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A Cysteine Scan of the Inner Vestibule of Cyclic Nucleotide–gated Channels Reveals Architecture and Rearrangement of the Pore

Galen E. Flynn¹ and William N. Zagotta^1,2

¹ Department of Physiology and Biophysics, University of Washington, Seattle, WA 98195
² Howard Hughes Medical Institute, University of Washington, Seattle, WA 98195

Address correspondence to William N. Zagotta, Department of Physiology and Biophysics, Box 357290, University of Washington, Seattle, WA 98195-7290. Fax: (206) 543-0934; E-mail: Zagotta{at}u.washington.edu

	ABSTRACT

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Cyclic nucleotide–gated (CNG) channels belong to the P-loop–containing family of ion channels that also includes KcsA, MthK, and Shaker channels. In this study, we investigated the structure and rearrangement of the CNGA1 channel pore using cysteine mutations and cysteine-specific modification. We constructed 16 mutant channels, each one containing a cysteine mutation at one of the positions between 384 and 399 in the S6 region of the pore. By measuring currents activated by saturating concentrations of the full agonist cGMP and the partial agonists cIMP and cAMP, we show that mutating S6 residues to cysteine caused both favorable and unfavorable changes in the free energy of channel opening. The time course of cysteine modification with 2-aminoethylmethane thiosulfonate hydrochloride (MTSEA) was complex. For many positions we observed decreases in current activated by cGMP and concomitant increases in current activated by cIMP and cAMP. A model where modification affected both gating and permeation successfully reproduced the complex time course of modification for most of the mutant channels. From the model fits to the time course of modification for each mutant channel, we quantified the following: (a) the bimolecular rate constant of modification in the open state, (b) the change in conductance, and (c) the change in the free energy of channel opening for modification of each cysteine. At many S6 cysteines, modification by MTSEA caused a decrease in conductance and a favorable change in the free energy of channel opening. Our results are interpreted within the structural framework of the known structures of KcsA and MthK. We conclude that: (a) MTSEA modification affects both gating and permeation, (b) the open configuration of the pore of CNGA1 channels is consistent with the structure of MthK, and (c) the modification of S6 residues disrupts the helical packing of the closed channel, making it easier for channels to open.

Key Words: cyclic nucleotide–gated channel • cysteine • protein structure • ion channel gating

	INTRODUCTION

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Ion channels are integral membrane proteins that regulate ion fluxes across membranes and thereby control membrane excitability. To regulate ion fluxes, channels undergo conformational changes that close and open their ion-permeable pore, a process referred to as gating. Gating is in turn allosterically regulated by changes in membrane voltage or binding of agonists. CNG channels are a prototypic ion channel gated by the binding of agonists. CNG channels are closed or opened in response to changes in the cytoplasmic concentrations of cyclic nucleotides (Yau and Baylor, 1989). Cyclic nucleotides bind directly to a cytoplasmic binding domain on the channel which, in turn, is allosterically coupled to the opening of the pore in these channels (Fesenko et al., 1985; Nakamura and Gold, 1987). When opened, CNG channels allow an influx of cations that depolarizes the plasma membrane. This is the regulatory mechanism that underlies the electrical response of photoreceptors and olfactory receptors to sensory stimuli (Burns and Baylor, 2001; Zufall and Munger, 2001). In addition, these channels are expressed in a number of other tissues where their function is less well characterized (for review see Kraus-Friedmann, 2000).

The first CNG channel cloned (CNGA1) was isolated from bovine rod photoreceptors (Kaupp et al., 1989). It shares sequence similarity with the voltage-gated ion channel superfamily, although CNG channels are only weakly voltage dependent (Yau and Baylor, 1989; Jan and Jan, 1990). Like voltage-gated potassium channels, CNG channels are composed of four subunits (Liu et al., 1996; Varnum and Zagotta, 1996). Each subunit contains cytoplasmic amino and carboxy termini, six transmembrane segments (S1–S6), a charged S4 region, and a P-loop between S5 and S6 (Kaupp et al., 1989; Molday et al., 1991; Wohlfart et al., 1992; Henn et al., 1995; Liu et al., 1996). Within each carboxy-terminal region, CNG channels have a cyclic nucleotide–binding domain (CNBD)^* that exhibits sequence similarity to other cyclic nucleotide-binding proteins (for review see Shabb and Corbin, 1992). Direct binding of cyclic nucleotides to this specialized domain leads to opening of the CNG channel pore.

CNG channel pores are thought to be structurally similar to that of other P-loop–containing ion channels (Goulding et al., 1993; Sun et al., 1996; Becchetti et al., 1999; Becchetti and Roncaglia, 2000; Liu and Siegelbaum, 2000; Flynn and Zagotta, 2001) (Fig. 1). The basic architectural plan of an ion channel pore was revealed by the crystal structure of KcsA, a bacterial potassium channel from Streptomyces lividans (Doyle et al., 1998). KcsA is a tetramer of identical subunits arranged with fourfold symmetry about a centrally located pore. A single KcsA subunit has two membrane-spanning helices, the outer and inner helices, and a reentrant P-loop. Each subunit of the tetramer contributes its P-loop to the formation of the pore. The P-loop starts from the extracellular side and enters the membrane as an -helix (pore helix) that then exits back extracellularly as an uncoiled strand. Within this strand, permeant cations are coordinated by the backbone carbonyl oxygens of the amino acids TVGYG, which are recognized as the signature sequence of K⁺-selective channels (Heginbotham et al., 1994). Intracellular to the selectivity filter is a large (10 Å in diameter; Doyle et al., 1998) water-filled vestibule. In the center of this cavity another permeant cation is stabilized through both electrostatic interactions with the pore helices and water molecules that hydrate the cation (Doyle et al., 1998; Roux and MacKinnon, 1999; Zhou et al., 2001). The membrane-spanning inner helices line the vestibule of the channel and cross the membrane at an angle to form a helix bundle on the intracellular side (Doyle et al., 1998). The helix bundle defines the intracellular entrance to the pore of KcsA that is thought to be in the closed state. To open the pore, a conformation change is thought to occur in the inner helix. An open conformation was revealed by the crystal structure of another ion channel, MthK (Jiang et al., 2002a,b). MthK is structurally similar to KcsA in the P-loop but exhibits a different conformation of the inner helix. In MthK, the inner helix contains a "gating hinge" near the top of the vestibule that bends this helix by 30°, creating a 12 Å opening on the intracellular side of the pore compared with the 4 Å opening of the helix bundle of KcsA. Amino acid conservation among a wide range of P-loop–containing channels suggests that the KcsA and MthK structures may serve as general models for the closed and open state conformations for this entire family of ion channels (Jiang et al., 2002b).

Using site-specific cysteine substitution, we have suggested previously that the cytoplasmic opening of the CNG channel pore is narrow when channels are closed and widens when channels open (Flynn and Zagotta, 2001). Substituting a cysteine (S399C, Fig. 1) at the cytoplasmic end of the S6 (the putative inner helix) in a cysteine-free variant of CNGA1 channels promotes channel closure through the spontaneous formation of an intersubunit disulfide bond. Since disulfide bonds are formed between cysteine residues 10 Å apart (Falke et al., 1988; Careaga and Falke, 1992), this result is consistent with a narrow cytoplasmic opening and the occurrence of a helix bundle similar to the one in KcsA. Furthermore, this disulfide bond forms much faster when channels are closed than when channels are open, suggesting a conformational change in the helix bundle of CNGA1 channels that widens the intracellular entrance of the pore. A widening of the intracellular entrance is necessary to explain the voltage-dependent block by large molecules such as tetrapentylammonium (TPeA) ions that enter the inner vestibule and block CNGA1 channels (Stotz and Haynes, 1996). Although the helix bundle defines a narrow cytoplasmic opening to the pore when channels are closed, its permeability to small cationic cysteine modifiers, such as Ag⁺ and 2-aminoethylmethane thiosulfonate hydrochloride (MTSEA), is state independent. These results are consistent with a model where the intracellular entrance of the pore of CNG channels widens during opening but is not itself the gate that controls permeation through the membrane (Flynn and Zagotta, 2001).

Several investigations have shown that the gating process in CNG channels is sensitive to perturbations of the pore structure (Sun et al., 1996; Bucossi et al., 1997; Fodor et al., 1997; Becchetti et al., 1999; Liu and Siegelbaum, 2000; Flynn et al., 2001). The sensitivity of the gating mechanism to mutational and biochemical perturbations provides useful information about the energetic and structural changes that occur during the final closed-to-open transition. In this study, we focused on the conformational change in the pore that is initiated by cyclic nucleotide binding to the CNBD. We replaced each residue in the S6 helix from 384 to 399 with cysteine (Fig. 1) and quantified changes in function in response to mutational and biochemical perturbations in the pore.

	MATERIALS AND METHODS

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Molecular Biology and Channel Expression
We introduced reporter cysteines into the S6 region of CNGA1_cys-free channels. CNGA1_cys-free is a cysteine-free variant of CNGA1 (Kaupp et al., 1989) with the mutations C35A, C169S, C186S, C314S, C481F, C505V, and C573V that remove all endogenous cysteine residues from the channel (Matulef et al., 1999). In addition, CNGA1_cys-free had a K2E mutation to create an Nco1 site and a FLAG^TM epitope (DYKDDDDK) that replaced the last five residues at the carboxy terminus. This construct was also used as the B protomer in tandem dimer experiments (see below). The cyclic nucleotide specificity of CNGA1_cys-free channels was the same as CNGA1 channels but the efficacies and apparent affinities for each cyclic nucleotide were higher primarily due to the C481F mutation (Matulef et al., 1999). For these experiments, PCR methods were used to replace a single codon in the cDNA encoding CNGA1_cys-free with the codon for cysteine. Each mutation was confirmed by DNA sequencing (PE Applied Biosystems, Inc.). 16 mutant channels were generated in total, one for each amino acid position between 384 and 399 in the CNGA1 channel protein. Nhe1-linearized cDNAs were transcribed in vitro to cRNAs using the T7 Message Machine kit (Ambion). For protein expression, Xenopus oocytes were injected with 40 nl of cRNA (1 µg/µl) and stored at 18°C for 4–10 d in a solution containing (in mM) 96 NaCl, 2 KCl, 1.8 CaCl₂, 1 MgCl₂, 5 HEPES at pH 7.6 (Zagotta et al., 1989). All chemicals were purchased from Sigma-Aldrich unless indicated otherwise.

For tandem dimers experiments, the coding sequences of two subunits (A and B protomers) were joined by a linker region in a single open reading frame and expressed in oocytes to form channels containing two A protomers and two B protomers (see previously described methods in Gordon and Zagotta, 1995b; Varnum et al., 1995). All A protomers included a COOH-terminal 21 amino acid peptide linker (Q₈IEGRQ₈A) followed by an Nco1 restriction site. All B protomers contained an Nco1 restriction site at the initial methionine. The entire coding sequence for the A protomer plus linker was excised and inserted into a pGEMHE vector containing the B protomer sequence. For heterodimeric S399C channel expression, the construct encoded a CNGA1_cys-free subunit (A protomer) connected to a S399C subunit (B protomer) separated by the peptide linker sequence. For homodimeric S399C channel expression, both A and B protomers contained the S399C mutation. DNA and mRNA were analyzed by agarose gel electrophoresis to confirm the presence of a single dimer-size construct before injection of mRNA into oocytes.

Electrophysiology
Electrophysiological data were collected from excised, inside-out patches of membrane from defolliculated oocytes using the patch-clamp technique (Hamill et al., 1981). All recordings were made at room temperature (20–22°C). Recording electrodes were fashioned from borosilicate glass (VWR International) that was polished to resistances of 0.2–1.0 m when filled with recording solutions. Inside each glass electrode, a 0.6% agar, 3 M KCl bridge was fit over the Ag-AgCl electrode wire to increase the stability of recordings. Electrodes were filled with a solution (extracellular) that contained (in mM): 130 NaCl, 3 HEPES, 0.5 niflumic acid, and 0.2 EDTA at pH 7.2. Niflumic acid was added to inhibit calcium-activated chloride currents endogenous to oocytes (White and Aylwin, 1990). The bath solution (intracellular) contained (in mM): 130 NaCl, 3 HEPES, and 0.2 EDTA at pH 7.2. To activate CNGA1_cys-free and cysteine mutant channels, saturating concentrations of cyclic nucleotides (2.5 mM cGMP, 16 mM cIMP, 16 mM cAMP) were applied to the intracellular surface of each patch using a rapid solution changer (RSC-100; Biologic). A three-step voltage protocol was applied to the patches that consisted of three 100 ms pulses to -60, +60, and -60 mV from a holding potential of 0 mV. Ionic currents were amplified and low-pass filtered at 2 kHz by an AxoPatch 200A (Axon Instruments, Inc.). The currents were digitized at 10 kHz using an ITC-16 board (Instrutech Corp.) interfaced to a Pentium 3 computer (Dell Computer Corp.) running Pulse software (HEKA Electronics, Inc.) and stored in files for offline analysis using Igro software (Wavemetrics, Inc.) or Excel software (Microsoft).

For all the experiments in this study, currents in the absence of cyclic nucleotide have been subtracted so that the data shown are only of currents through cyclic nucleotide-activated channels. After patch excision, we observed a slow increase in current through CNGA1_cys-free channels that required 20–25 min to stabilize. This increase in current has been attributed to dephosphorylation of the channel (Molokanova et al., 1999; Kramer and Molokanova, 2001). Most of our experiments were performed 30 min after patch excision to accommodate the increase in current that occurs after patch excision. A388C and S399C channels exhibited a spontaneous decline in current amplitude following patch excision. As a consequence, these mutants were examined immediately following patch excision and were analyzed relative to CNGA1_cys-free channels immediately after patch excision.

Cysteine Modification
We applied MTSEA (2-aminoethylmethane thiosulfonate hydrochloride) to the intracellular side of CNGA1_cys-free channels and each of the 16 mutant channels. Stock solutions of 100 mM MTSEA (Toronto Research Chemical) were made in water and stored at -80°C until just before application. MTSEA was diluted to the indicated concentration in the intracellular solution containing 2.5 mM cGMP. To study the kinetic effects of cysteine modification, mutant channels were partially modified by a brief exposure (10 s) to MTSEA applied to the intracellular side. Patches were then washed free of modifier and currents were activated by saturating concentrations of cGMP, cIMP, and cAMP. Measurements of cyclic nucleotide–activated currents were made at 2–3 ms after the initiation of +60- and -60-mV pulses in order to limit contamination by ion accumulation and depletion effects (Zimmerman et al., 1988). Currents in the absence of nucleotides were also measured. Patches were then reexposed to MTSEA and the cycle of solutions continued until no further changes in current were observed (see Fig. 6).

Determination of Mutational Effects on Gating
The effects on the free energy difference between the closed and open states caused by the cysteine mutation were quantified for each mutant channel. We assumed that the opening transition for a channel fully bound by agonist can be described by a simple closed-to-open equilibrium with equilibrium constant (L). The approach used actually measures resting-to-activated states but for simplicity will be referred to as closed and open states respectively. The behavior of the different agonists can be explained by L values specific for each agonist (L_cGMP, L_cIMP, and L_cAMP; Varnum et al., 1995; Varnum and Zagotta, 1996; Sunderman and Zagotta, 1999). The L values specific to each agonist were determined for each mutant channel assuming that the ratios L_cIMP/L_cGMP and L_cAMP/L_cGMP were fixed to 0.05 and 0.001, respectively, and were the same for all mutations. Currents activated by all three cyclic nucleotides were measured at both +60 and -60 mV. Analysis of the CNGA1_cys-free channels at both +60 and -60 mV indicated that the voltage dependence of L was very weak (e-fold per 460 ± 59 mV; n = 7). Therefore, we assumed that the L values were voltage independent. The maximum current when all channels are open (I_max) and L_cGMP was determined by minimizing the least squares difference between currents observed and currents calculated (cal) according to the following equations:

From these L values, the free energy difference between closed and open states was determined according to the relationship G_L,cNMP = -RTln(L_cNMP), where R is the ideal gas constant and T is temperature in Kelvin. All fits converged and the statistical results for each channel are shown as box plots or mean ± SEM as indicated.

Model for Determining the Modification Effects on Gating and Permeation
The rate of modification and the effects on gating and permeation were quantified by fitting a model to the time course of modification for each of the 16 cysteine mutant channels (see Fig. 8). Currents activated by all three cyclic nucleotides were measured at both +60 and -60 mV and plotted as a function of the cumulative exposure time in MTSEA. These time courses were fit with the equation below:

I(t) represents either I_cGMP, I_cIMP, or I_cAMP as a function of cumulative exposure time to MTSEA. L represents L_cGMP, L_cIMP, or L_cAMP as described above and was assumed to be voltage independent. I_max represents either I_{max,+60 mV} or I_{max,-60 mV} as described above. P is the probability that a subunit is modified and follows an exponential time course (P = 1 - e^(-t/)). The free parameters in this model represent the fractional change per subunit in the equilibrium constant of the opening transition (n), the fractional change in conductance per subunit (g), and time constant of modification per subunit (). All fits converged and the statistical results for each channel are shown as box plots or mean ± SEM as indicated.

Details for Analysis of Specific Mutants
A388C and S399C channels exhibited spontaneous declines in current amplitude after patch excision. In A388C channels, the decline in current was the result of desensitization since much of the current was recoverable after channels were closed for a period of time. However, a slow irreversible decline was also observed. The decline in S399C was attributed to the spontaneous formation of a disulfide bond (Flynn and Zagotta, 2001). Therefore, experiments on A388C and S399C channel were performed immediately after patch excision. Because we were unable to wait for the dephosphorylation of these two channels to reach steady-state before applying MTSEA, we analyzed these two channels relative to CNGA1_cys-free channels where experiments were performed immediately after patch excision.

Our analysis of the energetic effects on channel opening caused by cysteine mutations depends significantly on at least two cyclic nucleotides producing measurable currents with distinguishable amplitudes. G395C channels have such an unfavorable opening transition that the cGMP-activated currents were small and cIMP- and cAMP-activated currents were indiscernible from leak currents. However, MTSEA modification produced a dramatic increase in the currents activated by all three cyclic nucleotides and a pronounced inward rectification in G395C channels (see Fig. 12). Therefore, we used the modified channel to determined the maximum current (I_{max,-60 mV}) and the free energy of the opening transition (G_L,cGMP,MTSEA) at -60 mV after MTSEA modification. Based on the ratio of current activated at -60 mV by cGMP before MTSEA modification relative to I_{max,-60 mV} determined after modification, we calculated G_L,cGMP for G395C channels. In addition, for G392C channels after partial modification, currents activated by the three cyclic nucleotides were indistinguishable, and we were therefore unable to determine the change in the equilibrium constant caused by modification (n).

One of the limitations of the model is its inability to determine the number of subunits that are modified when the currents activated by all the cyclic nucleotides decay to zero with modification. This was the case for V384C, F387C, A388C, V391C, and G392C channels (see V384C Fig. 6). For these mutant channels we constrained the change in conductance (g) to 0.25.

The model describes effects of modification on gating and permeation and assumes that no change occurs in the cysteine residue other than MTSEA modification. However, we observed in G395C channels a behavior consistent with thiol-disulfide exchange when modification was slow relative to disulfide bond formation (see Fig. 15) (Creighton, 1984). To determine the effects of MTSEA modification on gating and permeation in G395C channels, currents activated by the three cyclic nucleotides were measured before and after rapid modification by 2 mM MTSEA. The free energy of the opening transition at -60 mV after modification (G_L,cGMP,MTSEA) was determined by simultaneously fitting the amplitudes of cyclic nucleotide–activated currents after rapid MTSEA exposure using the method of least squares. The energetic effects per subunit of G395C modification were calculated from the relationship G_L,MTSEA = (G_LcGMP,MTSEA - G_L,cGMP)/4 where G_L,cGMP was determine using the methods described above. Also, we calculated the change in conductance per subunit (g) as the ratio between the amplitude of the cGMP-activated current measured from modified G395C channels and the I_{max,+60 mV}, predicted from L_cGMP (see above), i.e., g = [1 - (I_cGMP,MTSEA/I_{max,+60 mV})]/4. The bimolecular rate constant of modification was not determined.

During the time course of MTSEA modification of S399C channels, the fractional activation by cIMP initially decreased and then increased, and the fractional activation by cAMP increased with a delay (see Fig. 16). This effect of modification on gating was different from the monotonic time course of the fractional activations of other S6 mutants in response to modification. Therefore, G_L,MTSEA for S399C channels could not be determined.

Mapping Function to Known Structures
Structural hypotheses were tested by mapping the parameters from the kinetic model onto two homology models of the CNGA1 S6 region made using the software Swiss-PDB viewer in conjunction with the Swiss-Model protein modeling server (Guex and Peitsch, 1997). The closed state of the CNGA1 pore region was modeled after KcsA (Doyle et al., 1998) while the open state was modeled after MthK (Jiang et al., 2002b). The angle of rotation made by each ß carbon in the inner helix of the KcsA and the MthK structures was measured relative to the central pore axis. A rotation angle was defined as the angle made by two lines when projected on a plane perpendicular to the pore axis; one connecting the ß carbon to the longitudinal axis of the inner helix and the second connecting the central pore axis to the longitudinal axis of the inner helix (Fig. 2 A; A108 shown). The second line represents an angle of 0° rotation. Angles of ß carbon rotation were measured for those KcsA and MthK residues that aligned with CNGA1 residues 384–399 (Fig. 1) . The angles of rotation were plotted versus amino acid number for both KcsA and MthK and a clear sinusoidal pattern of an helix was evident (Fig. 2 B). All structural models were analyzed and displayed using ViewerPro software (Accelrys).

View larger version (51K): [in this window] [in a new window]   FIGURE 1. Sequence alignment of the pore helix, selectivity filter, and inner helix for several channels of the P-loop–containing family. Amino acids in blue are conserved among several channels and amino acids in red are identical for all channels. The selectivity filter is highlighted by a yellow box, and inner helix residues that line the pore of KcsA are highlighted in gray boxes. The inner helix region between arrows shows residues that were substituted with cysteine in this study.

View larger version (40K): [in this window] [in a new window]   FIGURE 2. Angle of rotation away from the central pore axis of KcsA. (A) Side (top) and axial (bottom) view of the inner helix (ribbon) of KcsA in relation to the pore axis (red line, purple sphere). The pore axis (purple sphere) and the inner helix axis (gray sphere) are connected by a line (white) representing a 0° rotation angle. The ß carbon of alanine 108 in KcsA is shown in red along with the angle of rotation away from the pore axis (61°). (B) Beta carbon angles of KcsA residues 100–115 (squares) or MthK residues 84–98 (circles). Dashed line is a sine function fit to the data.

	RESULTS

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View larger version (21K): [in this window] [in a new window]   FIGURE 3. Effects of cysteine substitution on cyclic nucleotide-activated currents in CNGA1cys-free and three mutant channels. Representative currents traces from inside-out patches excised from Xenopus oocytes expressing either CNGA1cys-free or one of three mutant channels. Channels were activated by saturating concentrations of cGMP (green; 2.5 mM), cIMP (blue; 16 mM), cAMP (red; 16 mM). Currents were recorded in response to a three-step voltage protocol; 100-ms voltage steps to -60, +60 mV, and -60 mV from a holding potential of 0 mV (insert). The zero current level is indicated by a dashed line. Currents measured in the absence of cyclic nucleotides were subtracted from data shown. (A) CNGA1cys-free; bars: 1 nA, 20 ms. (B) M397C; bars: 1 nA, 20 ms. (C) T389C; bars: 50 pA, 20 ms. (D) I386C; bars: 50 pA, 20 ms.

SCHEME I

The values of these equilibrium constants for CNGA1_cys-free channels at 60 mV are shown in Table I . From the equilibrium constants, free energy differences between the closed and open states for CNGA1_cys-free channels were calculated for each cyclic nucleotide using the relation G_L,cNMP = -RTln(L_cNMP). These free energies are slightly lower (0.8 kcal/mol) than reported previously for CNGA1 channels and suggest that removal of endogenous cysteines caused a small stabilization in the opening transition but did not affect the agonist specificity (Varnum et al., 1995; Varnum and Zagotta, 1996; Matulef et al., 1999; Sunderman and Zagotta, 1999; Flynn and Zagotta, 2001; Johnson and Zagotta, 2001; Matulef and Zagotta, 2002).

For each of the mutant channels, we determined the energetic effect on opening. For these determinations, we assumed a fixed nucleotide specificity (G_L,cIMP - G_L,cGMP = 1.7 kcal/mol and G_L,cAMP - G_L,cGMP = 4.0 kcal/mol; Table I). These energies accurately account for differences in the amplitudes of cyclic nucleotide–activated currents of numerous mutant CNG channels where the mutations occurred at residues outside of the binding domain (Gordon and Zagotta, 1995a; Johnson and Zagotta, 2001). The maximum current (I_max) and the free energy difference of the opening transition in the presence of cGMP (G_L,cGMP) were determined by simultaneously fitting the amplitudes of cyclic nucleotide–activated currents with the above model (Scheme I) using the method of least squares. For G395C channels, cysteine replacement resulted in a channel with such an unfavorable opening transition that G_L,cGMP was calculated as described in MATERIALS AND METHODS. The G_L,cGMP values for each mutation were used to calculate the change in free energy of the closed-to-open transition caused by cysteine replacement using the relationship G_L,mutation = G_L,mutation - G_L,cys-free (Fig. 4) . As suggested by the apparent increase in the fractional activation by cIMP and cAMP in M397C channels, a cysteine at this site decreased the free energy difference of channel opening (-2.4 ± 0.06 kcal/mol, n = 5) shifting the equilibrium in favor of the open state. In contrast, a cysteine at T389C increased the free energy difference of channel opening (1.9 ± 0.062 kcal/mol, n = 4). For two positions, I390C and V391C (gray), a cysteine produced no significant difference in the channel opening transition compared with CNGA1_cys-free as determined by Student's t test (P > 0.2; for all other sites P < 0.005). From these data it is clear that mutations in the S6 perturbed the gating equilibrium in both directions, in favor of the open state (green) and in favor of the closed state (red) (Fig. 4).

View larger version (30K): [in this window] [in a new window]   FIGURE 4. Changes in the free energy of channel opening (GL,mutation) with cysteine replacement in the S6. Box plots of GL,mutation values for each mutant channel. For each box, the centerline is the median value, the edges of the boxes are the 25th and 75th percentiles, and the lines extending from the boxes are the 5th and 95th percentiles. Box plots were superimposed onto bar graphs extending for 0 to median GL,mutation for each mutant channel. Decreases (green) or increases (red) in GL,mutation relative to CNGA1cys-free are shown. Number of patches included in statistics is indicated in parentheses to the right of the mutant name.

View larger version (14K): [in this window] [in a new window]   FIGURE 5. Cyclic nucleotide–activated currents in CNGA1cys-free and V384C channels before and after modification by MTSEA. Representative currents traces are shown from CNGA1cys-free and V384C channels activated by saturating concentrations of cGMP (green; 2.5 mM), cIMP (blue; 16 mM), and cAMP (red; 16 mM) in response to a three step voltage protocol (see insert Fig. 3). (A and B) Representative currents are shown for CNGA1cys-free channels before and after 2 mM MTSEA was applied to the intracellular side while channels were held in the open state by 2.5 mM cGMP. The cumulative exposure time was 10 min. Bars: 100 pA, 20 ms (C and D) Representative currents are shown for V384C channels before and after 200 µM MTSEA was applied in the open state for a cumulative exposure time of 13 min. Bars: 100 pA, 20 ms.

View larger version (12K): [in this window] [in a new window]   FIGURE 6. Time course of cysteine modification in V384C channels. (A) Schematic diagram showing the solution-switching protocol for applying MTSEA in the open state of mutant channels. The intracellular side of a patch was exposed briefly to a solution of 2.5 mM cGMP containing MTSEA for 5–10 s. After each exposure, patches were washed in cGMP alone for 10 s. Currents were recorded in the presence of 2.5 mM cGMP, 16 mM cIMP, 16 mM cAMP, and 0 cNMP. Patches were exposed repeatedly to MTSEA for brief periods until no further change in current was observed. (B) Time course of modification by 200 µM MTSEA of V384C channels. Currents measured in cGMP (green square), cIMP (blue triangle), and cAMP (red diamond) at 60 mV were leak subtracted then plotted as a function of cumulative exposure time to MTSEA. Data are from the same patch as those shown in Fig. 5, C and D.

View larger version (26K): [in this window] [in a new window]   FIGURE 7. Complex kinetics of cysteine modification. (A) Representative currents are shown for N393C channels before (top) and after 2 min cumulative exposure to 20 µM MTSEA (middle). Plot of leak subtracted currents measured for each cyclic nucleotide recorded at 60 mV as a function of cumulative exposure time in MTSEA (bottom) is shown. Smooth lines through the data are from the model described in Fig. 8. Bars: 100 pA and 20 ms. (B) Currents recorded from S396C channels before and after 1 min cumulative exposure to 2 µM MTSEA. (C) Currents recorded from A388C channels before and after 100 s cumulative exposure to 20 µM MTSEA.

View larger version (18K): [in this window] [in a new window]   FIGURE 8. Kinetic model describing permeation and gating effects of cysteine modification. A model that predicts the time course of the changes in current for all three cyclic nucleotides caused by modifying each of four possible cysteines within the channel. The model is used to quantify the fractional change (n) in the equilibrium constant (L) between the closed (C) and open (O) states and the concomitant decrease in conductance through the open channel with each modification (m).

View larger version (18K): [in this window] [in a new window]   FIGURE 9. Effect of MTSET modification is reduced twofold in heterodimeric CNGA1cys-free/S399C channels. (A) Time course of MTSET modification of S399C/S399C homodimers and CNGA1cys-free/S399C heterodimers. A 2.5 mM cGMP solution containing 200 µM MTSET was applied to the intracellular side of each patch. Current amplitudes were normalized to the maximum cGMP-activated current for each patch. (B) Fraction of current modified by MTSET for homodimers and heterodimers. Bars show mean ± SEM (n = 3).

Open-state Accessibility of S6 Residues
One way to determine which residues line the pore is to quantify the accessibility of residues to cysteine modification. We investigated the open-state accessibility of the S6 residues by quantifying the second-order rate constants of MTSEA modification for each mutant channel. In the open state, S6 residues with side chains directed toward the center of the pore were predicted to modify more rapidly with MTSEA than residues with side chains directed away from the pore. From the time courses of modification of each mutant channel, time constants were determined from fits of our model to the time course data as described above. The inverse of was divided by the MTSEA concentration used to modify each mutant channel in order to obtain a second-order rate constant of modification for that site. Rate constants corresponding to each S6 site are plotted as box plots in Fig. 10 A. We found that most of the residues between 384 and 399 were accessible to MTSEA modification in the open state. Notable exceptions were L385C and I386C (white boxes) where the effects of MTSEA were not significantly different from that of CNGA1_cys-free channels (P > 0.05; Student's t test). Superimposed on these data is the sine function from Fig. 2 B that describes the angle of rotation of the ß carbons in KcsA and MthK relative to the pore axis. The data in Fig. 10 A are consistent with the sequence alignment shown in Fig. 1 and show that pore-lining residues were modified more rapidly by MTSEA then other S6 residues.

View larger version (47K): [in this window] [in a new window]   FIGURE 10. Rate constants of open-state cysteine modification for each S6 position. (A) Box plots of the bimolecular rate constants of MTSEA modification for each mutant channel. Dashed line is the sine function fit to the KcsA and MthK rotation angle data from Fig. 2. Colors correspond to median rate constants >104 M-1s-1 (red), between 103 M-1s-1 and 104/M/s (green), and between 101 M-1s-1 and 103 M-1s-1 (blue). Rate constants for L385C and I386C channels (white) were not statistically different from CNGA1cys-free (Student's t test, P > 0.6). (B) Homology models showing a bottom view of four S6 helices based on KcsA (top) and MthK (bottom), representing the closed and open conformations respectively. Sites corresponding to CNGA1 positions 384–399 are shown as space filled. Colors correspond to data in A.

MTSEA Modifies Permeation
Another way to determine which residues line the pore is to quantify the effects of modification on permeation. The largest conductance changes are expected for pore-lining residues from an electrostatic interaction between the positive charge on the modified cysteine and the permeant cations. One of the parameters of our model described the relative change in conductance caused by cysteine modification. In the model, we assumed that cysteine modification caused an identical change in all four subunits and that the effects of modification on conductance were additive. The fractional change in conductance per subunit due to modification (g) for each mutant channel is plotted relative to the amino acid number in Fig. 11 A. The sites with the largest MTSEA effect on the fractional conductance were those predicted by the sequence alignment and the homology models to project side chains toward the pore axis. When the g values from the kinetic model were mapped onto the homology models, the sites lining the pore were associated with the largest conductance changes (Fig. 11 B, red). These results are consistent with work published earlier describing the inhibitory effects of MTSET on some of these same mutant channels (Flynn and Zagotta, 2001). These effects on conductance were more indicative of pore-lining residues than the accessibility measurements. For example, MTSEA modification of the pore-lining residues F387C and V384C, which had slow rates, dramatically affected the conductance. Therefore, we conclude, from the data in Figs. 10 and 11, that the region between 384 and 399 of CNGA1 channels is an unbroken helix that lines the pore similar to KcsA and MthK. Perhaps more interestingly, pore-facing residues at the cytoplasmic end of the S6 (positions 395 and 399) had a somewhat smaller effect on conductance than expected for residues lining the pore. The 399 position in the KcsA homology model is within 4 Å from the central pore axis compared with 15 Å in the MthK homology model. The effect of modification on conductance at this position, therefore, is more consistent with a structural similarity between the open state of CNGA1 and MthK.

View larger version (50K): [in this window] [in a new window]   FIGURE 11. Changes in conductance due to MTSEA modification of S6 cysteines. (A) Box plots of the fractional conductance change per subunit (g) at 60 mV for each mutant channel determined from the model in Fig. 8. Dashed line is the sine function fit to the KcsA and MthK rotation angle data from Fig. 2. Colors correspond to median g values greater than 0.20 (red), between 0.10 and 0.20 (green), and <0.10 (blue). Because the rate constants of modification of L385C and I386C channels were not statistically different from CNGA1cys-free, box plots for these sites are shown in white. (B) Homology models showing sites color coded to correspond with data in A.

View larger version (49K): [in this window] [in a new window]   FIGURE 12. Changes in the free energy of channel opening (GL,MTSEA) with MTSEA modification of S6 cysteines. (A) Box plots and bar graphs of GL,MTSEA values per subunit determined for each mutant channel from the model in Fig. 8. Sites with a statistically significant decrease in the free energy change for the opening transition are shown in green. ND, not determined. (B) Homology models showing sites color coded to correspond with data in A.

View larger version (10K): [in this window] [in a new window]   FIGURE 13. MTSEA modification of G395C channels causes potentiation of cyclic nucleotide–activated currents. Representative currents for G395C channels before (A) and after (B) 2 mM MTSEA in 2.5 mM cGMP was applied to the intracellular side for 2 min. Bars: 200 pA, 20 ms.

View larger version (9K): [in this window] [in a new window]   FIGURE 14. Unliganded openings observed with large increases in the opening equilibrium constant after MTSEA modification. (A) Time course of modification by 20 µM MTSEA of A388C channels. Currents were measured at 60 mV, leak subtracted, and plotted as a function of cumulative exposure time to MTSEA. Shown are current amplitudes measured in 2.5 mM cGMP (IcGMP) (square) and current amplitudes measured in the absence of cyclic nucleotide (Isp) (circles). (B) Ratio Isp/IcGMP is plotted as a function of cumulative exposure time to MTSEA. Smooth line is a single exponential fit to the data of the equation Isp/IcGMP = 0.42 - 0.42e(-0.084t), where t is time.

View larger version (11K): [in this window] [in a new window]   FIGURE 15. Cysteine modification shows a high probability of unliganded openings for G392C channels. (A) Current amplitudes measured at +60 mV in the absence and presence of cyclic nucleotide. The solid line indicates the time when 2 mM MTSET in 2.5 mM cGMP was applied to the patch. The dashed line indicates the time when 2.5 mM cGMP was applied. Arrow indicates the change in current through unliganded channels. (B) Selected current traces showing the effects of MTSET modification on currents. Shown are currents in the absence of cyclic nucleotide (arrows 1 and 4) and in cGMP (arrows 2 and 3). Numbers correspond to points in A. Bars: 100 pA, 20 ms.

View larger version (15K): [in this window] [in a new window]   FIGURE 16. Partial MTSEA modification of G395C channels catalyzes thiol-disulfide exchange. (A) Current amplitudes measured at -60 mV while 2.5 mM cGMP was applied to the intracellular side of the patch (17 min, dashed line above the data). Currents were measured in 20 µM MTSEA in cGMP (solid line) until the effect reached steady-state. Amplitude of current measured in the absence of cyclic nucleotide is also shown. (B) Current amplitudes measured at -60 mV while 2.5 mM cGMP was applied (dashed line) to intracellular side of the patch. Currents were measured during a brief application (30 s, solid line) of 20 µM MTSEA in cGMP. (C) Reaction scheme depicting thiol-disulfide exchange.

View larger version (12K): [in this window] [in a new window]   FIGURE 17. Cysteine modification of S399C channels shows an additional gating component. (A) Time course of modification by 2 µM MTSEA of S399C channels. Shown are current amplitudes measured in 2.5 mM cGMP (green square), 16 mM cIMP (blue triangles), and 16 mM cAMP (red diamonds). (B) Ratios IcIMP/IcGMP and IcAMP/IcGMP are plotted as a function of cumulative exposure time to MTSEA.

	DISCUSSION

Top ABSTRACT INTRODUCTION MATERIALS AND METHODS RESULTS DISCUSSION REFERENCES

Cysteine Replacement in S6 Affects Gating
Mutations alone provide little direct information about protein structure. However, the sensitivity of the closed-to-open equilibrium to mutations does provide information about the structural elements involved in gating. We observed both favorable and unfavorable energetic effects on the closed-to-open equilibrium as a result of cysteine replacement between positions 384 and 399 in the S6 region of CNGA1 channels. The gating process is sensitive to mutational perturbations in most of the S6 helix, suggesting that the S6 is involved in gating. However, there was no pattern to these effects that could be correlated to any structural elements. It is difficult to make direct comparisons of the energetic effects caused by mutations between different positions when the perturbations are different for each positions, i.e., each substitution involves a different starting residue. In this study, a direct comparison is made possible by comparing the energetic effects of modification between positions where all the perturbations were the same, i.e., cysteine to cysteine + modifier.

Gating effects of S6 mutations are not unique to CNG channels. In both the six transmembrane and two transmembrane families of P-loop–containing channels, mutations in the cytoplasmic half of the inner helix have been reported to alter gating (Shyng et al., 1997; Lu et al., 1999; Enkvetchakul et al., 2000; Lippiat et al., 2000; Li-Smerin et al., 2000; Loussouarn et al., 2000; Espinosa et al., 2001; Panchenko et al., 2001; Sadja et al., 2001; Hackos et al., 2002; Jin et al., 2002; Lu et al., 2002; Yifrach and MacKinnon, 2002). Mutations in the inner helix have been shown to have both favorable and unfavorable energetic effects on the closed-to-open equilibrium. In the most extreme cases, the energetic effects were so favorable that the channels were constitutively active (Minor et al., 1999; Yi et al., 2001a; Yi et al., 2001b; Sadja et al., 2001; Zeidner et al., 2001; Hackos et al., 2002). We show here that G392C, and A388C after modification, also make the channels open in the absence of ligand. These results suggest that the inner helix is important for channel gating of many P-loop–containing channels.

Some of the cysteine mutations in the S6 produced novel gating effects. Currents through S399C channels spontaneously declined immediately after patch excision. We demonstrated previously that the decline in S399C current is due to the formation of a disulfide bond between the S399C residues from different subunits (Flynn and Zagotta, 2001). An intersubunit disulfide bond also formed in V391C channels, but required the presence of the mild oxidizing agent copper:phenanthroline (Flynn and Zagotta, 2001). Currents through G395C channels decline after a brief exposure to MTSEA. This unexpected result suggests a possible disulfide bond between G395C residues that is catalyzed by thiol-disulfide exchange (Creighton, 1984). All of these results are consistent with the prediction that V391C, G395C, and S399C are pointing toward the center of the pore. The distance between each of these sites on neighboring or diagonal subunits is <10 Å in the KcsA homology model, which is within a suitable range for disulfide bond formation (Creighton, 1984, 1993; Falke et al., 1988; Careaga and Falke, 1992). Another novel gating effect is observed in A388C channels. Currents through A388C channels spontaneously decline to a new steady-state level when the patch was exposed to a saturating cGMP solution (unpublished data). This decline in current was partially recoverable when the channels were closed by exposure to a cyclic nucleotide-free solution. Thus, A388C channels desensitized.

Changes in Gating and Permeation Occur as S6 Cysteines are Modified
Many of the cysteine mutant CNG channels exhibited a complex response to modification by positively charged MTSEA when applied in the open state. We observed a decrease in cGMP-activated currents and a concomitant increase in cIMP- and cAMP-activated currents in several mutant channels. Because of these disparate effects on CNG currents, we designed a kinetic model that describes the effects of modification in terms of alterations in both gating and permeation. The model predicts changes in current for all three cyclic nucleotides caused by modifying each of four possible cysteines within the channel. Three key assumptions in the model were (a) each modification produced an equal and identical change in the free energy difference between the closed and open states, (b) each modification produced an equal and identical change in the conductance, and (c) each modification occurs independently. The first of these three assumptions is reasonable and is the prediction of a concerted opening conformational change. A number of mutations have been observed to have energetic effects that are nearly additive when different numbers of subunits are mutated (Liu et al., 1996, 1998; Varnum and Zagotta, 1996). However, it has also been proposed that CNG channels gate by a dimer-of-dimer mechanism where the energy effects are nonadditive (Liu et al., 1998). The first assumption is also violated if disulfide bonds occur or if there are interactions between modifiers, such as electrostatic interactions between positive charges. The second of these three assumptions is not well supported theoretically; however, it has been shown previously that modification by 1, 2, 3, or 4 MTS reagents produced an approximately linear decreased in the single-channel conductance (Lu et al., 1999; Loussouarn et al., 2001). It is also supported by the observation that the current decline due to MTSET modification of S399C is twofold less in heterodimeric channels in which the number of modifiable cysteines is reduced by two relative to homodimeric channels. For the third assumption, one possible violation occurs when modification is state dependent and produces a significant change in the open probability of the channel. In this study, all modification was performed by applying MTSEA in combination with a 2.5 mM concentration of cGMP. The open probability of most of the mutant channels in 2.5 mM cGMP was near one, with the noted exception of G395C channels. In all cases where modification changed the open probability, the change increased an already high open probability for these channels. In addition, we have shown previously that residues deep in the pore (e.g., V391C) do not exhibit state-dependent modification by MTSEA (Flynn and Zagotta, 2001) and therefore state-dependent modification is not a significant issue. A second possible violation of the third assumption is that the rate of modification is affected by electrostatic or steric interactions from a previous modification. This seems likely for G395C and S399C channels where the model did not produce an adequate fit. However, the success of the model in fitting the data at most positions suggests that these assumptions are reasonable. These assumptions are required for the systematic analysis presented here, and we believe the qualitative conclusions reached in this study are not strongly dependent on the specifics of the model.

We used our model to analyze modification of cysteine residues introduced at positions 384–399 in the S6 of CNGA1 channels. By analyzing the effects of modification, we can directly compare the results from multiple positions with identical perturbations at each position. For each site, we determined three different properties of the modification: (a) the bimolecular rate of modification in the open state, (b) the change in conductance with modification, and (c) the change in the closed-to-open equilibrium with modification. In the open state, we observed modification rates that ranged over five orders of magnitude. The effects of modification on permeation ranged from little to complete, but were always negative thus resulting in a decrease in conductance. The effects of modification on the closed-to-open transition were variable but were always positive, thus increasing the cAMP-activated currents. The success of the model in fitting the data from a great majority of positions in the S6 suggests that the model provides a good description of the gating and permeation effects of modification.

Structural Homology with KcsA and MthK
CNGA1 channels share sequence similarity with KcsA and MthK channels. We have further confirmed a structural homology between these channels and CNGA1 channels by showing that CNGA1 residues predicted by these structures to point toward the center of the pore were those sites modified with the fastest rate constants (Fig. 10). We also found that changes in the fractional conductance per subunit caused by attaching a positively charged group to cysteine residues in the S6 region were largest for those sites predicted to point toward the center of the pore, as though the positively-charged cysteine interacted with the permeant ion (Fig. 11). Perhaps more interestingly, pore-facing residues at the cytoplasmic end of the S6 (positions 395 and 399) had a somewhat smaller effect on conductance than other residues lining the pore. This result is perhaps more consistent with a structural similarity between CNGA1 and MthK where the cytoplasmic end of the inner helix veers away from the central axis of the pore.

Nature of the Physical Changes in the S6 of CNGA1
The MthK structure was solved in the presence of a calcium ligand that shifts the opening transition of these channels to favor the open state (Jiang et al., 2002a,b). Superimposition of KcsA and MthK suggests a structural model for the closed-to-open transition in the inner helices; with KcsA being closed and MthK being open. The physical model suggests that in the closed state the inner helices are close together, forming a helical bundle on the cytoplasmic side of the channel (Doyle et al., 1998). In the open state, the inner helices are separated by a bend at a conserved glycine disrupting the helical bundle and widening the entrance to the pore (Jiang et al., 2002b). This glycine hinge is just one residue upstream of the region we investigated and may mark a site within CNG channels for a break or bend in the S6 helix.

A model where the inner helices separate during channel opening is consistent with our results. MTSEA modification of many S6 sites resulted in a decrease in free energy for channel opening (Fig. 12). This decrease might be caused if MTSEA disrupted the helical packing of the closed channel, making it easier for the channels to open. We have shown previously that S399C forms an intersubunit disulfide bond when channels are closed but not when channels are open (Flynn and Zagotta, 2001). This intersubunit disulfide bond might lock the cytoplasmic ends of two S6 helices in the closed configuration (Flynn and Zagotta, 2001). In the KcsA structure, the residues corresponding to S399C are close together (within 5–8 Å), whereas in MthK structure the residues are predicted to be far apart (separate by 20–30 Å). The closed and open conformations of CNGA1 channels therefore may be similar to KcsA and MthK, respectively.

Conclusions
By perturbing the S6 region through mutations and biochemical modifications, we have shown the importance of this region as part of the gating mechanism for CNG channels and gained some insights into the structure and rearrangement of the pore. We conclude that: (a) modification effects both gating and permeation, (b) the open configuration of the pore of CNGA1 channels is consistent with the structure of MthK, and (c) the modification of the S6 disrupts the helical packing of the closed channel making it easier for the channels to open.

	FOOTNOTES

 
^* Abbreviations used in this paper: CNBD, cyclic nucleotide–binding domain; IRK, inward rectifier; MTSEA, 2-aminoethylmethane thiosulfonate hydrochloride; MTSET, 2-trimethylammonioethylmethane thiosulfonate hydrochloride; TPeA, tetrapentylammonium.

	ACKNOWLEDGMENTS

 
We would like to thank Drs. Matthew C. Trudeau, Jie Zheng, and Leon D. Islas for comments on this manuscript. We wish to thank Dr. Jos Van Schagen for helpful discussions and for computer algorithms that calculate coupled differential equations. We wish to acknowledge the members of the Zagotta lab who have contributed in many ways to the success of these experiments, especially the technical expertise of Kevin Black, Shellee Cunnington, Gay Sheridan, and Heidi Utsugi.

This work was funded by National Eye Institute (EY10329) and the Howard Hughes Medical Institute. W.N. Zagotta is an associate investigator within the Howard Hughes Medical Institute.

Olaf S. Andersen served as editor.

Submitted: 14 February 2003
Revised: 23 April 2003
Accepted: 2 May 2003

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