xA-H104C mutant by DTT (20 mM). E, Pretreatment of C13xA-H104C with Cu/P reduced the fractional current level upon inhibition by MTSET from 0.27 to 0.77 without changing the time constant (∼91 s). F, For the mutant C14xA-H104C channel with C104 only, pretreatment of Cu/P did not affect inhibition of the channel by MTSET. The best-fit single exponential decay gave the following time constant and fractional current level at steady-state (in parenthesis): 56 s (0.26). All plots were generated as explained in Fig. 2 legend, except the plot in panel B, which is not normalized.

Fig. 6 Summary of the modulation of Kv4.1 wild-type and mutant channels by reagents that target the T1 Zn++ site. A, Percent inhibition; B, best-fit inhibition rate constant. For H2O2 and NO, the differences against C14xA are statistically significant at p<0.05 (∗), <0.01(∗∗) and <0.001 (∗∗∗) (one-way ANOVA test, n=4-8). For MMTS and MTSEA, the control is C13xA-H104A. Fig. 7 Evidence of a redox switch in the T1 Zn++ site of Kv4.1 channels. A, Time course of the normalized peak current from C11xA (solid symbols) and C13xA (hollow symbols) mutants. GSH (10 mM) reversed the inhibition of the C11xA mutant by NO. In contrast, the inhibition of the C13xA mutant was not reversed. B, Time course of the normalized peak current from C11xA (solid symbols) and C14xA (hollow symbols) mutants. The combined presence of 1 mM SNAP and 1 mM GSH served as a redox buffer and thus reduced the fast inhibition of the C11xA channel by NO (Fig. 2). The best-fit single exponential decay gave the following time constant and fractional current level at steady-state (in parenthesis): 119 s (0.66). The C14xA channel was not responsive. C, FPLC profile of the Kv4.2-T1 protein before (upper) and after (below) treatment with reduced GSH (2 mM) for 20 min. AU280 = normalized absorbance units at 280 nm. The expected elution of the T1 tetramer and the T1 monomer are marked as “T” and “M” below the peak, respectively. “V-V” represents the void volume. The T1 domain Zn++ content was not changed upon GSH treatment, as determined by atomic absorbance spectroscopy (Experimental Procedures). D, Outward macropatch currents induced by the Kv4.1-C11xA ternary complex (Experimental Procedures). From the same patch under the conditions indicated in the graph, these currents were evoked by a depolarizing pulse from -100 to +80 mV. The inside-out patch was first exposed to a saturating concentration (10 µM) of internally-applied ZnCl2, and then to 1 mM GSH 5 min later.

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

14

Fig. 8 Western blot analysis of Kv4.1 sensitivities to disulfide cross-linking upon exposure to NO. A, Simplified views of the T1 Zn++ site residues in WT, C12xA-b and C14xA. B, Kv4.1 cDNAs (WT, C12xA-b and C14xA) were co-transfected into T293 cells with KChIP3 and EGFP plasmids. Transfected cells were incubated with or without MAHMA-NONOate for 20 min immediately before solubilization in SDS sample buffer, SDS-PAGE gel electrophoresis and Western blotting for Kv4.1 protein (anit-Kv4.1 antibody; Experimental Procedures). The solubilized sample was divided in two equal portions and half of each sample was reduced with DTT prior to loading the gel. The Kv4.1 construct used and treatment received for each sample are indicated by (+) signs above the respective lanes. Under reduced film exposure conditions a reduction in the monomer band intensity is clearly evident following NO induced cross-linking (data not shown). Fig. 9 Zn++-dependent modulation of the Kv4.1 channel inhibition by NO and H2O2. Upper, Simplified views of the T1 Zn++ site residues in WT/C11xA (A), C12xA-a (B) and C12xA-b (C). The blue spheres represent the Zn++ atoms as found in the crystal structure of the isolated Kv4.2-T1 domain (2). The thick and thin dash lines represent strong and weak bond, respectively. Below, time-dependent inhibition of C11xA (solid) and C14xA (hollow symbols) by internally-applied MAHMA-NONOate (100 µM) (D) or H2O2 (5.8 mM) (E) in the presence (grey) or absence (black) of a saturating concentration ZnCl2 (10 µM). C14xA is a control mutant without any intracellular cysteines. D, Zn++ dependence of the inhibition of Kv4.1 Zn++ site mutants by MAHMA-NONOate. The solid lines are the best-fit single exponential decays with the following time constants and fractional current levels at steady-state (in parenthesis): 42 s (0.12), 117 s (0.59) and 146 s (0.69) for C11xA, C11xA+Zn++ and C14xA, respectively. E, Zn++ dependence of the inhibition of Kv4.1 Zn++ site mutants by H2O2. The solid lines are the best-fit single exponential decays with the following time constants and fractional current levels at steady-state (in parenthesis): 217 s (0.19), 308 s (0.36) and 142 s (0.58) for C11xA, C11xA+Zn++ and C14xA, respectively. F, Inhibition of the C12xA-a channel by MTSET (200 µM) in the absence (black) and presence (grey) of 10 µM ZnCl2. The solid lines are the best-fit single exponential decays with the following time constants and fractional current levels at steady-state (in parenthesis): 39 s (0.21) and 108 s (0.67) for C11xA and C11xA+ Zn++, respectively. G, Zn++ dependence of the Cu/P-induced inhibition of the Kv4.1 mutant C12xA-b with C110, C132 and H104 in the Zn2+ site only. The presence of Zn++ increases the fractional current level from 0.52 to 0.83 and the time constant from 65 s to 202 s. All plots were generated as explained in Fig. 2 legend. Fig. 10 Working hypothesis of the Kv4 channel inhibition by three oxidants. For WT and C11xA (A), Zn++ is bound to C131, C132 and H104 in the T1 domain of the intact Kv4.1 channel. C110 interacts with Zn++ weakly and thus the T1-T1 interface is dynamic. Dashed lines represent the boundaries of the T1-T1 interface. Even if C110 is oxidized by H2O2 or NO, failure to form an H-bond or a disulfide bond across the T1-T1 interface prevents the modulation of the channel. However, once Zn++ is released, nitrosylation of C110 induces the formation of a disulfide bridge with C132. High concentration of H2O2 may highly oxidize C110 and thus form a thiolsulfinate with C132, which cannot be reversed by GSH (33). In any case, straight-jacketing the interface causes channel inhibition, as reported previously (9). For C13xA (B), oxidation of C110 by H2O2 or NO or MTSEA induces an inter-subunit H-bond with the imidazole group of H104 and thus inhibits the channel activity. The nitrosylated C110 may be oxidized further by NO to sulfinic acid. The presence of GSH can only reverse the disulfide bond and the nitrosylation of C110.

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

WSPC2JBC

15

Table I. Nomenclature and cysteine content of Kv4.1 constructs.

Kv4.1 Construct Number of Intracellular

Cys∗

Cys at the Zn++ site¶

Wild-type 14§ C110, C131, C132

C3xA 11‡ None

C11xA 3 C110, C131, C132

C12xA-a 2 C131, C132

C12xA-b 2 C110, C132

C12xA-c 2 C110, C131

C13xA 1 C110

C13xA-H104A 1 C110

C13xA-H104C 2 C110, C104

C14xA 0 None

C14xA-H104C 1 C104

* All constructs include four extracellular cysteines: C209, C223 and C233, C338. ¶ Except for three instances, H104 is left intact in most constructs. § Fourteen intracellular cysteines available: C105, C110, C131, C132, C257, C322, C392, C467, C484, C490, C532, C533,C589 and C642. ‡ Eleven intracellular cysteines available: C105, C257, C322, C392, C467, C484, C490, C532, C533, C589 and C642.

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from

AFig. 1 Wang et al.

B

90o

ext

int

VSD + PD

T1

Kv4-T1

C132

C131C110

H104

C132

C131C110

H104

by guest on August 11, 2019 http://www.jbc.org/ Downloaded from

0 200 4000.0

0.4

0.8

1.2

0 200 4000.0

0.4

0.8

1.2

0 200 4000.0

0.4

0.8

1.2

NO-50 ms200 pA

C -

C -

NO-

C -NO-

CB

E

H

F

50 ms

50 pA

I p/I p

-max

NO

I p/I p

-max

Exposure Time (s)

Exposure Time (s)

I p/I p

-max

Exposure Time (s)

I

50 ms

50 pA

Fig. 2 Wang et al.

C132

C131

C110

H104

C132

C131

C110

H104

A132

A131

A110

H104

A

D

G

WT

C11xA

C3xA

by guest on August 11, 2019 http://www.jbc.org/ Downloaded from

Fig. 3 Wang et al.

A132

A131

A110

H1040 200 400

0.0

0.4

0.8

1.2

0 200 4000.0

0.4

0.8

1.2

0 200 4000.0

0.4

0.8

1.2

0 400 8000.0

0.4

0.8

1.2

0 200 4000.0

0.4

0.8

1.2

0 150 300 4500.0

0.4

0.8

1.2

C14xA

I p/I p

-max

C14xANO

C13xAC13xA

C12xA-aC12xA-a

I p/I p

-max

IH

E F

C

I p/I p

-max

Exposure Time (s)

Control Pre MMTS

B

I p/I p

-max

I p/I p

-max

I p/I p

-max

I p/I p

-max

I p/I p

-max

I p/I p

-max

Exposure Time (s)

Control Pre MMTS

Exposure Time (s) Exposure Time (s)

Exposure Time (s) Exposure Time (s)

Internal External

H2O2

Control TPEN

A

D

G

C14xA

C12xA-a

C13xA

C132

C131

A110

H104

A132

A131

C110

H104

by guest on August 11, 2019 http://www.jbc.org/ Downloaded from

Fig. 4 Wang et al.C13xA C13xA-H104A

0 200 4000.0

0.4

0.8

1.2

0 300 6000.0

0.4

0.8

1.2

0 400 8000.0

0.4

0.8

1.2

0 150 300 4500.0

0.4

0.8

1.2

Exposure Time (s)

I p/I p

-max

C13xAC13xA-H104A

MMTSI p

/I p-m

axI p

/I p-m

ax

C13xAC13xA-H104A

MTSEA

Exposure Time (s)

I p/I p

-max

Exposure Time (s)

F

D

E

C13xAC13xA-H104A

H2O2 C

Exposure Time (s)

C13xAC13xA-H104A

NO

A B

A132

A131

C110

H104

A132

A131

C110

A104

by guest on August 11, 2019 http://www.jbc.org/ Downloaded from

Fig. 5 Wang et al. C14xA-H104C

A132

A131

A110

C104

C13xA-H104C

A132

A131

C110

C104

A B

0 200 4000.0

0.4

0.8

1.2

0 200 4000.0

0.4

0.8

1.2

0 400 8000

100

200

0 150 300 4500.0

0.4

0.8

1.2

I p/I p

-max

Exposure Time (s)

C13xA-H104C C13xA-H104C, Pre Cu/P

MTSET

Exposure Time (s)

FE

D

I p/I p

-max

C13xA-H104C C14xA-H104C

Cu/PC

Exposure Time (s)

Peak

cur

rent

(pA

)

C13xA-H104C

Cu/P DTT

I p/I p

-max

Exposure Time (s)

C14xA-H104C Pre Cu/PMTSET

by guest on August 11, 2019 http://www.jbc.org/ Downloaded from

C13xA-H104A

C3xA

C14xA

C13xA

C12xA-c

C12xA-b

C12xA-a

C11xA

WT

0 40 80

0 40 80

% Inhibition0 1 2 3 4 5

0 1 2 3 4 5

Rate Constant (10-2 s-1)

NO

H2O2

MTSEA

MMTS

Fig. 6 Wang et al.

*** ******

******

******** ***

*** **

*

***

***

**

A B

by guest on August 11, 2019 http://www.jbc.org/ Downloaded from

-150 0 150 300 450 6000.0

0.4

0.8

1.2

-100 0 100 200 300 4000.0

0.4

0.8

1.2

Exposure Time (s)

I p/I p

-max

C11xA C13xA

NO GSH

Cell-attached Inside-out 10 µM ZnCl2 1 mM GSH

50 ms

50 p

A

C11xA

I p/I p

-max

Exposure Time (s)

C14xAC11xA

SNAP+GSH

Fig. 7 Wang et al.

A

B

C

D

V-V T

V-V M

Kv4.2-T1

Kv4.2-T1 + GSH

by guest on August 11, 2019 http://www.jbc.org/ Downloaded from

WT C12xA-b

A

B

C14xAFig. 8 Wang et al.

}

}

X-linked

Monomer

200 -120 -100 -

53 -

37 -

(Kd)

}

}

X-linked

Monomer

200 -120 -100 -

53 -

37 -

(Kd)

X-linked

Monomer

200 -120 -100 -

53 -

37 -

(Kd)

200 -120 -100 -

53 -

37 -

(Kd)

++++++------DTT+++---+++---NONOate+--+--+--+--C12xA-b-+--+--+--+-C14xA--+--+--+--+WT

++++++------DTT+++---+++---NONOate+--+--+--+--C12xA-b-+--+--+--+-C14xA--+--+--+--+WT

MW(Kd)

C132

C131

C110

H104

C132

A131

C110

H104

A132A131

A110

H104

by guest on August 11, 2019 http://www.jbc.org/ Downloaded from

0 100 200 3000.0

0.4

0.8

1.2

0 250 500 7500.0

0.4

0.8

1.2

0 250 500 750 10000.0

0.4

0.8

1.2

0 200 4000.0

0.4

0.8

1.2

Exposure Time (s)

C12xA-b + Zn++

C12xA-bCu/P

I p/I p

-max

Exposure Time (s)

C12xA-a + Zn++

C12xA-aMTSET

I p/I p

-max

I p/I p

-max

G

ED

Exposure Time (s)

C14xA C11xA C11xA + Zn++

H2O2

F Exposure Time (s)

I p/I p

-max

C14xA C11xA C11xA + Zn++

NO

Fig. 9 Wang et al.WT / C11xA C12xA-a C12xA-bA B C

C132

C131

C110H104

C132

C131

A110H104

C132

A131

C110

H104

by guest on August 11, 2019 http://www.jbc.org/ Downloaded from

Fig. 10 Wang et al.

NN

His104

SHCys110

Zn2+

S

Cys132

Zn2+

SO

SCys132

Cys110

SS

Cys110

Cys132

H2O2

NO

GSH

S

Cys131

NHN

His104

SHCys110

SH

Cys132

SH

Cys131

SO

O-

Cys110

SNOCys110

NNH

His104

H2O2

NO

MTSEA

NO

N+H

HH S

S

Cys110

NHN

His104

NHN

His104

NHN

His104

SHCys110

HONO

A

B

WTor

C11xA

C13xA

T1-T1 T1-T1

T1-T1

by guest on August 11, 2019 http://www.jbc.org/ Downloaded from

Guangyu Wang, Candace Strang, Paul J. Pfaffinger and Manuel Covarrubias+-dependent redox switch in the intracellular T1-T1 interface of a Kv channel2Zn

published online March 1, 2007J. Biol. Chem. 

  10.1074/jbc.M609182200Access the most updated version of this article at doi:

 Alerts:

  When a correction for this article is posted• 

When this article is cited• 

to choose from all of JBC's e-mail alertsClick here

Supplemental material:

  http://www.jbc.org/content/suppl/2007/03/06/M609182200.DC1

by guest on August 11, 2019

http://ww

w.jbc.org/

Dow

nloaded from