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Communication

Deglycosylation of Shaker KV channels affects voltage sensing and the open–closed transition

Angelica Lopez-Rodriguez  Correspondence email, Miguel Holmgren  Correspondence email
DOI: 10.1085/jgp.201711958 | Published June 7, 2018

Abstract

Most membrane proteins are subject to posttranslational glycosylation, which influences protein function, folding, solubility, stability, and trafficking. This modification has been proposed to protect proteins from proteolysis and modify protein–protein interactions. Voltage-activated ion channels are heavily glycosylated, which can result in up to 30% of the mature molecular mass being contributed by glycans. Normally, the functional consequences of glycosylation are assessed by comparing the function of fully glycosylated proteins with those in which glycosylation sites have been mutated or by expressing proteins in model cells lacking glycosylation enzymes. Here, we study the functional consequences of deglycosylation by PNGase F within the same population of voltage-activated potassium (KV) channels. We find that removal of sugar moieties has a small, but direct, influence on the voltage-sensing properties and final opening–closing transition of Shaker KV channels. Yet, we observe that the interactions of various ligands with different domains of the protein are not affected by deglycosylation. These results imply that the sugar mass attached to the voltage sensor neither represents a cargo for the dynamics of this domain nor imposes obstacles to the access of interacting molecules.

Introduction

N-glycoproteins are highly abundant within the proteome (Apweiler et al., 1999). Most membrane proteins, as they travel along the secretory pathway, are exposed to a diverse and complex set of sequential glycosylation reactions that have important roles in protein folding and other biological functions (Varki, 1993; Moremen et al., 2012). A wide range of ion channels, including voltage-activated Na+, Ca2+, and K+ (KV); acetylcholine receptors; transient receptor potential channels; and cyclic nucleotide–gated channels, have been shown to be glycosylated (Covarrubias et al., 1989; Recio-Pinto et al., 1990; Deal et al., 1994; Santacruz-Toloza et al., 1994; Schwalbe et al., 1995; Gurnett et al., 1996; Thornhill et al., 1996; Castillo et al., 1997; Petrecca et al., 1999; Shi and Trimmer, 1999; Zhang et al., 1999; Freeman et al., 2000; Pabon et al., 2000; Ufret-Vincenty et al., 2001a; Zhu et al., 2001, 2003; Bennett, 2002; Much et al., 2003; Faillace et al., 2004; Sutachan et al., 2005; Wirkner et al., 2005; Meighan et al., 2013; Watanabe et al., 2015).

Voltage-activated K+ (KV) channels are involved in shaping action potentials as well as tuning the pattern of electrical signals in neurons (Hille, 2001). The membrane core of these channels is formed by four identical or homologous α subunits that assemble as a complex that contains a central ion permeation pathway and four peripheral voltage-sensor domains (VSDs; Long et al., 2005). Each α subunit comprises six transmembrane segments from which the first four (S1–S4) constitute the VSD and the last two (S5–S6) constitute the pore. Diversity of this family of ion channels derives from the large number of members (Southan et al., 2016) as well as posttranscriptional (Holmgren and Rosenthal, 2015) and posttranslational modifications, such as phosphorylation (Drain et al., 1994; Ivanina et al., 1994; Levin et al., 1996) and glycosylation (Deal et al., 1994; Santacruz-Toloza et al., 1994; Schwalbe et al., 1995; Thornhill et al., 1996; Petrecca et al., 1999; Shi and Trimmer, 1999; Freeman et al., 2000; Khanna et al., 2001; Ufret-Vincenty et al., 2001b; Zhu et al., 2001, 2003; Watanabe et al., 2003, 2004, 2007, 2015; Napp et al., 2005; Sutachan et al., 2005; Fujita et al., 2006; Johnson and Bennett, 2008; Hall et al., 2015).

Glycosylation in KV channels influences cell surface expression and stability (Petrecca et al., 1999; Khanna et al., 2001; Watanabe et al., 2004, 2015; Fujita et al., 2006; Hall et al., 2015) and in some instances changes the functional properties of the channel (Thornhill et al., 1996; Freeman et al., 2000; Ufret-Vincenty et al., 2001b; Watanabe et al., 2003, 2007; Napp et al., 2005; Sutachan et al., 2005; Johnson and Bennett, 2008). Many members of the KV1 channel family are glycosylated within the VSD. Because the movements of this domain are coupled to opening of the channel, its glycosylation has been shown to influence the speed of channel opening and the steady-state voltage dependence of activation (Thornhill et al., 1996; Watanabe et al., 2003, 2007; Johnson and Bennett, 2008). Here, we set to determine the direct influence of sugar moieties attached to the VSD on the structural dynamics of this domain. Because the functional consequences of glycosylation in ion channels have commonly been assessed by comparing different populations of ion channels (e.g., glycosylated channels with N→Q mutant channels or expression of channels in cell lines with distinct restrictions in the glycosylation process), we implemented an approach that directly assesses the role of glycosylation on the VSD function within the same population of channels. We studied the consequences of deglycosylation by PNGase F on the gating currents (Bezanilla, 2000) and the binding/unbinding of hanatoxin, a voltage-sensor toxin (Swartz, 2007). We found that deglycosylation of Shaker KV channels occurs in minutes, giving rise to ion channels with rightward displaced charge distribution on the voltage axis. This shift was not accompanied by changes in the kinetics of the gating currents, consistent with the notion that sugars attached to the VSD do not represent a cargo to the dynamics of this domain. Nonetheless, as previously reported (Thornhill et al., 1996; Watanabe et al., 2003, 2007; Johnson and Bennett, 2008), a reduction in the speed of activation at positive potentials and deactivation at negative potentials was observed from macroscopic ionic currents after PNGase F treatment, indicating that sugars might influence the C↔O transition. To isolate this transition, we introduced the triple-mutant ILT into our Shaker KV construct (Smith-Maxwell et al., 1998a,b). Deglycosylation by PNGase F to these channels also slowed the opening of these channels. These results combined suggest that glycosylation has a direct influence on the opening and closing of KV channels. Finally, we found that glycosylation does not alter the interactions of either gating-modifier toxins or pore blockers with the Shaker KV channels.

Materials and methods

Protein extraction and Western blotting

Xenopus laevis oocytes were injected with cRNA of noninactivating (Δ6–46) Shaker KV channels (Hoshi et al., 1990) that contained a c-myc epitope (Glu-Gln-Lys-Leu-Ile-Ser-Glu-Glu-Asp-Leu) inserted at the C terminus (after Val638) and incubated in ND96 solution (in mM: 96 NaCl, 2 KCl, 1 MgCl2, 1.8 CaCl2, and 5 HEPES, pH 7.6) at 17°C. 24 h after injection, oocytes were exposed to 10 mg/ml PNGase F diluted 1:10 in ND96. The enzyme activity was monitored by removing 10 oocytes at different times, supplemented with 100 mM glycine and lysed in 200 µl buffer H (1% Triton X-100, 100 mM NaCl, and 20 mM Tris-HCl, pH 7.4). Lysates were rocked at room temperature for 15 min and then centrifuged at 13,000 rpm for 3 min. The pellet was discarded, and the supernatant was analyzed by Western blot by using anti–Myc mouse monoclonal antibody at a dilution of 1:5,000 (Clontech) and detected with a secondary, goat anti–mouse antibody conjugated to horseradish peroxidase at a dilution of 1:10,000 (Pierce). Membranes were developed by SuperSignal WestFemto (Thermo Fisher Scientific) and visualized by chemiluminescence by using a FluorChem E Imager (Cell Biosciences).

Electrophysiological recordings

Xenopus oocytes were injected with in vitro transcripts of three different Shaker KV channel constructs: noninactivating (Δ6–46) Shaker KV channels (Hoshi et al., 1990) to determine the voltage dependence of the relative probability of opening, hanatoxin-sensitive Shaker KV channels (Milescu et al., 2013) to assess toxin binding, and nonconductive Shaker KV channels W434F (Perozo et al., 1993) to measure gating currents. Oocytes were incubated at 17°C for 24–48 h in ND96 solution before recordings. Currents were acquired before treatment, then PNGase F was applied for 5 min in the recording chamber and washed out thoroughly before subsequent recordings.

Two-microelectrode voltage-clamp currents were acquired with an OC-725C oocyte amplifier (Warner Instruments) and digitized by using a Digidata 1321A interface and pCLAMP 10.3 software (Axon Instruments). Data were filtered at 1 kHz and digitized at 10 kHz. Microelectrode resistances were between 0.1 and 1.2 MΩ when filled with 3 M KCl. Solutions for recording contained 50 mM KCl, 50 mM NaCl, 1 mM MgCl2, 0.3 mM CaCl2, and 5 mM HEPES, pH 7.4. Voltage steps to +40 mV in 10-mV increments were given from a holding potential of −80 mV, which were returned to −50 mV by using a P/−4 subtracting protocol. Toxins were applied for 5 min before acquisitions were initiated.

For currents acquired by using a cut-open voltage-clamp oocyte technique (Taglialatela et al., 1992), we exclusively used the animal pole for recording. Oocytes were clamped with a Dagan CA-1B high-performance oocyte clamp. Data were acquired at 10 kHz and filtered at 5 kHz. A 0.2- to 0.3-MΩ pipette and ground-pool were filled with 3M Tris HCl, while bridges were filled with 3M Na-MES in 3% agarose. Oocytes were permeabilized with 0.4% saponin in internal solution. For ionic current experiments, the internal solution contained (in mM): 110 potassium glutamate, 10 HEPES-NMG, and 10 EGTA-NMG, pH 7.3, and the external solution (in mM): 2.5 KCl, 120 NaMES, 1.8 CaCl2, and 10 HEPES-NMG, pH 7.6. Gating-current experiments were performed with the same external solution, but the internal solution contained (in mM): 120 NMG-glutamate, 10 HEPES-NMG, and 10 EGTA-NMG, pH 7.3. A P/−4 subtracting protocol was used. Toxins used in this study were provided by K. Swartz (National Institutes of Health, Bethesda, MD).

Data analyzing

A weighted time constant was calculated when a double-exponential fit was used. The time constant of the activating or deactivating phases of ionic currents was determined by using distinct exponential fits.

PNGase F extraction

Origami (DE3) Escherichia coli competent cells transformed with pET22b (PNGase F) construct were grown at 37°C until absorbance at 600 nm reached 1.0 in Lysogeny broth medium supplemented with 15 mg/ml kanamycin, 50 mg/ml ampicillin, and 12.5 mg/ml tetracycline. The culture was then induced with 0.5 mM isopropyl β-D-1 thiogalactopyranoside and grown in a shaker incubator overnight at 20°C. Cells were harvested, washed with Tris buffered saline, and resuspended in lysis buffer (50 mM Tris, pH 8.0, 150 mM NaCl, 1 mM PMSF, 0.5 mM MgCl2, 0.5 mg/ml lysozyme, and 25 μg/ml deoxyribonuclease I). Cells were lysed by using EmulsiFlex-C5 (Avestin). The cell lysate was ultracentrifuged at 40,000 rpm for 1 h by using a TL45 rotor (Beckman). The soluble fraction was loaded on a 5-ml Ni-NTA prepacked column equilibrated with 150 mM NaCl, 10 mM imidazole, and 50 mM HEPES, pH 8.0, and eluted with a gradient from 10 to 500 mM imidazole. Peak fractions were dialyzed with two changes by using a storage buffer containing 150 mM NaCl, 50 mM HEPES, pH 8.0, and 10% glycerol. The purified protein was concentrated and flash frozen until use. Preliminary experiments were performed with a PNGase F gift from M. Mayer (National Institutes of Health, Bethesda, MD).

Results

PNGase F enzymatic activity in Xenopus oocytes

To assess the temporal course of PNGase F enzymatic activity, we incubated 70 Xenopus oocytes expressing Shaker KV channels with 1 mg/ml PNGase F and proceeded to remove 10 oocytes at different times (Fig. 1). The WT Shaker KV channel expressed in oocytes has two predominant glycosylation products (Santacruz-Toloza et al., 1994): a small (∼70-kD) form and a large (∼100-kD) form (Fig. 1, line t0). Even within 3 min of PNGase F exposure, most of the Shaker KV channel protein is deglycosylated, migrating as a smaller (∼65-kD) band (Fig. 1, lines 3–30), similar to Shaker KV channels in which their glycosylation sites have been mutated (Santacruz-Toloza et al., 1994). These results show that PNGase F acts rapidly, providing an amenable opportunity to study the role of glycosylation on the function of the channels in the same population of proteins.

Figure 1.
Figure 1.

Temporal course of PNGase F action on Shaker KV channel. Western blot showing the reduction of molecular weight after enzymatic digestion of the sugar moiety with PNGase F (1 mg/ml; n = 5).

Fig. 2 A (control) shows a family of K+ currents in response to voltage protocol pulses from −80 mV to +40 mV, every 20 mV. Immediately after acquiring these data, the oocyte was exposed to PNGase F for 5 min to allow complete enzymatic digestion. After removal of the enzyme from the recording chamber, the oocyte was subjected to the same voltage protocol (Fig. 2 B). Shaker KV channels clearly activated and deactivated more slowly after PNGase F exposure. The time the currents took to rise from 10% to 90% increased by approximately two- to threefold after deglycosylation (Fig. 2 C). Deglycosylation also slowed deactivation, with the time the tail currents took to decline to 10% increased from 14.7 ± 1.8 ms to 20.7 ± 2.6 ms before and after deglycosylation, respectively. These results are comparable to previous observations obtained from Shaker KV or rat KV1.1 channels expressed in mammalian cell lines (Thornhill et al., 1996; Watanabe et al., 2003; Johnson and Bennett, 2008). In those instances, however, functional comparisons were performed with different populations of channels. For example, the population of deglycosylated channels derived from channels in which the glycosylated sites have been mutated or from WT channels expressed in mammalian cell lines in which glycosylation has been impaired. Even though the activation and deactivation kinetics of deglycosylated channels changed, the relative probability of opening at different voltage is similar (Fig. 2 D).

Figure 2.
Figure 2.

Effects of deglycosylation on the ionic currents carried by Shaker KV channels. (A and B) Ionic currents from an oocyte expressing Shaker KV channels before (A) and after (B) 5-min exposure to PNGase F. Ionic currents were recorded by using the two-microelectrode voltage-clamp technique. These currents were elicited by 50-ms voltage steps from a holding potential of −80 to +40 mV, every 10 mV. (C) Time rise from 10% to 90% current activation. (D) Relative conductance plots before (black) and after (red) PNGase F treatment. Error bars in C and D represent mean ± SEM, (n = 5). DeGly, deglycosylation.

Because we externally applied PNGase F to the oocytes, deglycosylation should have occurred in all glycoproteins at the cell surface. To assess whether the removal of sugars from the Shaker KV channels has a direct influence on function of the protein, we developed a Shaker KV construct with both glycosylation sites mutated to aspartate (N259D–N263D), the end product at the site of cleavage by the enzyme (Tarentino and Plummer, 1994). Expressing this Shaker KV construct in untreated oocytes would imply that all native glycoproteins remain intact while the N259D–N263D Shaker KV mutant channels lack sugar moieties. Fig. 3 shows that the rise times from 10% to 90% of the current activation of these channels are comparable to those from WT channels treated with PNGase F. As a comparison, we also plotted the rise times of a Shaker KV construct with the traditional mutations (N259Q–N263Q) to study glycosylation and protein function (Fig. 3, blue triangles). PNGase F treatment to oocytes expressing these glycosylation-deficient Shaker KV channels did not alter the kinetics of activation or deactivation (Fig. 4, B and C). These results combined provide support to the idea that the functional consequences of deglycosylation of WT KV channels are specific to the removal of sugars from the channels.

Figure 3.
Figure 3.

Activation from glycosylation-deficient mutant Shaker KV channels. Rise time from 10% to 90% current activation from N259D–N263D mutant Shaker KV channels (n = 3) and N259Q–N263Q mutant Shaker KV channels (n = 3) and, for comparative purposes, those after PNGase F treatment. These data derived from ionic currents recorded using the two-microelectrode voltage clamp technique. Error bars represent mean ± SEM. DeGly, deglycosylation.

Figure 4.
Figure 4.

PNGase F treatment of glycosylation-deficient mutant Shaker KV channels. (A) Normalized and superimposed ionic current traces from WT Shaker KV channels before and after PNGase F treatment, shown for comparative purposes. (B and C) Normalized and superimposed ionic current recordings before and after PNGase F treatment of oocytes expressing glycosylation-deficient N259D–N263D Shaker KV channels (B, n = 3) and N259Q–N263Q Shaker KV channels (C, n = 3). These experiments were performed with the same batch of PNGase F enzyme. These ionic currents were elicited in response to a voltage step to +60 mV from a holding potential of −80 mV and were recorded by using the cut-open oocyte voltage-clamp technique. DeGly, deglycosylation.

Effect of deglycosylation on gating currents

We next asked whether sugar removal affects the motions of the VSD. The VSD is formed by the first four transmembrane segments of the channel (S1–S4). Within the S4, there are several arginines that are responsible for sensing voltage across the membrane (Aggarwal and MacKinnon, 1996; Seoh et al., 1996). In response to a voltage step, these charges rearrange resulting in nonlinear currents known as the gating currents (Bezanilla, 2000). Because the two glycosylation sites in Shaker KV channels are within the VSD (Santacruz-Toloza et al., 1994), it is conceivable that the sugar mass attached to the VSD might influence its motions. We measured gating currents using a nonconductive (W434F) Shaker KV channel construct (Perozo et al., 1993) using the cut-open voltage-clamp technique (Taglialatela et al., 1992). Fig. 5 A shows superimposed gating current traces in response to four voltage steps acquired from the same oocyte before (black) and after (red) PNGase F treatment. Two effects of deglycosylation were observed: (1) a reduction of 18 ± 4% (n = 7) of the total amount of charge (Fig. 5 B) and (2) a small shift of ∼6 mV toward positive potentials of the steady-state charge distribution (Fig. 5, B–D). Interestingly, this shift was not accompanied by changes in the kinetics of the gating currents (Fig. 5 A), as can be appreciated by the relaxations of the “on” gating currents. Further, the early development of the rising phase in the “off” currents, which is observed at −48 mV before glycosylation, is apparent after deglycosylation at about −44 mV.

Figure 5.
Figure 5.

Effect of deglycosylation on gating currents from the nonconducting (W434F) mutant Shaker KV channel. (A) Gating currents from an oocyte expressing nonconductive Shaker KV channels before (black) and after (red) 5-min exposure to PNGase F. Gating currents were elicited in response to voltage steps from −100 mV to the voltage shown on the traces by using the cut-open oocyte voltage clamp. (B) Voltage dependence of the gating charge before and after PNGase F treatment from the experiment shown in A. Solid lines represent Boltzmann fits. The best parameter values for the total amount of charge (Qtot) and the midpoint voltage (V1/2) were 2.42 nC and −55.4 mV before treatment and 1.94 nC and −48.7 mV after PNGase F treatment, respectively. The reduction in Qtot was observed in all experiments. It progresses with a slow time constant (∼60 min) to ∼30% of the initial value of Qtot. The reason for this response is unknown presently, but it appears to be specific for the nonconducting W434F mutant channel. (C) Normalized voltage dependence of the steady-state charge distribution. The plot shows the charge distribution before (black symbols) and after (red symbols) PNGase F treatment from seven experiments. In all experiments, the V1/2 shifted to the right in the voltage axes. (D) Box plot of the changes in V1/2 from the seven experiments.

Effect of deglycosylation on the activation and deactivation of ILT mutant channels

That PNGase F treatment produced little functional consequences in the movement of the voltage sensor suggests that the effects of deglycosylation on the activation and deactivation of the WT Shaker KV channels might originate from changes in the opening and closing events. To isolate this transition, we used the ILT mutant Shaker KV channels, which shift both the relative probability of opening by ∼80 mV toward positive potentials (Smith-Maxwell et al., 1998a,b) and the steady-state charge distribution of the gating currents toward negative potentials by ∼40 mV (Ledwell and Aldrich, 1999). Fig. 6 A shows ionic currents from ILT mutant channels before and after PNGase F treatment at activation voltages of +100 and +120 mV from a holding potential of −100 mV. Clearly, deglycosylation slowed the opening of these channels. The rise times from 10% to 90% of the current activation changed by 5.4 ± 1.0 ms at +100 mV and 4.7 ± 1.1 ms at +120 mV (n = 4). ILT channels close fast at −100 mV (Fig. 6), requiring nearly perfect linear capacity subtraction to have an appreciation of the effect of deglycosylation on closing. Like WT channels, the changes in deactivation are small, yet observed in all four experiments, two of them shown in Fig. 6 B.

Figure 6.
Figure 6.

Effect of deglycosylation on ionic currents from the ILT mutant Shaker KV channel. (A) Normalized ionic current traces elicited from a holding potential of −100 mV to +100 mV (left) and +120 mV (right), before (black) and after (red) PNGase F treatment. (B) Normalized tail currents before (black) and after (red) PNGase F treatment upon return from the voltage step to +120 mV. Left: Zoomed-in traces depicted in A (+120 mV). Right: Traces from a different oocyte. Ionic currents from ILT channels were P/6 subtracted with subpulses of opposite polarity from the holding potential. These experiments were performed by using the cut-open oocyte voltage clamp. DeGly, deglycosylation.

Effect of deglycosylation on the binding of ligands

In addition to changes in protein function, glycosylation might interfere directly with the binding and unbinding of ligands that interact with the protein in question. We tested the consequences of deglycosylation of Shaker KV channels with three ligands: hanatoxin (HaTx), agitoxin (AgTx), and 4-amino pyridine (4AP). HaTx interacts with the VSD (Swartz and MacKinnon, 1997a,b; Li-Smerin and Swartz, 2000, 2001; Phillips et al., 2005; Herrington et al., 2006; Milescu et al., 2007, 2009, 2013), whereas the remaining two bind to the pore domain: AgTx to the exterior pore, and 4AP to the internal pore (Kirsch and Drewe, 1993; Kirsch et al., 1993; Garcia et al., 1994; Gross and MacKinnon, 1996).

To test HaTx, we used a Shaker KV construct in which five residues in the third transmembrane segment were mutated to the corresponding amino acids of KV2.1 (Shaker Δ5; Milescu et al., 2013) to make the Shaker KV channel sensitive to HaTx (Milescu et al., 2013). Fig. 7 A shows current recordings before and after exposure to 200 nM HaTx in the same oocyte. The voltage steps shown are −50, −40, 0, and +40 mV from a holding potential of −80 mV. In the presence of HaTx, ionic currents could be elicited at more negative potentials compared with the control, consistent with the toxin stabilizing the open state (Milescu et al., 2013). Fig. 7 B shows a comparable experiment with a different oocyte, except that before HaTx exposure, the oocyte was treated with PNGase F. Similar to untreated KV Shaker channels, HaTx increased the open probability at negative potentials so ionic current could be detected at −50 and −40 mV. The changes by HaTX on the relative probability of opening are also comparable for untreated and PNGase F–treated channels (Fig. 7 C).

Figure 7.
Figure 7.

Effect of deglycosylation on the binding of Hanatoxin. (A and B) Current traces represent ionic currents in response to voltage steps to −50, −40, 0, and +40 mV from a holding potential of −80 mV. Tail currents shown were in response to voltage of −50 mV. In A, 200 nM HaTx was tested on unmodified channels (n = 5) and in B after deglycosylation by PNGase F (n = 4). (C) Relative probability of opening estimated from the tail currents. Data in the presence of HaTx were normalized with respect to the control (black and gray symbols) or immediately after PNGase F treatment (red and cyan symbols). These experiments were performed by using the two-microelectrode voltage-clamp technique. Error bars represent mean ± SEM. DeGly, deglycosylation.

Fig. 8 shows the consequences of deglycosylation on the blockade of Shaker KV channels by either AgTx (100 nM; Fig. 8 A) or 4AP (0.1 mM; Fig. 8 B). In both cases, these pore blockers inhibited similarly untreated and PNGase F–treated channels.

Figure 8.
Figure 8.

Effect of deglycosylation on the binding of pore blockers. (A and B) Current traces represent ionic currents in response to voltage steps to −50, −40, 0, and +40 mV from a holding potential of −80 mV. Tail currents shown were in response to voltage of −50 mV. A compares the inhibition by 100 nM AgTx before (top) and after (bottom) PNGase F treatment (n = 2). At +40 mV, AgTx blocked 48% of the ionic current before and 62% after treatment, respectively. At 0 mV, it inhibited 48% of the ionic current before and 63% treatment, respectively. B compares the inhibition by 0.1 mM 4AP before (top) and after (bottom) PNGase F treatment (n = 2). At +40 mV, 4AP blocked 55% of the ionic current before and 53% after treatment, respectively. At 0 mV, it inhibited 48% of the ionic current before and 50% after treatment, respectively. These experiments were performed by using the two-microelectrode voltage-clamp technique. DeGly, deglycosylation.

Discussion

Many KV channels are known to be subject to glycosylation as they traffic toward the cell membrane. In some instances, glycosylation influences cell surface expression (Petrecca et al., 1999; Khanna et al., 2001; Watanabe et al., 2004, 2007, 2015; Fujita et al., 2006; Hall et al., 2015), whereas in others it might also impact protein function (Schwalbe et al., 1995; Thornhill et al., 1996; Freeman et al., 2000; Ufret-Vincenty et al., 2001b; Watanabe et al., 2003, 2007; Napp et al., 2005; Sutachan et al., 2005; Johnson and Bennett, 2008). Shaker KV channel contains two glycosylation sites within the VSD of the protein (Santacruz-Toloza et al., 1994). In a previous study (Johnson and Bennett, 2008), the kinetics of Shaker KV channels expressed in CHO cell lines with either normal or deficient glycosylation were shown to be altered by the extent of sugar incorporation into the channel. Here, we aimed to assess whether glycosylation has a direct influence on the function of Shaker KV channels’ VSD, the domain where the glycans are attached.

We studied the role of sugars on the channel’s function by assessing their presence and absence within the same population of ion channels. We chose PNGase F to remove sugars because it acts within minutes (Fig. 1), and the Asn at the cleavage site becomes an Asp, which has the potential to restore the electrostatic nature of the sugars removed, and its influence can be directly assessed by site-directed mutagenesis. There are some advantages of this approach. For example, (a) in some instances, mutation of glycosylation sites might not be a viable alternative because of failures in cell membrane expression; (b) the small functional consequences observed in the gating currents (Fig. 5, C and D) would have been extremely difficult to discern comparing two populations of channels; and (c) the effect on the Qtot from the gating currents would have remained undetected by analyzing two populations of channels. Yet, the method needs to be carefully controlled because PNGase F treatment is expected to act on many native glycosylated proteins that might indirectly influence the protein under study. Further, this approach is limited to the study of the presence or not of sugar moieties; therefore, the role of partial glycosylation is inaccessible.

We found that deglycosylation by PNGase F slows down channel activation and deactivation with an undetected shift in the relative probability of opening, suggesting that the effect of deglycosylation on the rates of opening and closing is similar. This is similar to previous observations in which KV channels were expressed in cell lines with different degrees of glycosylation capabilities (Thornhill et al., 1996; Johnson and Bennett, 2008) or when functional comparisons were made between WT and glycosylation-deficient N→Q mutant channels expressed in the same cell line (Watanabe et al., 2003, 2007). To obtain direct evidence of the effect of glycans on the VSDs of KV channels, we assessed the influence of deglycosylation by PNGase F on the gating currents. The steady-state distribution of the gating currents is shifted by 6 mV toward more depolarized potentials after deglycosylation (Fig. 5). This shift appears to be a pure voltage bias to the charge moving transitions carrying no changes in the kinetics of the gating currents. This implies that the mass of the sugar attached to the VSD does not influence the sensors’ movements in response to voltage steps.

The functional consequences observed by PNGase F treatment of WT Shaker KV channels likely originated from the direct removal of sugar moieties from the channels. First, the activation kinetics of glycosylation-deficient mutant Shaker KV (N259Q–N263Q) and Shaker KV (N259D–N263D) are slower than WT channels (Fig. 3), comparable to those measured after WT channels were deglycosylated. And second, PNGase F treatment of glycosylation-deficient mutant Shaker KV channels produces no changes in the opening and closing of the channels (Fig. 4), indicating that the functional consequences observed with WT channels are not mediated by the deglycosylation of a native membrane protein that could have been interacting with Shaker KV channels.

The changes in channels’ opening and closing by deglycosylation cannot be explained by the shift in the steady-state distribution of the VSD transitions, suggesting they should originate from a direct effect on the C↔O transition. We isolated this transition by introducing the triple ILT mutations in the VSD. This channel construct has a distribution of the relative probability of opening that is shifted by ∼80 mV toward positive potentials, and a charge distribution of the gating currents shifted ∼40 mV in the opposite direction (Smith-Maxwell et al., 1998a,b; Ledwell and Aldrich, 1999). Deglycosylation by PNGase F had qualitatively similar consequences on the opening and closing of ILT channels (Fig. 6), like those observed in WT channels. These results lead to the conclusion that sugar moieties attached to the S1–S2 linker influence the motions of the pore domain accompanying the opening and closing of the channels. By using kinetic modeling of ionic currents derived from mammalian KV1.1 WT and the glycosylation-deficient mutant channel KV1.1 (N207Q), the opening transition C→O had to be reduced approximately threefold to account for the slowdown in activation (Watanabe et al., 2003). As in Shaker KV channels, mammalian KV1.1 are glycosylated within the VSD at the linker between transmembrane segments S1–S2.

The association between the VSDs and the C↔O transition in Shaker KV channels had initially been focused to regions of the VSD directly involved in charge movement steps. For example, the ILT mutations in the transmembrane segment S4 produced a substantial shift (∼40 mV) of the charge movement distribution, which allowed a practical separation between the VSD movements and the C↔O transition (Smith-Maxwell et al., 1998a,b; Ledwell and Aldrich, 1999). This ILT mutant construct has also been used to show that the dynamics of the S4 are directly linked to the movements of the S6 that determine the C↔O transition (Pathak et al., 2005) as well as to establish that HaTx binding to the VSDs can directly stabilize the open state of the channel (Milescu et al., 2013). Additionally, by bridging the S4 and S5 transmembrane segments with metal ions (Lainé et al., 2003) and studying the state dependence of bridge formation, it was demonstrated they formed in the open state (Phillips and Swartz, 2010). However, by using statistical coupling analysis, it has also been proposed that the external end of transmembrane segment S1 forms an interface with the pore helix of KV channels (Lee et al., 2009). Therefore, it is conceivable that sugars bound to the S1–S2 linker might influence the state of this interface to determine the kinetics of the C↔O transition. In fact, mutations of some residues at the external end of transmembrane segment S5, the pore helix, and their connecting linker have shown substantial changes in the C↔O transition (Yifrach and MacKinnon, 2002). Further, it has recently been reported noncanonical interactions between adjacent VSDs and pore domains indicate a far more extensive functional relationship of these domains than structures predict (Carvalho-de-Souza and Bezanilla, 2017).

The presence of sugars at the external side of Shaker KV channels does not appear to possess a stearic hindrance to access and binding of interacting molecules such as toxins and blockers of both VSD and the pore region (Figs. 7 and 8). The lack of changes in the interactions of HaTx-targeting VSDs by deglycosylation is compatible with the general mechanism of action of this toxin. On the one hand, HaTx access their binding site through the membrane (Lee and MacKinnon, 2004; Phillips et al., 2005; Milescu et al., 2007), so sugars at the external side of the protein are not in the path of voltage-sensor toxins. On the other hand, HaTx interacts with the surface of the VSD delineated by S3 and S4 transmembrane segments (Milescu et al., 2007, 2009), which are facing the membrane at the opposite direction as S1 and S3. Even though it is not surprising that the interactions of pore-blocking molecules such as AgTx and 4AP are not influenced by glycans, these results indicate that the ∼20-kD sugar moiety has a rather confined surface of interaction with the channel.

Acknowledgments

We thank Mark Mayer and Kenton Swartz for providing the PNGase F and toxins used in this work and Joseph Mindell, Deepa Shrikumar, Chhavi Mathur, and Pablo Miranda-Fernandez for technical support. We also thank Kenton Swartz for reading the manuscript.

This work was supported by The Intramural Research Program of the National Institute of Neurological Disorders and Stroke, National Institutes of Health to M. Holmgren.

The authors declare no competing financial interests.

Author contributions: A. Lopez-Rodriguez and M. Holmgren conceived and designed the experiments, performed the experiments, analyzed the data, contributed reagents, materials, and analysis tools, and wrote the paper.

Richard W. Aldrich served as editor.

  • Submitted: 29 November 2017
  • Revision received 23 April 2018
  • Accepted: 15 May 2018
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References

  1. ↵
    1. Aggarwal, S.K., and
    2. R. MacKinnon
    . 1996. Contribution of the S4 segment to gating charge in the Shaker K+ channel. Neuron. 16:1169–1177. doi:10.1016/S0896-6273(00)80143-9
    OpenUrlCrossRefPubMed
  2. ↵
    1. Apweiler, R.,
    2. H. Hermjakob, and
    3. N. Sharon
    . 1999. On the frequency of protein glycosylation, as deduced from analysis of the SWISS-PROT database. Biochim. Biophys. Acta. 1473:4–8. doi:10.1016/S0304-4165(99)00165-8
    OpenUrlCrossRefPubMed
  3. ↵
    1. Bennett, E.S.
    2002. Isoform-specific effects of sialic acid on voltage-dependent Na+ channel gating: Functional sialic acids are localized to the S5-S6 loop of domain I. J. Physiol. 538:675–690. doi:10.1113/jphysiol.2001.013285
    OpenUrlCrossRefPubMed
  4. ↵
    1. Bezanilla, F.
    2000. The voltage sensor in voltage-dependent ion channels. Physiol. Rev. 80:555–592. doi:10.1152/physrev.2000.80.2.555
    OpenUrlCrossRefPubMed
  5. ↵
    1. Carvalho-de-Souza, J., and
    2. F. Bezanilla
    . 2017. Non-canonical interactions between voltage sensors and pore domain in Shaker K+-channel. Biophys. Journal. doi:10.1016/j.bpj.2016.11.892
    OpenUrlCrossRef
  6. ↵
    1. Castillo, C.,
    2. M.E. Díaz,
    3. D. Balbi,
    4. W.B. Thornhill, and
    5. E. Recio-Pinto
    . 1997. Changes in sodium channel function during postnatal brain development reflect increases in the level of channel sialidation. Brain Res. Dev. Brain Res. 104:119–130. doi:10.1016/S0165-3806(97)00159-4
    OpenUrlCrossRefPubMed
  7. ↵
    1. Covarrubias, M.,
    2. C. Kopta, and
    3. J.H. Steinbach
    . 1989. Inhibitors of asparagine-linked oligosaccharide processing alter the kinetics of the nicotinic acetylcholine receptor. J. Gen. Physiol. 93:765–783. doi:10.1085/jgp.93.5.765
    OpenUrlAbstract/FREE Full Text
  8. ↵
    1. Deal, K.K.,
    2. D.M. Lovinger, and
    3. M.M. Tamkun
    . 1994. The brain Kv1.1 potassium channel: In vitro and in vivo studies on subunit assembly and posttranslational processing. J. Neurosci. 14:1666–1676. doi:10.1523/JNEUROSCI.14-03-01666.1994
    OpenUrlAbstract/FREE Full Text
  9. ↵
    1. Drain, P.,
    2. A.E. Dubin, and
    3. R.W. Aldrich
    . 1994. Regulation of Shaker K+ channel inactivation gating by the cAMP-dependent protein kinase. Neuron. 12:1097–1109. doi:10.1016/0896-6273(94)90317-4
    OpenUrlCrossRefPubMed
  10. ↵
    1. Faillace, M.P.,
    2. R.O. Bernabeu, and
    3. J.I. Korenbrot
    . 2004. Cellular processing of cone photoreceptor cyclic GMP-gated ion channels: a role for the S4 structural motif. J. Biol. Chem. 279:22643–22653. doi:10.1074/jbc.M400035200
    OpenUrlAbstract/FREE Full Text
  11. ↵
    1. Freeman, L.C.,
    2. J.J. Lippold, and
    3. K.E. Mitchell
    . 2000. Glycosylation influences gating and pH sensitivity of I(sK). J. Membr. Biol. 177:65–79. doi:10.1007/s002320001100
    OpenUrlCrossRefPubMed
  12. ↵
    1. Fujita, T.,
    2. I. Utsunomiya,
    3. J. Ren,
    4. Y. Matsushita,
    5. M. Kawai,
    6. S. Sasaki,
    7. K. Hoshi,
    8. T. Miyatake, and
    9. K. Taguchi
    . 2006. Glycosylation and cell surface expression of Kv1.2 potassium channel are regulated by determinants in the pore region. Neurochem. Res. 31:589–596. doi:10.1007/s11064-006-9056-4
    OpenUrlCrossRefPubMed
  13. ↵
    1. Garcia, M.L.,
    2. M. Garcia-Calvo,
    3. P. Hidalgo,
    4. A. Lee, and
    5. R. MacKinnon
    . 1994. Purification and characterization of three inhibitors of voltage-dependent K+ channels from Leiurus quinquestriatus var. hebraeus venom. Biochemistry. 33:6834–6839. doi:10.1021/bi00188a012
    OpenUrlCrossRefPubMed
  14. ↵
    1. Gross, A., and
    2. R. MacKinnon
    . 1996. Agitoxin footprinting the shaker potassium channel pore. Neuron. 16:399–406. doi:10.1016/S0896-6273(00)80057-4
    OpenUrlCrossRefPubMed
  15. ↵
    1. Gurnett, C.A.,
    2. M. De Waard, and
    3. K.P. Campbell
    . 1996. Dual function of the voltage-dependent Ca2+ channel alpha 2 delta subunit in current stimulation and subunit interaction. Neuron. 16:431–440. doi:10.1016/S0896-6273(00)80061-6
    OpenUrlCrossRefPubMed
  16. ↵
    1. Hall, M.K.,
    2. D.A. Weidner,
    3. M.A. Edwards, and
    4. R.A. Schwalbe
    . 2015. Complex N-glycans influence the spatial arrangement of voltage gated potassium channels in membranes of neuronal-derived cells. PLoS One. 10:e0137138. doi:10.1371/journal.pone.0137138
    OpenUrlCrossRef
  17. ↵
    1. Herrington, J.,
    2. Y.P. Zhou,
    3. R.M. Bugianesi,
    4. P.M. Dulski,
    5. Y. Feng,
    6. V.A. Warren,
    7. M.M. Smith,
    8. M.G. Kohler,
    9. V.M. Garsky,
    10. M. Sanchez, et al.
    2006. Blockers of the delayed-rectifier potassium current in pancreatic beta-cells enhance glucose-dependent insulin secretion. Diabetes. 55:1034–1042. doi:10.2337/diabetes.55.04.06.db05-0788
    OpenUrlAbstract/FREE Full Text
  18. ↵
    1. Hille, B.
    2001. Ion Channels of Excitable Membranes. Sinauer Associates, Inc., Sunderland, MA. 814 pp.
  19. ↵
    1. Holmgren, M., and
    2. J.J. Rosenthal
    . 2015. Regulation of ion channel and transporter function through RNA editing. Curr. Issues Mol. Biol. 17:23–36.
    OpenUrl
  20. ↵
    1. Hoshi, T.,
    2. W.N. Zagotta, and
    3. R.W. Aldrich
    . 1990. Biophysical and molecular mechanisms of Shaker potassium channel inactivation. Science. 250:533–538. doi:10.1126/science.2122519
    OpenUrlAbstract/FREE Full Text
  21. ↵
    1. Ivanina, T.,
    2. T. Perets,
    3. W.B. Thornhill,
    4. G. Levin,
    5. N. Dascal, and
    6. I. Lotan
    . 1994. Phosphorylation by protein kinase A of RCK1 K+ channels expressed in Xenopus oocytes. Biochemistry. 33:8786–8792. doi:10.1021/bi00195a021
    OpenUrlCrossRefPubMed
  22. ↵
    1. Johnson, D., and
    2. E.S. Bennett
    . 2008. Gating of the shaker potassium channel is modulated differentially by N-glycosylation and sialic acids. Pflugers Arch. 456:393–405. doi:10.1007/s00424-007-0378-0
    OpenUrlCrossRefPubMed
  23. ↵
    1. Khanna, R.,
    2. M.P. Myers,
    3. M. Lainé, and
    4. D.M. Papazian
    . 2001. Glycosylation increases potassium channel stability and surface expression in mammalian cells. J. Biol. Chem. 276:34028–34034. doi:10.1074/jbc.M105248200
    OpenUrlAbstract/FREE Full Text
  24. ↵
    1. Kirsch, G.E., and
    2. J.A. Drewe
    . 1993. Gating-dependent mechanism of 4-aminopyridine block in two related potassium channels. J. Gen. Physiol. 102:797–816. doi:10.1085/jgp.102.5.797
    OpenUrlAbstract/FREE Full Text
  25. ↵
    1. Kirsch, G.E.,
    2. C.C. Shieh,
    3. J.A. Drewe,
    4. D.F. Vener, and
    5. A.M. Brown
    . 1993. Segmental exchanges define 4-aminopyridine binding and the inner mouth of K+ pores. Neuron. 11:503–512. doi:10.1016/0896-6273(93)90154-J
    OpenUrlCrossRefPubMed
  26. ↵
    1. Lainé, M.,
    2. M.C. Lin,
    3. J.P. Bannister,
    4. W.R. Silverman,
    5. A.F. Mock,
    6. B. Roux, and
    7. D.M. Papazian
    . 2003. Atomic proximity between S4 segment and pore domain in Shaker potassium channels. Neuron. 39:467–481. doi:10.1016/S0896-6273(03)00468-9
    OpenUrlCrossRefPubMed
  27. ↵
    1. Ledwell, J.L., and
    2. R.W. Aldrich
    . 1999. Mutations in the S4 region isolate the final voltage-dependent cooperative step in potassium channel activation. J. Gen. Physiol. 113:389–414. doi:10.1085/jgp.113.3.389
    OpenUrlAbstract/FREE Full Text
  28. ↵
    1. Lee, S.Y., and
    2. R. MacKinnon
    . 2004. A membrane-access mechanism of ion channel inhibition by voltage sensor toxins from spider venom. Nature. 430:232–235. doi:10.1038/nature02632
    OpenUrlCrossRefPubMed
  29. ↵
    1. Lee, S.Y.,
    2. A. Banerjee, and
    3. R. MacKinnon
    . 2009. Two separate interfaces between the voltage sensor and pore are required for the function of voltage-dependent K(+) channels. PLoS Biol. 7:e47. doi:10.1371/journal.pbio.1000047
    OpenUrlCrossRefPubMed
  30. ↵
    1. Levin, G.,
    2. D. Chikvashvili,
    3. D. Singer-Lahat,
    4. T. Peretz,
    5. W.B. Thornhill, and
    6. I. Lotan
    . 1996. Phosphorylation of a K+ channel alpha subunit modulates the inactivation conferred by a beta subunit. Involvement of cytoskeleton. J. Biol. Chem. 271:29321–29328. doi:10.1074/jbc.271.46.29321
    OpenUrlAbstract/FREE Full Text
  31. ↵
    1. Li-Smerin, Y., and
    2. K.J. Swartz
    . 2000. Localization and molecular determinants of the Hanatoxin receptors on the voltage-sensing domains of a K+ channel. J. Gen. Physiol. 115:673–684. doi:10.1085/jgp.115.6.673
    OpenUrlAbstract/FREE Full Text
  32. ↵
    1. Li-Smerin, Y., and
    2. K.J. Swartz
    . 2001. Helical structure of the COOH terminus of S3 and its contribution to the gating modifier toxin receptor in voltage-gated ion channels. J. Gen. Physiol. 117:205–218. doi:10.1085/jgp.117.3.205
    OpenUrlAbstract/FREE Full Text
  33. ↵
    1. Long, S.B.,
    2. E.B. Campbell, and
    3. R. Mackinnon
    . 2005. Crystal structure of a mammalian voltage-dependent Shaker family K+ channel. Science. 309:897–903. doi:10.1126/science.1116269
    OpenUrlAbstract/FREE Full Text
  34. ↵
    1. Meighan, S.E.,
    2. P.C. Meighan,
    3. E.D. Rich,
    4. R.L. Brown, and
    5. M.D. Varnum
    . 2013. Cyclic nucleotide-gated channel subunit glycosylation regulates matrix metalloproteinase-dependent changes in channel gating. Biochemistry. 52:8352–8362. doi:10.1021/bi400824x
    OpenUrlCrossRefPubMed
  35. ↵
    1. Milescu, M.,
    2. J. Vobecky,
    3. S.H. Roh,
    4. S.H. Kim,
    5. H.J. Jung,
    6. J.I. Kim, and
    7. K.J. Swartz
    . 2007. Tarantula toxins interact with voltage sensors within lipid membranes. J. Gen. Physiol. 130:497–511. doi:10.1085/jgp.200709869
    OpenUrlAbstract/FREE Full Text
  36. ↵
    1. Milescu, M.,
    2. F. Bosmans,
    3. S. Lee,
    4. A.A. Alabi,
    5. J.I. Kim, and
    6. K.J. Swartz
    . 2009. Interactions between lipids and voltage sensor paddles detected with tarantula toxins. Nat. Struct. Mol. Biol. 16:1080–1085. doi:10.1038/nsmb.1679
    OpenUrlCrossRefPubMed
  37. ↵
    1. Milescu, M.,
    2. H.C. Lee,
    3. C.H. Bae,
    4. J.I. Kim, and
    5. K.J. Swartz
    . 2013. Opening the shaker K+ channel with hanatoxin. J. Gen. Physiol. 141:203–216. doi:10.1085/jgp.201210914
    OpenUrlAbstract/FREE Full Text
  38. ↵
    1. Moremen, K.W.,
    2. M. Tiemeyer, and
    3. A.V. Nairn
    . 2012. Vertebrate protein glycosylation: Diversity, synthesis and function. Nat. Rev. Mol. Cell Biol. 13:448–462. doi:10.1038/nrm3383
    OpenUrlCrossRefPubMed
  39. ↵
    1. Much, B.,
    2. C. Wahl-Schott,
    3. X. Zong,
    4. A. Schneider,
    5. L. Baumann,
    6. S. Moosmang,
    7. A. Ludwig, and
    8. M. Biel
    . 2003. Role of subunit heteromerization and N-linked glycosylation in the formation of functional hyperpolarization-activated cyclic nucleotide-gated channels. J. Biol. Chem. 278:43781–43786. doi:10.1074/jbc.M306958200
    OpenUrlAbstract/FREE Full Text
  40. ↵
    1. Napp, J.,
    2. F. Monje,
    3. W. Stühmer, and
    4. L.A. Pardo
    . 2005. Glycosylation of Eag1 (Kv10.1) potassium channels: intracellular trafficking and functional consequences. J. Biol. Chem. 280:29506–29512. doi:10.1074/jbc.M504228200
    OpenUrlAbstract/FREE Full Text
  41. ↵
    1. Pabon, A.,
    2. K.W. Chan,
    3. J.L. Sui,
    4. X. Wu,
    5. D.E. Logothetis, and
    6. W.B. Thornhill
    . 2000. Glycosylation of GIRK1 at Asn119 and ROMK1 at Asn117 has different consequences in potassium channel function. J. Biol. Chem. 275:30677–30682. doi:10.1074/jbc.M005338200
    OpenUrlAbstract/FREE Full Text
  42. ↵
    1. Pathak, M.,
    2. L. Kurtz,
    3. F. Tombola, and
    4. E. Isacoff
    . 2005. The cooperative voltage sensor motion that gates a potassium channel. J. Gen. Physiol. 125:57–69. doi:10.1085/jgp.200409197
    OpenUrlCrossRefPubMed
  43. ↵
    1. Perozo, E.,
    2. R. MacKinnon,
    3. F. Bezanilla, and
    4. E. Stefani
    . 1993. Gating currents from a nonconducting mutant reveal open-closed conformations in Shaker K+ channels. Neuron. 11:353–358. doi:10.1016/0896-6273(93)90190-3
    OpenUrlCrossRefPubMed
  44. ↵
    1. Petrecca, K.,
    2. R. Atanasiu,
    3. A. Akhavan, and
    4. A. Shrier
    . 1999. N-linked glycosylation sites determine HERG channel surface membrane expression. J. Physiol. 515:41–48. doi:10.1111/j.1469-7793.1999.041ad.x
    OpenUrlCrossRefPubMed
  45. ↵
    1. Phillips, L.R., and
    2. K.J. Swartz
    . 2010. Position and motions of the S4 helix during opening of the Shaker potassium channel. J. Gen. Physiol. 136:629–644. doi:10.1085/jgp.201010517
    OpenUrlAbstract/FREE Full Text
  46. ↵
    1. Phillips, L.R.,
    2. M. Milescu,
    3. Y. Li-Smerin,
    4. J.A. Mindell,
    5. J.I. Kim, and
    6. K.J. Swartz
    . 2005. Voltage-sensor activation with a tarantula toxin as cargo. Nature. 436:857–860. doi:10.1038/nature03873
    OpenUrlCrossRefPubMed
  47. ↵
    1. Recio-Pinto, E.,
    2. W.B. Thornhill,
    3. D.S. Duch,
    4. S.R. Levinson, and
    5. B.W. Urban
    . 1990. Neuraminidase treatment modifies the function of electroplax sodium channels in planar lipid bilayers. Neuron. 5:675–684. doi:10.1016/0896-6273(90)90221-Z
    OpenUrlCrossRefPubMed
  48. ↵
    1. Santacruz-Toloza, L.,
    2. Y. Huang,
    3. S.A. John, and
    4. D.M. Papazian
    . 1994. Glycosylation of shaker potassium channel protein in insect cell culture and in Xenopus oocytes. Biochemistry. 33:5607–5613. doi:10.1021/bi00184a033
    OpenUrlCrossRefPubMed
  49. ↵
    1. Schwalbe, R.A.,
    2. Z. Wang,
    3. B.A. Wible, and
    4. A.M. Brown
    . 1995. Potassium channel structure and function as reported by a single glycosylation sequon. J. Biol. Chem. 270:15336–15340. doi:10.1074/jbc.270.25.15336
    OpenUrlAbstract/FREE Full Text
  50. ↵
    1. Seoh, S.A.,
    2. D. Sigg,
    3. D.M. Papazian, and
    4. F. Bezanilla
    . 1996. Voltage-sensing residues in the S2 and S4 segments of the Shaker K+ channel. Neuron. 16:1159–1167. doi:10.1016/S0896-6273(00)80142-7
    OpenUrlCrossRefPubMed
  51. ↵
    1. Shi, G., and
    2. J.S. Trimmer
    . 1999. Differential asparagine-linked glycosylation of voltage-gated K+ channels in mammalian brain and in transfected cells. J. Membr. Biol. 168:265–273. doi:10.1007/s002329900515
    OpenUrlCrossRefPubMed
  52. ↵
    1. Smith-Maxwell, C.J.,
    2. J.L. Ledwell, and
    3. R.W. Aldrich
    . 1998a. Role of the S4 in cooperativity of voltage-dependent potassium channel activation. J. Gen. Physiol. 111:399–420. doi:10.1085/jgp.111.3.399
    OpenUrlAbstract/FREE Full Text
  53. ↵
    1. Smith-Maxwell, C.J.,
    2. J.L. Ledwell, and
    3. R.W. Aldrich
    . 1998b. Uncharged S4 residues and cooperativity in voltage-dependent potassium channel activation. J. Gen. Physiol. 111:421–439. doi:10.1085/jgp.111.3.421
    OpenUrlAbstract/FREE Full Text
  54. ↵
    1. Southan, C.,
    2. J.L. Sharman,
    3. H.E. Benson,
    4. E. Faccenda,
    5. A.J. Pawson,
    6. S.P. Alexander,
    7. O.P. Buneman,
    8. A.P. Davenport,
    9. J.C. McGrath,
    10. J.A. Peters, et al. NC-IUPHAR
    . 2016. The IUPHAR/BPS Guide to PHARMACOLOGY in 2016: Towards curated quantitative interactions between 1300 protein targets and 6000 ligands. Nucleic Acids Res. 44(D1):D1054–D1068. doi:10.1093/nar/gkv1037
    OpenUrlCrossRefPubMed
  55. ↵
    1. Sutachan, J.J.,
    2. I. Watanabe,
    3. J. Zhu,
    4. A. Gottschalk,
    5. E. Recio-Pinto, and
    6. W.B. Thornhill
    . 2005. Effects of Kv1.1 channel glycosylation on C-type inactivation and simulated action potentials. Brain Res. 1058:30–43. doi:10.1016/j.brainres.2005.07.050
    OpenUrlCrossRefPubMed
  56. ↵
    1. Swartz, K.J.
    2007. Tarantula toxins interacting with voltage sensors in potassium channels. Toxicon. 49:213–230. doi:10.1016/j.toxicon.2006.09.024
    OpenUrlCrossRefPubMed
  57. ↵
    1. Swartz, K.J., and
    2. R. MacKinnon
    . 1997a. Hanatoxin modifies the gating of a voltage-dependent K+ channel through multiple binding sites. Neuron. 18:665–673. doi:10.1016/S0896-6273(00)80306-2
    OpenUrlCrossRefPubMed
  58. ↵
    1. Swartz, K.J., and
    2. R. MacKinnon
    . 1997b. Mapping the receptor site for hanatoxin, a gating modifier of voltage-dependent K+ channels. Neuron. 18:675–682. doi:10.1016/S0896-6273(00)80307-4
    OpenUrlCrossRefPubMed
  59. ↵
    1. Taglialatela, M.,
    2. L. Toro, and
    3. E. Stefani
    . 1992. Novel voltage clamp to record small, fast currents from ion channels expressed in Xenopus oocytes. Biophys. J. 61:78–82. doi:10.1016/S0006-3495(92)81817-9
    OpenUrlCrossRefPubMed
  60. ↵
    1. Tarentino, A.L., and
    2. T.H. Plummer Jr
    . 1994. Enzymatic deglycosylation of asparagine-linked glycans: Purification, properties, and specificity of oligosaccharide-cleaving enzymes from Flavobacterium meningosepticum. Methods Enzymol. 230:44–57. doi:10.1016/0076-6879(94)30006-2
    OpenUrlCrossRefPubMed
  61. ↵
    1. Thornhill, W.B.,
    2. M.B. Wu,
    3. X. Jiang,
    4. X. Wu,
    5. P.T. Morgan, and
    6. J.F. Margiotta
    . 1996. Expression of Kv1.1 delayed rectifier potassium channels in Lec mutant Chinese hamster ovary cell lines reveals a role for sialidation in channel function. J. Biol. Chem. 271:19093–19098. doi:10.1074/jbc.271.32.19093
    OpenUrlAbstract/FREE Full Text
  62. ↵
    1. Ufret-Vincenty, C.A.,
    2. D.J. Baro,
    3. W.J. Lederer,
    4. H.A. Rockman,
    5. L.E. Quinones, and
    6. L.F. Santana
    . 2001a. Role of sodium channel deglycosylation in the genesis of cardiac arrhythmias in heart failure. J. Biol. Chem. 276:28197–28203. doi:10.1074/jbc.M102548200
    OpenUrlAbstract/FREE Full Text
  63. ↵
    1. Ufret-Vincenty, C.A.,
    2. D.J. Baro, and
    3. L.F. Santana
    . 2001b. Differential contribution of sialic acid to the function of repolarizing K(+) currents in ventricular myocytes. Am. J. Physiol. Cell Physiol. 281:C464–C474. doi:10.1152/ajpcell.2001.281.2.C464
    OpenUrlCrossRefPubMed
  64. ↵
    1. Varki, A.
    1993. Biological roles of oligosaccharides: All of the theories are correct. Glycobiology. 3:97–130. doi:10.1093/glycob/3.2.97
    OpenUrlCrossRefPubMed
  65. ↵
    1. Watanabe, I.,
    2. H.G. Wang,
    3. J.J. Sutachan,
    4. J. Zhu,
    5. E. Recio-Pinto, and
    6. W.B. Thornhill
    . 2003. Glycosylation affects rat Kv1.1 potassium channel gating by a combined surface potential and cooperative subunit interaction mechanism. J. Physiol. 550:51–66. doi:10.1113/jphysiol.2003.040337
    OpenUrlCrossRefPubMed
  66. ↵
    1. Watanabe, I.,
    2. J. Zhu,
    3. E. Recio-Pinto, and
    4. W.B. Thornhill
    . 2004. Glycosylation affects the protein stability and cell surface expression of Kv1.4 but Not Kv1.1 potassium channels. A pore region determinant dictates the effect of glycosylation on trafficking. J. Biol. Chem. 279:8879–8885. doi:10.1074/jbc.M309802200
    OpenUrlAbstract/FREE Full Text
  67. ↵
    1. Watanabe, I.,
    2. J. Zhu,
    3. J.J. Sutachan,
    4. A. Gottschalk,
    5. E. Recio-Pinto, and
    6. W.B. Thornhill
    . 2007. The glycosylation state of Kv1.2 potassium channels affects trafficking, gating, and simulated action potentials. Brain Res. 1144:1–18. doi:10.1016/j.brainres.2007.01.092
    OpenUrlCrossRefPubMed
  68. ↵
    1. Watanabe, I.,
    2. J. Zhu,
    3. E. Recio-Pinto, and
    4. W.B. Thornhill
    . 2015. The degree of N-glycosylation affects the trafficking and cell surface expression levels of Kv1.4 potassium channels. J. Membr. Biol. 248:187–196. doi:10.1007/s00232-014-9756-7
    OpenUrlCrossRef
  69. ↵
    1. Wirkner, K.,
    2. H. Hognestad,
    3. R. Jahnel,
    4. F. Hucho, and
    5. P. Illes
    . 2005. Characterization of rat transient receptor potential vanilloid 1 receptors lacking the N-glycosylation site N604. Neuroreport. 16:997–1001. doi:10.1097/00001756-200506210-00023
    OpenUrlCrossRefPubMed
  70. ↵
    1. Yifrach, O., and
    2. R. MacKinnon
    . 2002. Energetics of pore opening in a voltage-gated K(+) channel. Cell. 111:231–239. doi:10.1016/S0092-8674(02)01013-9
    OpenUrlCrossRefPubMed
  71. ↵
    1. Zhang, Y.,
    2. H.A. Hartmann, and
    3. J. Satin
    . 1999. Glycosylation influences voltage-dependent gating of cardiac and skeletal muscle sodium channels. J. Membr. Biol. 171:195–207. doi:10.1007/s002329900571
    OpenUrlCrossRefPubMed
  72. ↵
    1. Zhu, J.,
    2. I. Watanabe,
    3. B. Gomez, and
    4. W.B. Thornhill
    . 2001. Determinants involved in Kv1 potassium channel folding in the endoplasmic reticulum, glycosylation in the Golgi, and cell surface expression. J. Biol. Chem. 276:39419–39427. doi:10.1074/jbc.M107399200
    OpenUrlAbstract/FREE Full Text
  73. ↵
    1. Zhu, J.,
    2. I. Watanabe,
    3. A. Poholek,
    4. M. Koss,
    5. B. Gomez,
    6. C. Yan,
    7. E. Recio-Pinto, and
    8. W.B. Thornhill
    . 2003. Allowed N-glycosylation sites on the Kv1.2 potassium channel S1-S2 linker: Implications for linker secondary structure and the glycosylation effect on channel function. Biochem. J. 375:769–775. doi:10.1042/bj20030517
    OpenUrlAbstract/FREE Full Text
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This is a work of the U.S. Government and is not subject to copyright protection in the United States. Foreign copyrights may apply. http://www.rupress.org/terms/https://creativecommons.org/licenses/by-nc-sa/4.0/This article is distributed under the terms of an Attribution–Noncommercial–Share Alike–No Mirror Sites license for the first six months after the publication date (see http://www.rupress.org/terms/). After six months it is available under a Creative Commons License (Attribution–Noncommercial–Share Alike 4.0 International license, as described at https://creativecommons.org/licenses/by-nc-sa/4.0/).
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The Journal of General Physiology: 151 (10)

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October 7, 2019
Volume 151, No. 10

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