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. 2016 Aug 19;5:e16093. doi: 10.7554/eLife.16093

Histidine phosphorylation relieves copper inhibition in the mammalian potassium channel KCa3.1

Shekhar Srivastava 1,2,3, Saswati Panda 1,2,3, Zhai Li 1,2,3, Stephen R Fuhs 4, Tony Hunter 4, Dennis J Thiele 5,6, Stevan R Hubbard 1,3,*, Edward Y Skolnik 1,2,3,*
Editor: Roger J Davis7
PMCID: PMC5005030  PMID: 27542194

Abstract

KCa2.1, KCa2.2, KCa2.3 and KCa3.1 constitute a family of mammalian small- to intermediate-conductance potassium channels that are activated by calcium-calmodulin. KCa3.1 is unique among these four channels in that activation requires, in addition to calcium, phosphorylation of a single histidine residue (His358) in the cytoplasmic region, by nucleoside diphosphate kinase-B (NDPK-B). The mechanism by which KCa3.1 is activated by histidine phosphorylation is unknown. Histidine phosphorylation is well characterized in prokaryotes but poorly understood in eukaryotes. Here, we demonstrate that phosphorylation of His358 activates KCa3.1 by antagonizing copper-mediated inhibition of the channel. Furthermore, we show that activated CD4+ T cells deficient in intracellular copper exhibit increased KCa3.1 histidine phosphorylation and channel activity, leading to increased calcium flux and cytokine production. These findings reveal a novel regulatory mechanism for a mammalian potassium channel and for T-cell activation, and highlight a unique feature of histidine versus serine/threonine and tyrosine as a regulatory phosphorylation site.

DOI: http://dx.doi.org/10.7554/eLife.16093.001

Research Organism: None

Introduction

KCa2.1, KCa2.2, KCa2.3 and KCa3.1 (also called SK1–4) are encoded by the KCNN genes and respond to calcium via calmodulin, which is constitutively bound to the cytoplasmic region of these channels (Adelman et al., 2012). KCa2.1, KCa2.2 and KCa2.3 are expressed predominantly in neurons, contributing to medium afterhyperpolarization, whereas KCa3.1 plays a key role in the activation of T cells, B cells and mast cells (Feske et al., 2015). Potassium efflux via KCa3.1 is required to maintain a negative membrane potential, which provides the electrical gradient for sustained calcium influx via calcium release-activated channels (CRACs) and subsequent production of cytokines (Feske et al., 2015).

A unique feature of KCa3.1 relative to the other KCa channels is its regulation by histidine phosphorylation. We showed previously that His358 of KCa3.1 is phosphorylated (pHis358) by nucleoside diphosphate kinase-B (NDPK-B) (Di et al., 2010; Srivastava et al., 2006b), which, along with NDPK-A, are the only two mammalian protein histidine kinases identified to date (Attwood and Wieland, 2015). We also showed that KCa3.1 activation requires phosphatidylinositol 3-phosphate (PI(3)P) (Srivastava et al., 2006a), generated by a class II phosphatidylinositol 3-kinase (PI3K-C2β) (Srivastava et al., 2009), and that KCa3.1 is negatively regulated by protein histidine phosphatase-1 (PHPT1), which dephosphorylates pHis358 (Srivastava et al., 2008), and by myotubularin-related protein-6 (MTMR6), which dephosphorylates PI(3)P (Srivastava et al., 2005). In addition, we recently identified phosphoglycerate mutase-5 (PGAM5) as a histidine phosphatase that specifically dephosphorylates the catalytic histidine (His118) in NDPK-B. By dephosphorylating NDPK-B, PGAM5 negatively regulates T-cell receptor signaling by inhibiting NDPK-B-mediated histidine phosphorylation and activation of KCa3.1 (Panda et al., 2016).

We reported previously that mutation of His358 (H358N) converted KCa3.1 into a channel that, like the other three KCa channels, requires only calcium-calmodulin for activation (Srivastava et al., 2006b). Furthermore, swapping 14 residues of KCa3.1 containing His358 with the equivalent residues of KCa2.3 converted the latter into a channel that required NDPK-B and PI(3)P for activation (Srivastava et al., 2006a). These studies highlighted the autonomous role of His358 and proximal residues in the regulation of KCa3.1.

Although histidine phosphorylation is well characterized in prokaryotic two-component systems used in chemotaxis and other sensing systems (Hess et al., 1988), it is poorly characterized in eukaryotes (Klumpp and Krieglstein, 2009), in part because phosphohistidine is more labile than phosphotyrosine or phosphoserine/threonine. In addition to KCa3.1, histidine phosphorylation of several mammalian proteins by NDPKs has been reported, including the β subunit of heterotrimeric G proteins and the transient receptor potential vanilloid-5 (TRPV5) channel (Attwood and Wieland, 2015; Cai et al., 2014; Klumpp and Krieglstein, 2009). However, the functional consequences of histidine phosphorylation of these eukaryotic proteins, and the mechanisms whereby histidine phosphorylation regulates their activity, are poorly understood. The regulation of KCa3.1 by histidine phosphorylation has emerged as the clearest example in a mammalian protein of the functional importance of this post-translational event, yet the molecular basis for His358-mediated regulation of KCa3.1 is unknown.

The special role of histidine in KCa3.1 inhibition, together with the knowledge that histidine is a common ligand in metal-ion coordination, led us to hypothesize that the four copies of His358 in the cytoplasmic domains of the homotetrameric channel coordinate a metal ion, which renders KCa3.1 refractory to the conformational changes induced by calcium binding to calmodulin. Here, we provide evidence for copper-mediated inhibition of KCa3.1 from patch-clamping studies of KCa3.1 in human embryonic kidney (HEK) 293 cells and in mouse embryonic fibroblasts (MEFs) from copper transporter-1 (Ctr1) knockout mice. Moreover, we show that copper inhibition of KCa3.1 is relevant in a physiologic context, namely, regulation of CD4+ T-cell activation.

Results

KCa3.1 is activated by metal chelators and inhibited by copper in whole-cell membrane patches

To test the hypothesis that KCa3.1 is inhibited by His358-mediated metal binding, we first used whole-cell patch clamping to measure the effect of the cell-permeable metal chelator TPEN (N,N,N′,N′-tetrakis(2-pyridylmethyl) ethylenediamine) on the current from HEK 293 cells stably transfected with GFP-KCa3.1 (293-KCa3.1 cells). TPEN is a high-affinity chelator (KD < 10–10) of several metal ions, including zinc, copper, iron, nickel and manganese (Smith and Martell, 1975; Treves et al., 1994). As shown in Figure 1A,B,E, the addition of 20 µM TPEN to the cell bath markedly increased the measured current from these cells, consistent with metal-ion inhibition of KCa3.1. We identified the KCa3.1-mediated current as a major contributor to the current from these cells since it was largely blocked by 1 µM TRAM-34, a KCa3.1-specific inhibitor (Wulff et al., 2000).

Figure 1. Metal chelators activate and copper inhibits KCa3.1 in whole-cell membrane patches.

Figure 1.

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(A–D) (i) Representative current vs. voltage (IV) plot recorded from a 293-KCa3.1(WT) cell (A–C) or a 293-KCa3.1(H358N) cell (D). After obtaining the basal current, TPEN (20 µM) (A, B) or TTM (20 µM) (C, D) was perfused in the bath solution, followed by the addition of either ZnCl2 (100 µM) (A) or CuCl2 (100 µM) (B–D), and then TRAM-34 (1 µM). (A–D) (ii) Representative trace of current (pA/pF) recorded at +60 mV using ramp protocol applied every 10 s (sweep frequency). The timing of additions (TPEN, CuCl2, etc.) is indicated by the arrows. (E, F) Summary bar graph of TRAM-34-sensitive current at +60 mV for data measured from 293-KCa3.1(WT) cells (E) or from 293-KCa3.1(H358N) cells (F). In (E), +ZnCl2 is to be compared with +TPEN (i.e. ZnCl2 added after treatment with TPEN), and +CuCl2 with +TTM (i.e. CuCl2 added after treatment with TTM). Data are displayed as mean ± the standard error of the mean (SEM) (n = 10 (E) or 8 (F) cells). *p≤0.01 versus Basal and for +CuCl2 versus +TTM.

DOI: http://dx.doi.org/10.7554/eLife.16093.002

Histidine often serves as a ligand for zinc and copper (Dokmanić et al., 2008). We therefore tested whether addition of these ions could reverse the TPEN-mediated activation of KCa3.1. Addition of 100 µM zinc (ZnCl2) did not alter substantially the measured current from 293-KCa3.1 cells (Figure 1A,E), but addition of 100 µM copper (CuCl2) to the bath caused a rapid decrease in the current (Figure 1B,E), consistent with channel inhibition.

Because copper but not zinc inhibited KCa3.1 when added to cells, we also tested the copper-selective chelator tetrathiomolybdate (TTM) (Brewer et al., 2006). Similar to TPEN, addition of 20 µM TTM to 293-KCa3.1 cells caused a substantial increase in the measured current, which was reversed by the addition of 100 µM CuCl2 (Figure 1C,E).

To determine whether the observed copper inhibition of KCa3.1 was mediated by His358, we transfected HEK 293 cells with the KCa3.1 mutant H358N and tested the effect of TTM and copper on channel activity. In contrast to wild-type (WT) KCa3.1, TPEN did not activate substantially, nor did copper inhibit substantially, KCa3.1 H358N (Figure 1D,F), demonstrating that copper inhibition of KCa3.1 is strictly dependent on His358.

KCa3.1 is activated by TTM and inhibited by copper in isolated inside-out membrane patches, dependent on the phosphorylation state of His358

The direct effects of TTM and copper on KCa3.1 channel activity were further explored in inside-out (I/O) patch-clamp experiments using membrane patches pulled from 293-KCa3.1 cells. These experiments were designed to explore the model that KCa3.1 is inhibited by copper binding to His358, which is released by GTP/ATP-dependent NDPK-B phosphorylation of His358 or, in vitro, by chelation by TTM, with either inhibitory-release mechanism requiring calcium binding to calmodulin for channel activation. We first determined whether the addition of 20 µM TTM to the bath could activate KCa3.1, which it did (Figure 2A). We then added 10 µM copper (CuCl2) to the bath, which decreased the current to the basal level. Re-application of TTM activated the channel (Figure 2A), demonstrating that inhibition and activation by copper and TTM, respectively, are reversible. NDPK-B also activated copper-inhibited KCa3.1, to a similar extent as TTM (Figure 2B). However, while NDPK-B required GTP to be present to activate KCa3.1 (Figure 2C), as GTP (or ATP) is necessary for NDPK-B to phosphorylate His358, TTM did not require GTP to activate KCa3.1 (Figure 2A,B), consistent with TTM simply chelating copper from His358.

Figure 2. TTM activates and copper inhibits KCa3.1 in inside-out membrane patches, dependent on His358 phosphorylation.

Figure 2.

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Representative recordings of channel activity versus time from I/O patches isolated from 293-KCa3.1 (A–F) or parental 293 (G) cells. All recordings were done in 300 nM free Ca2+ and 500 µM GTP unless otherwise specified. TTM (20 µM), CuCl2 (10 µM), NDPK-B (10 µg/ml) and TRAM-34 (1 µM) were added as indicated. Shown are representative patch data from three independent experiments. (A–C) were performed in the presence of EGTA (5 mM) and (D–F) in the absence of EGTA in the bath solution. (E,F) were done in the presence of 10 µM free calcium in the bath solution.

DOI: http://dx.doi.org/10.7554/eLife.16093.003

Addition of 1 µM TRAM-34 suppressed channel activity to the basal level in all I/O patch experiments (Figure 2A–D,F), confirming that the measured activity is due to KCa3.1. The TTM-mediated current increase in the I/O patches was further confirmed to be KCa3.1-specific by demonstrating that addition of TTM after treatment with TRAM-34 did not increase the current (Figure 2A), as well as by the inability of either TTM or NDPK-B to elicit an increase in current in I/O patches isolated from parental (non-KCa3.1-transfected) 293 cells (Figure 2G).

To provide further evidence that phosphorylation by NDPK-B is the key event in regulating copper sensitivity of KCa3.1, we assessed the effects of TTM and copper on KCa3.1 activity after phosphorylation by NDPK-B. We found that copper did not inhibit KCa3.1 channel activity following prior activation by NDPK-B (Figure 2C), which would be predicted if phosphorylation of the imidazole ring of His358 abrogates copper binding. This is in contrast to the ability of copper to inhibit a TTM-activated channel (Figure 2A,B), in which case copper is free to bind to His358 once TTM is removed. Moreover, the addition of TTM did not lead to further activation of KCa3.1 (Figure 2C), as there should be little or no remaining His358-bound copper for TTM to chelate.

The experiments in Figure 2A–C were performed in the presence of EGTA to buffer calcium in the bath solution. In addition to chelating calcium, EGTA also chelates copper (Xiao and Wedd, 2010). While 10 µM CuCl2 was required to inhibit KCa3.1 in an I/O patch in the presence of EGTA, 100 nM CuCl2 was sufficient to inhibit in the absence of EGTA (Figure 2D). Furthermore, the ability of copper to inhibit KCa3.1 is inversely dependent on the calcium concentration: approximately 1 µM CuCl2 was required for inhibition at 10 µM CaCl2 (Figure 2E), versus 100 nM CuCl2 at 300 nM CaCl2 (Figure 2D). The copper (Figure 2E) and TRAM-34-inhibited currents (Figure 2F) in 10 µM calcium, following the addition of TTM, were the same, indicating that the copper-inhibited current is specific to KCa3.1.

We showed previously that activation of KCa3.1 requires both His358 phosphorylation by NDPK-B and calcium binding to KCa3.1-bound calmodulin (Srivastava et al., 2006b). We therefore tested whether calcium is also required for activation by TTM. Addition of 20 µM TTM to an I/O patch in low calcium (30 nM) did not result in KCa3.1 activation (Figure 3A), indicating that calcium is also required for TTM activation of KCa3.1.

Figure 3. Calcium is required for activation of KCa3.1 by NDPK-B or TTM in inside-out membrane patches.

Figure 3.

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Representative recordings of channel activity versus time from I/O patches isolated from 293-KCa3.1 cells as described in Figure 2. All recordings were done in 500 µM GTP and either 30 or 300 nM free Ca2+ as indicated. TTM (20 µM), NDPK-B (10 µg/ml), CuCl2 (10 µM) and TRAM-34 (1 µM) were added as indicated. Shown are representative patch data from three independent experiments.

DOI: http://dx.doi.org/10.7554/eLife.16093.004

If copper bound to His358 is chelated by TTM in low calcium, we anticipated that perfusing the patch with high calcium (300 nM) following TTM would be sufficient to activate. Surprisingly, addition of 300 nM calcium to an I/O patch following 20 µM TTM did not activate KCa3.1, although it did activate other non-KCa3.1 channels that were not inhibited by TRAM-34 (Figure 3A). Rather, TTM and high calcium needed to be present simultaneously for activation (Figure 3A). A similar result was obtained for activation of KCa3.1 by NDPK-B (Figure 3B). These data suggest that a calcium/calmodulin-induced conformational change in the channel is required for TTM and NDPK-B to gain access to copper and His358, respectively. Addition of copper inhibited the TRAM-34-sensitive current, but not the current from other channels activated by calcium (Figure 3A), providing further evidence that copper specifically inhibits KCa3.1.

Basal KCa3.1 current is elevated in transfected MEFs from Ctr1-/- mice

To further explore our finding that copper inhibits KCa3.1 activity, we measured current from GFP-KCa3.1-transfected MEFs derived from mice in which the gene for the copper transporter Ctr1 was deleted (Lee et al., 2002). As shown in Figure 4A–C, the basal current is approximately three-fold higher in the KCa3.1-transfected Ctr1-/- MEFs than in the transfected WT MEFs expressing similar amounts of GFP-KCa3.1 (only GFP+ cells exhibiting similar levels of immunofluorescence were patched). Furthermore, in contrast to the transfected WT MEFs (Figure 4A,C), addition of 20 µM TTM had little effect on the current from the transfected Ctr1-/- MEFs (Figure 4B,C). Addition of exogenous copper (10 µM) inhibited KCa3.1 current in the transfected Ctr1-/- MEFs as well as the WT MEFs (Figure 4A–C). Entry of copper into Ctr1-/- MEFs was presumably mediated by a Ctr1-independent process, as has been described (Lee et al., 2002). The failure of exogenous TTM to increase KCa3.1 current in WT MEFs to levels similar to those in Ctr1-/- MEFs (Figure 4A,B) is likely due to incomplete chelation of copper bound to His358.

Figure 4. Basal KCa3.1 current is elevated in transfected MEFs from Ctr1-/- mice.

Figure 4.

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(A, B) Representative current vs. voltage (IV) plot recorded from a KCa3.1-transfected MEF from a wild-type (WT) mouse (A) or a Ctr1-/-(B) mouse. After obtaining the basal current, TTM (20 µM) was perfused in the bath solution followed by CuCl2 (10 µM) and then TRAM-34 (1 µM). (C) Summary bar graph of the TRAM-34-sensitive current at +60 mV for data measured from MEFs from WT or Ctr1-/- mice. For statistical analysis, one-way-ANOVA was used, and the Bonferroni test was applied to compare the mean values. Data are displayed as mean ± SEM (n = 10 cells). *p≤0.01 versus Basal in WT MEFs and for +CuCl2 versus +TTM. (D) Exofacial HA-tagged KCa3.1 was transfected into WT or Ctr1-/- MEFs, and cell surface expression was assessed by FACs analysis following staining with anti-HA antibodies and with anti-mouse FITC antibodies in non-permeabilized cells. (i) Flow cytometry results of WT and Ctr1-/- controls stained with only the secondary anti-mouse FITC antibody, or of WT + KCa3.1-HA and Ctr1-/- + KCa3.1-HA MEFs stained with both anti-HA and anti-mouse FITC antibodies. (ii) Mean fluorescence intensity (MFI) of WT + KCa3.1-HA and Ctr1-/- + KCa3.1-HA MEFs.

DOI: http://dx.doi.org/10.7554/eLife.16093.005

To rule out the possibility that the increased KCa3.1 current in the Ctr1-/- MEFs was due to an increase in surface expression of KCa3.1 in these MEFs, we transfected WT and Ctr1-/- MEFs with KCa3.1 bearing an exofacial hemagglutinin (HA) epitope tag inserted into the extracellular loop between transmembrane helices S3 and S4 (Srivastava et al., 2005). Previous studies (Srivastava et al., 2005; Syme et al., 2003) have demonstrated that KCa3.1 with this tag functions normally and can be specifically detected at the plasma membrane following staining of non-permeabilized cells with anti-HA antibody. FACS analysis of non-permeabilized cells demonstrated that the surface levels of KCa3.1 in the two types of MEFs were very similar (Figure 4D).

CD4+ T cells derived from Ctr1+/- mice are hyperactivated and endogenous KCa3.1 is hyperphosphorylated on histidine and exhibits increased activity

Finally, we tested whether copper inhibition of KCa3.1 is physiologically relevant in the context of CD4+ T-cell activation. By effluxing potassium ions, KCa3.1 maintains a negative membrane potential required for additional calcium influx. Because Ctr1-/- mice are embryonically lethal, we assessed T-cell activation in CD4+ T cells from mice that were heterozygous for Ctr1. The level of cytoplasmic copper in the Ctr1+/- splenocytes is approximately 50% of that in WT splenocytes (Lee et al., 2001). As shown in Figure 5A, endogenous KCa3.1 channel activity was markedly increased in CD4+ T cells isolated from Ctr1+/- mice compared to those from WT littermate controls. Consistent with increased KCa3.1 activity, by blotting with a monoclonal antibody specific for 3-pHis (Fuhs et al., 2015), we observed significantly increased histidine phosphorylation of (endogenous) KCa3.1 in T cells from Ctr1+/- mice versus those from WT littermate controls, with no appreciable difference in KCa3.1 expression levels (Figure 5B). In accord with elevated KCa3.1 activity, T-cell receptor-stimulated (by anti-CD3/CD28 antibodies) calcium influx and cytokine production—interleukin-2 (IL-2), interferon-γ (IFN-γ) and tumor necrosis factor-α (TNFα)—were also substantially increased in Ctr1+/- T cells (Figure 5C–E).

Figure 5. CD4+ T cells from Ctr1+/- mice are hyperactivated.

Figure 5.

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Naive CD4+ T cells were isolated from spleens of wild-type (WT) or Ctr1+/- mice and activated with anti-CD3 and -CD28 antibodies for 48 hr. (A) KCa3.1 channel activity was determined by whole-cell patch clamping as in Figure 1. Shown is a summary bar graph of TRAM-34 (1 µM; KCa3.1) and ShK (1 nM; Kv1.3)-sensitive currents at +60 mV (n = 12 (WT) or 15 (Ctr1+/-)). For statistical analysis, one-way-ANOVA was used, and the Bonferroni test was applied to compare the mean values. Data are displayed as mean ± SEM (n = 10 cells). *p≤0.01 vs TRAM-34-sensitive current in WT. (B) KCa3.1 was immunoprecipitated from lysates of CD4+ T cells from WT or Ctr1+/- mice and probed with a monoclonal antibody to 3-pHis (clone SC56-2) or KCa3.1 as indicated. (C) Activated cells were rested overnight, loaded with Fura-2, AM and attached to a poly-L-lysine-coated coverslip for 20 min. Calcium imaging was then performed in unstimulated cells and following stimulation with anti-CD3 and -CD28 antibodies (n = 80–100 cells). (D) ELISA to quantify IL-2 and IFN-γ in the supernatants. Statistical significance was calculated using Student’s t-test (*p<0.05, **p<0.01, n.s.: p>0.05; not significant as compared to WT). (E) Representative intracellular flow cytometry detecting cytokine expression following restimulation with anti-CD3 and -CD28 antibodies in the presence of brefeldin-A for 4 hr. Cells not re-stimulated with anti-CD3 and -CD28 were stained and analyzed in a similar manner and served as controls for setting the cut-off limits for cytokine production. Data are representative of two independent experiments.

DOI: http://dx.doi.org/10.7554/eLife.16093.006

Discussion

KCa3.1 is unique among the four calcium-activated KCa potassium channels in that activation requires phosphorylation by NDPK-B of a specific histidine residue, His358, in the cytoplasmic domain (Srivastava et al., 2006b). The mechanism by which (non-phosphorylated) His358 is inhibitory to KCa3.1 was not known. The specificity of His358 inhibition—the conservative substitution asparagine completely disinhibits the channel—pointed to an inhibitory mechanism for which histidine is uniquely suited.

One important biochemical feature of histidine is its role in coordinating metal ions, through one of the two nitrogen atoms (N1 and N3) in the imidazole ring. Metal ions for which histidine serves as a ligand include copper, nickel, iron, zinc and manganese (Zheng et al., 2008). In homotetrameric KCa3.1, four copies of His358 are present in the cytoplasmic region, related (on average) by C4 symmetry. Of the metal ions listed above, only copper (Cu(II)) is found to be coordinated solely by four histidine residues (Dokmanić et al., 2008).

These considerations suggested the possibility that the KCa3.1 inhibitory mechanism, mediated by His358, involves copper binding. In the present study, we provide strong support for this hypothesis. Moreover, we show that copper inhibition of KCa3.1 is physiologically relevant for CD4+ T-cell activation; KCa3.1 in T cells deficient in intracellular copper (from Ctr1+/- mice) exhibits higher activity, which results in increased calcium influx and cytokine production (Figure 5). Although the BK and Shaker potassium channels and TASK-3, a two-pore-domain potassium channel, have also been shown to be inhibited by copper (Gruss et al., 2004; Ma et al., 2008), the inhibition mechanism is distinct from that in KCa3.1 because copper acts extracellularly in these other channels, and there is no evidence that these effects are regulated by histidine phosphorylation.

How would His358 be positioned spatially to bind a copper ion? His358 is located just C-terminal to the calmodulin-binding domain in KCa3.1 and 18 residues N-terminal to the predicted C-terminal coiled-coil region, which crystallographic analysis shows forms a four-helix bundle (S.R.H., unpublished data). Coiled-coil prediction software indicates that there is a reasonable probability that the region encompassing His358 also forms a coiled-coil/four-helix bundle, with His358 occupying an inward-facing ‘a’ position in the heptad repeat, which would position the four copies of His358 to coordinate a copper ion (via N3 in the imidazole ring) on the four-fold axis. Biophysical studies are ongoing to validate this structural model and demonstrate direct binding of copper to His358.

Our model for the phosphorylation- and calcium-dependent activation of KCa3.1 is presented in Figure 6. In the absence of calcium (basal state), copper binding to the four copies of His358 in homotetrameric KCa3.1 maintains low channel activity by opposing the calcium/calmodulin-mediated conformational changes that induce channel opening. Upon a marked increase in intracellular calcium levels due to T-cell stimulation (for example), calcium binding to channel-associated calmodulin induces conformational changes in the channel, distorting the four-helix bundle locally (the C-terminal four-helix bundle is maintained) and partially destabilizing copper binding to His358, thus exposing it to phosphorylation by NDPK-B. Because stable copper binding requires tetravalent coordination, phosphorylation of a single His358 residue (on N3) in the tetramer is probably sufficient to abrogate copper binding and inhibition of KCa3.1. Unencumbered by copper, the channel-opening conformational changes induced by calcium proceed in the same manner in KCa3.1 as in the other three KCa channels. Consistent with our model, (i) NDPK-B or TTM activation of KCa3.1 required calcium to be present concurrently (Figure 3), (ii) copper did not inhibit NDPK-B-activated (phosphorylated) KCa3.1 (Figure 2C), and (iii) KCa3.1 was hyperphosphorylated on histidine (and more active) in CD4+ T cells from Ctr1+/- mice (Figure 5B), which contain lower intracellular copper levels.

Figure 6. Model for KCa3.1 activation by calcium and histidine phosphorylation.

Figure 6.

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For clarity, only two of the four KCa3.1 subunits and calmodulin (CaM) are shown. In the basal state (no calcium; left panel), the CaM C lobe (green sphere) is bound to the N-terminal segment of the calmodulin-binding domain (CBD, yellow) in KCa3.1 (Schumacher et al., 2004). The transmembrane S6 helices close off the channel on the cytoplasmic side. At the C terminus of KCa3.1 is a coiled-coil region that forms a four-helix bundle (violet cylinders; S.R.H., unpublished data). Based on coiled-coil prediction software, it is probable that the region containing His358, which is just C-terminal to the CBD, also forms a four-helix bundle (maroon cylinders), with His358 (pentagon) occupying an inward-facing ‘a’ position in the heptad repeat. This would position the four copies of His358 for coordination of a Cu(II) ion on the axis of the four-helix bundle, which would stabilize the four-helix bundle and act to resist the conformational changes induced by calcium binding to the CaM N lobe (blue sphere). An increase in intracellular calcium induces conformational changes in the CBD that partially destabilize copper binding (middle panel), providing access to His358 for phosphorylation by NDPK-B (or copper chelation by TTM). Upon phosphorylation of His358, copper binding is abrogated, and the calcium-induced conformational changes in the CBD lead to channel opening (right panel; exact mechanism not known) (Adelman et al., 2012; Sachyani et al., 2014).

DOI: http://dx.doi.org/10.7554/eLife.16093.007

The metal-binding properties of histidine and its ability to be phosphorylated distinguish histidine from the common phosphorylatable residues in eukaryotes, serine/threonine and tyrosine. Although our model for KCa3.1 inhibition and activation might be unique for a potassium channel, we recently demonstrated that the TRPV5 calcium channel is also activated by NDPK-B-mediated histidine phosphorylation (Cai et al., 2014). Whether metal-ion coordination by histidine, disrupted through histidine phosphorylation, is the basis for regulation of TRPV5, or of any of the 1- and 3-pHis-containing proteins recently identified with anti-pHis antibodies (Fuhs et al., 2015), remains to be explored.

Our finding that limiting intracellular copper can enhance activation of T cells has therapeutic implications. It was recently shown that copper chelators such as penicillamine, used to treat Wilson disease (copper-overload disorder), were able to slow growth of oncogenic BRAFV600E-driven tumors in mice, through inactivation of MEK1 (Brady et al., 2014). In addition to directly targeting BRAFV600E in tumors, treatment with checkpoint inhibitors to enhance immune-cell killing has led to dramatic response rates (Pardoll, 2012). Thus, treatment of patients harboring BRAFV600E driven-tumors with copper chelators has the potential to not only block BRAF activation of the MAP kinase pathway, but also to enhance T-cell-mediated killing of these tumors via enhanced activation of KCa3.1.

Materials and methods

Materials

TPEN (Sigma-Aldrich, St. Louis, MO, Cat # P4413), TTM (Sigma-Aldrich, Cat # 323446), CuCl2 (Sigma-Aldrich, Cat # C6641) and ZnCl2 (Acros, Geel, Belgium, Cat # 318170100) were purchased. TRAM-34 was obtained from Heike Wulff (University of California–Davis). Ctr1-/- MEFs and Ctr1+/- splenocytes have been described previously (Lee et al., 2001, 2002).

Whole-cell patch clamping

HEK 293 cells stably transfected with GFP-tagged human KCa3.1 (293-KCa3.1) (in vector pEGFP-C1), either WT or H358N, or MEFs from WT or Ctr1-/- mice transiently transfected with mCherry-tagged KCa3.1 were used for electrophysiological studies. Cells were plated on 15-mm diameter coverslips and, after 24 hr, patch clamping was performed at room temperature as described (Srivastava et al., 2005), using a pipette solution containing 100 mM K-gluconate, 30 mM KCl, 1.2 mM MgCl2, 5 mM EGTA, 4.3 mM CaCl2 and 10 mM HEPES, pH 7.2, with 1 N KOH (calculated free Ca2+, 1 µM), and a bath solution containing 140 mM NaCl, 5 mM KCl, 1 mM CaCl2, 1 mM MgCl2, 10 mM glucose and 10 mM HEPES, pH 7.4. Patch-clamp pipettes had resistances ranging between 3 and 3.5 MΩ. Current-voltage (I-V) relationships were measured using ramp voltage-clamp protocols (at 15-s intervals) from a holding potential of −70 to −120 mV, followed by ramp depolarization to +60 mV (symmetrical ramp rate of 0.18 mV·ms−1). The current-voltage relationship was obtained by plotting the current during the depolarizing ramp phase as a function of the corresponding voltage. Membrane currents were filtered (−3 dB at 1 kHz) and digitized at 10 kHz (pClamp 10.3 with Digidata 1440A ADC interface; Molecular Devices, Sunnyvale, CA). Cell capacitance and pipette series resistances were compensated (usually > 80%). Whole-cell current density was expressed as picoamperes per picofarad (pA/pF).

Inside-out (I/O) patch clamping

293-KCa3.1 cells were used for single-channel recordings, performed using the I/O patch-clamp mode. Data were obtained using standard patch-clamp techniques with an Axopatch-200B amplifier (Molecular Devices). Currents were recorded at room temperature (21–23°C), low-pass filtered (-3-dB cut-off frequency at 1 kHz), and recorded on computer disk at a sampling frequency of 5 kHz (Clampex; Molecular Devices). The pipette resistance was 3–4 MΩ when filled with a solution containing 145 mM KCl, 1 mM MgCl2, 1 mM CaCl2 and 5 mM HEPES, pH 7.4. The bath solution contained 141 mM KCl, 5 mM EGTA, 1 mM MgCl2, 3.3 mM CaCl2, 10 mM glucose and 5 mM HEPES, pH 7.2. The calcium activity was calculated to be 300 nM (Win-MAX Chelator software) (Bers et al., 1994). Data were analyzed using pClamp 10.3 (Molecular Devices) and Origin 7.0 (OriginLab, Northampton, MA) software. GST-NDPK-B was expressed in E. coli (pGEX-4T-2, GE Healthcare, Chicago, IL) and purified by Glutathione-Sepharose chromatography (GE Healthcare).

Characterization of CD4+ T cells isolated from WT and Ctr1+/- mice

Mouse CD4+ T cells were isolated from spleens of WT and Ctr1+/- mice using the mouse CD4+ T-cell isolation kit from Miltenyi Biotec (Bergisch Gladbach, Germany) according to the manufacturer’s protocol. We routinely obtained > 95% CD4+ T cells as assessed by FACS. Whole-cell patch clamping was then performed following stimulation with anti-CD3 (5 µg/ml) and anti-CD28 (2 µg/ml) antibodies for 48 hr, as described (Srivastava et al., 2008). For intracellular cytokine staining, cells were rested overnight and then re-stimulated with anti-CD3 and -CD28 antibodies for 4 hr in the presence of brefeldin-A (1 µg/ml, Sigma-Aldrich). Intracellular staining was performed using the Fixation/Permeabilization kit (BD Biosciences, Franklin Lakes, NJ, Cat # 554714) and antibodies to: IL-2 (Cat # JES6-5H4), IFN-γ (Cat # XMG1.2) or TNFα (Cat # MP6-XT22), according to the manufacturer’s (eBioscience, San Diego, CA) instructions. Cells were analyzed on a LSRII HTS flow cytometer (BD Biosciences), and the results were analyzed with Treestar (FlowJo, Ashland, OR).

Intracellular calcium flux

Mouse CD4+ T cells were loaded with 5 µm Fura-2, AM (Molecular Probes, Eugene, OR) in RPMI 1640 medium and 10% FBS for 30 min at 22–25°C. Cells were then attached to a poly-L-lysine-coated coverslip for 20 min, and calcium imaging was done with an IX81 epifluorescence microscope (Olympus, Tokyo, Japan) and OpenLab imaging software (Improvision, Conventry, UK) as described (Di et al., 2010). For single-cell analysis, 340/380 nm Fura-2 emission ratios of > 100 cells per experiment were analyzed. Background fluorescence obtained from regions containing no cells was digitally subtracted from each image. To compare the different groups, the 340/380 nm ratio was normalized to 1 by dividing the fluorescence values at different time points to the cellular fluorescence at time 0. Experiments were independently repeated at least three times.

Immunoprecipitation and western blotting

Mouse CD4+ T cells were lysed in lysis buffer (100 mM NaCl, 10% glycerol, 1 mM EDTA, 5 mM MgCl2, 0.5% NP-40 and 50 mM Tris, pH 8.0, along with freshly added 1 mM DTT, protease and phosphatase inhibitors) and immunoprecipitated with anti-KCa3.1 antibodies (Alomone Labs, Jerusalem, Israel, Cat # ALM-051). After washing three times with lysis buffer, bound proteins were eluted using SDS loading buffer (pH 8.8), separated by SDS-PAGE (pH 8.8), and immunoblotted with anti-KCa3.1 or rabbit monoclonal anti-3-pHis (clone SC56-2) antibodies (Fuhs et al., 2015).

FACS analysis of MEFs transfected with exofacial HA-tagged KCa3.1

KCa3.1 containing a hemagglutinin (HA) epitope tag inserted into the extracellular loops between transmembrane helices S3 and S4 was transfected into Ctr1-/- and WT MEFs. To assess the plasma membrane levels of KCa3.1, cells were fixed with 4% paraformaldehyde for 10 min on ice and washed three times with PBS. The non-permeabilized fixed cells were incubated with anti-HA antibody (1:200 dilution) for 1 hr on ice, washed three times with PBS, and stained with secondary anti-mouse FITC-conjugated antibody (1:500 dilution) for 30 min on ice. Cells were then washed three times with PBS prior to analysis by flow cytometry. Cells incubated with only the secondary antibody served as the negative control. Flow cytometry was performed using LSR II HTS flow cytometer (BD Biosciences), and the results were analyzed using Treestar (FlowJo).

Acknowledgements

We thank Senena Corbalán-García (University of Murcia) for helpful discussions, Min Ha Hwang (Duke University) for mice dissections, and Heike Wulff (University of California–Davis) for TRAM-34. This work was supported in part by NIH grants R01CA194584 (TH), R01DK074192 (DJT), R21AI107443 (SRH), and R01GM084195 (EYS). TH is a Frank and Else Schilling American Cancer Society Professor and holds the Renato Dulbecco Chair in Cancer Research.

Funding Statement

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Funding Information

This paper was supported by the following grants:

  • National Institute of Allergy and Infectious Diseases R21AI107443 to Stevan R Hubbard.

  • National Institute of Diabetes and Digestive and Kidney Diseases R01DK074192 to Dennis J Thiele.

  • National Institute of General Medical Sciences R01GM084195 to Edward Y Skolnik.

  • National Cancer Institute R01CA194584 to Tony Hunter.

Additional information

Competing interests

TH: Senior editor, eLife.

The other authors declare that no competing interests exist.

Author contributions

SS, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article.

SP, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article.

ZL, Acquisition of data, Contributed unpublished essential data or reagents.

SRF, Drafting or revising the article, Contributed unpublished essential data or reagents.

TH, Drafting or revising the article, Contributed unpublished essential data or reagents.

DJT, Drafting or revising the article, Contributed unpublished essential data or reagents.

SRH, Conception and design, Analysis and interpretation of data, Drafting or revising the article.

EYS, Conception and design, Analysis and interpretation of data, Drafting or revising the article.

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eLife. 2016 Aug 19;5:e16093. doi: 10.7554/eLife.16093.008

Decision letter

Editor: Roger J Davis1

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for submitting your article "Histidine phosphorylation relieves copper inhibition in the mammalian potassium channel KCa3.1" for consideration by eLife. Your article has been reviewed by three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Ivan Dikic as the Senior Editor.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

This study that establishes a potential mechanism for the regulation of KCa3.1 channels in CD4 T cells by histidine phosphorylation on His358. The authors propose that Cu++ ions would coordinate with four His358 residues thus rendering the channel refractory to the conformational changes initiated by the binding of Ca2+ to CaM. This mechanism is blocked by His358 phosphorylation. This may have important physiological significance in the context of understanding CD4 T cell activation.

Essential revisions:

1) Copper binding to His358 is inferred from indirect studies, but is not directly demonstrated – this should be acknowledged. The dose response of copper (Figure 1) shows that 1µM had no effect, 10µM shows an inhibitory effect, and that 100µM was saturating. How does this dose response compare with physiological intracellular copper concentrations?

2) The concentration of free Cu++ in the solutions used for the whole cell and inside-out patch clamp experiments is a concern. The expected Cu++ concentration in the pipette solution used in whole cell experiments and in the bath solution for the inside-out experiments should be specified. EDTA has a binding affinity of 10-18M for Cu++ and it is likely to be the same for EGTA. Under such conditions, both solutions should be Cu++ free and yet channel activity appears to be low in the whole cell and inside-out recordings presented in Figures 1 and 2 in control conditions.

3) Figure 2A shows that channel activity remains high after TTM removal in 300nM Ca2+ (the effect of TTM is not reversible) consistent with the bathing solution being Cu++ free and TTM being capable of chelating Cu++ from the Cu++- H358 complex. However, channel activity appears low in 300nm Ca2+ after TTM treatment in 30nM Ca2+. One would have expected channel activity to be high in this case if TTM had succeeded to chelate Cu++ from the Cu++-H358 complex in 30nM Ca2+ conditions. The authors need to discuss how the action of TTM might be modulated by Ca2+ and may involve more than chelating Cu++.

4) Segments of the inside-out recording in Figure 2A, B and D should also be presented at a higher time resolution to compare single channel activity at 300nm Ca2+ + low Cu++, 10μM CuCl2 and TTM conditions. The authors also need to comment on the finding that in Figure 2A, B and D, channel activity appears greater in CuCl2 conditions than in control before TTM application in 300nm Ca2+ conditions.

5) The proposed mechanism implies that Cu++ coordination by four H358 is state dependent, with Cu++ binding taking place in the channel-closed configuration. Accordingly, the potency of CuCl2 to decrease channel activity following TTM treatment should be Ca2+ dependent with CuCl2 being less potent in saturating Ca2+ conditions (10μM Ca2+). The Ca2+ dependency of CuCl2 mediated inhibition after TTM treatment should be presented in inside-out experiments to confirm state dependency.

6) The significance of the results illustrated in Figure 3A–B would be clearer if information on the mechanisms underlying Cu++ cellular homeostasis, and on the expected basal Cu++ intracellular level is provided. If channel expression is not affected in Ctr1-/- cells, why is there a two fold difference in current intensity following TTM treatment between WT and Ctr1-/- cells assuming that TTM can chelate all the Cu++ in the cells?

7) The structural constraints related to Cu++ coordinating four His358 residues should be discussed. What are the minimal distances between adjacent imidazole nitrogen atoms to insure Cu++ coordination? How does the proposed structural model for the channel closed state impact on the possibility of a dimer-dimer structure for the channel open state? The model presented in Figure 5 presumes that this dimer-dimer structure does not prevail in the channel open state configuration.

8) The increase in KCa3.1 channel activity in Ctr1-/+ CD4 T cells was predicted by the patch clamping studies. However, the increase in His358 phosphorylation was not predicted. This finding raises questions concerning mechanism. Is His358 phosphorylation a passive response to lack of copper binding to His358? This response differs from the major conclusion of this study – that phosphorylation prevents copper interactions with KCa3.1. The observation suggests a more complicated relationship between copper binding and histidine phosphorylation than the authors' conclusions.

9) Very limited information is provided concerning statistical analysis. For example, bar graphs and error bars are presented in Figure 4, but it is unclear whether these data represent mean {plus minus} SEM or something else, and the number of biological replicates is unclear. What statistical test was performed? Are differences statistically significant? This information should be presented for all figure panels.

eLife. 2016 Aug 19;5:e16093. doi: 10.7554/eLife.16093.009

Author response

1) Copper binding to His358 is inferred from indirect studies, but is not directly demonstrated – this should be acknowledged. The dose response of copper (Figure 1) shows that 1µM had no effect, 10µM shows an inhibitory effect, and that 100µM was saturating. How does this dose response compare with physiological intracellular copper concentrations?

We have acknowledged in the revised Discussion that we do not have direct evidence of copper binding to His358.

We typically do our patch clamping in the presence of EGTA to buffer cytosolic and bath calcium. As pointed out by the reviewers, EGTA is also a potent copper chelator. We have now repeated the inside/out (I/O) patch clamp experiments without EGTA (Figure 2D) so as to more accurately determine the lowest free copper concentration that inhibits. These experiments demonstrated that 100nm CuCl2 inhibits KCa3.1, which is a 100-fold lower than with EGTA present.

Most of the intracellular copper is sequestered, and free intracellular copper levels are very low. This has led to much speculation as to how copper metalloenzymes become loaded with copper. Some of the ideas include copper metallochaperones that function to load copper onto enzymes and presumably KCa3.1. Thus, it is very difficult, and likely not relevant, to extrapolate the amount of copper needed to inhibit in vitro (I/O patches) to intracellular copper concentrations.

2) The concentration of free Cu++in the solutions used for the whole cell and inside-out patch clamp experiments is a concern. The expected Cu++concentration in the pipette solution used in whole cell experiments and in the bath solution for the inside-out experiments should be specified. EDTA has a binding affinity of 10-18M for Cu++and it is likely to be the same for EGTA. Under such conditions, both solutions should be Cu++free and yet channel activity appears to be low in the whole cell and inside-out recordings presented in Figures 1 and 2 in control conditions.

This is an excellent point raised by the reviewers. See response #1 above.

3) Figure 2A shows that channel activity remains high after TTM removal in 300nm Ca2+ (the effect of TTM is not reversible) consistent with the bathing solution being Cu++free and TTM being capable of chelating Cu++from the Cu++-H358 complex. However, channel activity appears low in 300nm Ca2+ after TTM treatment in 30nm Ca2+. One would have expected channel activity to be high in this case if TTM had succeeded to chelate Cu++from the Cu++-H358 complex in 30nm Ca2+ conditions. The authors need to discuss how the action of TTM might be modulated by Ca2+ and may involve more than chelating Cu++.

This is another excellent point raised by the reviewers, which we missed. We have now repeated the same experiment with NDPK-B and found that treatment with NDPK-B (and GTP) followed by high calcium also does not activate KCa3.1. Rather, calcium needs to be present simultaneously for both TTM and NDPK-B to activate. This makes perfect physiological sense as this requirement would prevent aberrant activation of the channel under sub-threshold stimuli. We hypothesize that a calcium/calmodulin-induced conformational change in the channel provides access to His358 for phosphorylation by NDPK-B. We have modified our model to incorporate this finding, which is presented in the new Figure 6 and in the Discussion.

4) Segments of the inside-out recording in Figure 2A, B and D should also be presented at a higher time resolution to compare single channel activity at 300nm Ca2+ + low Cu++, 10μM CuCl2 and TTM conditions. The authors also need to comment on the finding that in Figure 2A, B and D, channel activity appears greater in CuCl2 conditions than in control before TTM application in 300nm Ca2+ conditions.

The data shown in Figures 2 and 3 are mean patch current. The single channel activity will not be apparent with higher time resolution due to the existence of many channels in each patch. If the reviewers would like, we can also present the results as mean patch in the form of a bar graph for additional clarity.

High calcium results in activation of other potassium channels in the I/O patch that are not KCa3.1, which is evident from the failure of the KCa3.1 inhibitor to block this current. This finding provides further evidence that copper specifically inhibits KCa3.1, as copper only inhibits the TRAM-34-inhibitable current (KCa3.1) and not the non-KCa3.1 current induced by calcium. This is not evident in the other tracings, as baseline current shown is in 300nm calcium, not 30nm, as in the old Figure 2A and the new Figure 3A.

5) The proposed mechanism implies that Cu++coordination by four H358 is state dependent, with Cu++binding taking place in the channel closed configuration. Accordingly, the potency of CuCl2 to decrease channel activity following TTM treatment should be Ca2+ dependent with CuCl2 being less potent in saturating Ca2+ conditions (10μM Ca2+). The Ca2+ dependency of CuCl2 mediated inhibition after TTM treatment should be presented in inside-out experiments to confirm state dependency.

We have now performed this experiment and indeed found that copper inhibition of KCa3.1 is dependent on calcium levels, with a 10-fold higher concentration of copper (1μM) required to inhibit the channel under saturating calcium concentrations (new Figure 2E). These results have been incorporated into our model (see Discussion and the new Figure 6).

6) The significance of the results illustrated in Figure 3A–B would be clearer if information on the mechanisms underlying Cu++cellular homeostasis, and on the expected basal Cu++intracellular level is provided. If channel expression is not affected in Ctr1-/- cells, why is there a two fold difference in current intensity following TTM treatment between WT and Ctr1-/- cells assuming that TTM can chelate all the Cu++in the cells?

See response #1 above with respect to free copper levels in cells. We think that the difference in activity is due to the failure of TTM (applied exogenously) to chelate all of the copper bound to KCa3.1 in WT MEFs. In fact, this is consistent with our findings that calcium binding to calmodulin is required for TTM to chelate copper away from KCa3.1 (new Figure 3A). Since intracellular calcium concentrations are unlikely to be saturating, TTM should not be able to chelate all of the channel-bound copper.

7) The structural constraints related to Cu++coordinating four His358 residues should be discussed. What are the minimal distances between adjacent imidazole nitrogen atoms to insure Cu++coordination? How does the proposed structural model for the channel closed state impact on the possibility of a dimer-dimer structure for the channel open state? The model presented in Figure 5 presumes that this dimer-dimer structure does not prevail in the channel open state configuration.

As mentioned in the previous version of the manuscript, we have unpublished crystallographic data showing that the C-terminal region of KCa3.1 forms a four-helix bundle. Suggestively, in this structure, His389 (31 residues C-terminal to the phosphorylation site, His358) occupies an inward-facing ‘a’ position in the heptad repeat, and the four copies of His389 (via N3 in the imidazole ring), along with an axial water molecule, bind a Cu(II) ion on the four-fold axis. (CuCl2 was added prior to crystallization trials.) The N3 (imidazole ring)-Cu(II) distance is 2.0Å (C4 symmetry–all four the same), and the O (H2O)-Cu(II) distance is 2.4Å. This structure demonstrates that four histidine residues in the ‘a’ position of a four-helix bundle (in a homotetramer) are capable of binding a Cu(II) ion on the four-fold axis. Based on coiled-coil prediction software (see Minor Point #5 below), our model is that His358 is also positioned in an ‘a’ position in a four-helix bundle, coordinating a Cu(II) ion on the four-fold axis similar to His389.

Because our crystal structure of the C-terminal four-helix bundle is unpublished, and because of the confusion that could arise by showing copper coordinated by His389 instead of His358 (the subject of the present study), we prefer to simply state in the Discussion that His358 is predicted to occupy an inward-facing ‘a’ position, and that the four copies would be positioned to coordinate (via N3) a Cu(II) ion on the four-fold axis.

For several reasons, we believe that the 2:2 SK2 (KCa2.2):calmodulin (CaM) structure reported by Schumacher et al. (Nature 410, 1120 (2001)) is artifactual and that their dimer-dimer model for the open-channel state is not correct. The fundamental reason for believing their model is incorrect is that the dimer-dimer model is incompatible with the C4 symmetry of the homotetrameric SK/KCa channels. We believe that the 2:2 complex they observed in solution and in the crystal structure (with calcium present) is an artifact of the C-terminal 6xHis-tag included, which contained a heterologous leucine residue (LEHHHHHH) that interacts with the second molecule of CaM in the dimer (i.e., this might help to stabilize the 2:2 complex). Our solution data (unpublished), as well as data published by Halling et al. (J. Gen. Physiol. 143, 231 (2014)), show unequivocally a 1:1 SK2 (or SK4):CaM complex, with or without calcium present. Four 1:1 complexes can be assembled into a structure that is consistent with the C4 symmetry of the channel. Such a model was proposed by Sachyani et al. (Structure 22, 1582 (2014)) for Kv7.1, which is also activated by calcium-CaM and shows sequence similarity with the KCa channels in the CaM binding domain. We favor this model for KCa3.1-CaM in the open state, which is why this paper and not the Schumacher et al. (2001) paper is referenced in the new Figure 6 legend.

8) The increase in KCa3.1 channel activity in Ctr1-/+ CD4 T cells was predicted by the patch clamping studies. However, the increase in His358 phosphorylation was not predicted. This finding raises questions concerning mechanism. Is His358 phosphorylation a passive response to lack of copper binding to His358? This response differs from the major conclusion of this study – that phosphorylation prevents copper interactions with KCa3.1. The observation suggests a more complicated relationship between copper binding and histidine phosphorylation than the authors' conclusions.

We have now addressed these points in the model presented in the Discussion and in the new Figure 6.

9) Very limited information is provided concerning statistical analysis. For example, bar graphs and error bars are presented in Figure 4, but it is unclear whether these data represent mean {plus minus} SEM or something else, and the number of biological replicates is unclear. What statistical test was performed? Are differences statistically significant? This information should be presented for all figure panels.

Detailed statistical methods along with the significance levels have been added to figure legends where indicated.

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