 |
INTRODUCTION |
Both voltage-gated and Ca2+-activated K+
(KCa)1 channels
are widely expressed in immune cells, including human T lymphocytes. Drugs that block voltage-gated Kv1.3 channels inhibit T lymphocyte activation and proliferation, volume regulation, and cell-mediated cytotoxicity (for review, see Ref. 1). Inasmuch as these functions involve Ca2+ influx through channels activated by depletion
of Ca2+ stores, one widely proposed role for K+
channels is to maintain a negative membrane potential and large driving
force for Ca2+ entry. However, the relative roles of
KCa versus Kv1.3 channels in these cell
functions are not known, partly owing to the previous lack of potent
KCa blockers that do not also block Kv1.3 channels.
Two KCa channels have been found in lymphocytes and
lymphocytic cell lines. They differ in biophysical and pharmacological properties (2-6). An apamin-sensitive, small conductance channel (7-8
pS) is the prevalent KCa channel in the commonly used
Jurkat T cell line (4) and is also present in rat T and human B
lymphocytes (2, 6). However, a corresponding apamin-sensitive
whole-cell current has not been identified in normal human T cells,
perhaps a result of channel rundown we observed after cell disruption (2). Instead, a KCa channel we first described (2, 3) is
the prevalent KCa channel in resting and activated human T lymphocytes. It is a charybdotoxin-sensitive, inwardly rectifying channel (15-35 pS in symmetrical K+ solutions (5, 6)) that
is commonly called "IK," for intermediate conductance
KCa. Recently, a molecular candidate for IK was cloned from
a human placental cDNA library (hSK4 (7)) and subsequently from
human pancreas (hIK1 (8)) and a human lymph node library (hKCa4
(9)).
IK current increases in the 3-4 days following activation of human T
cells (5). Thus, it is anticipated that this KCa current will be especially important for secondary immune responses of activated T cells (lymphoblasts), including proliferation and volume
regulation. The regulatory volume decrease (RVD) that follows T cell
swelling is known to depend on K+ (and Cl
)
channels (10-12). Although Kv1.3 is involved in RVD in some resting T
cells (13), the relative contribution of IK versus Kv1.3
channels is not known, either in resting human T cells or in
lymphoblasts. We previously reported that intracellular
Ca2+ rises immediately after human T cells are exposed to a
hypotonic shock (14); thus, we predicted that KCa currents
would also subserve RVD.
In the present study we cloned hSK4 from human T lymphoblasts,
expressed the channels stably in CHO cells, and compared the salient
biophysical and pharmacological properties of the native and cloned
channels. All intrinsic properties examined were indistinguishable, supporting the view that hSK4 homotetramer forms the
subunit of the
IK channel of lymphoblasts. We found that hSK4 mRNA expression is
strongly up-regulated after T cell activation; thus we predicted (and
observed) an increased role for IK current in lymphoblasts compared
with resting T cells. Although hSK4 is functionally a KCa
channel, that is activated by a rise in intracellular Ca2+,
it was recently reported that brain SK channels are not gated directly
by Ca2+ but rather by Ca2+ interacting with
calmodulin that is irreversibly bound to the channel (15). We have
presented preliminary data showing that the lymphoblast IK current is
inhibited by antagonists of calmodulin and CaM kinase (16, 17). We now
show details of this inhibition of native IK current and that
calmodulin binds directly to the hSK4 channel protein in a
Ca2+-dependent manner. The CaM binding domain
resides in the proximal part of the C terminus, since binding to this
region occurs in the absence of flanking sequence and is eliminated in
constructs lacking this region. Unlike the study of heterologously
expressed brain SK channels (15), we provide evidence for additional
modulation of hSK4 in lymphocytes, results that have important
implications for the existence of accessory molecules and cell-specific
KCa channel regulation.
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EXPERIMENTAL PROCEDURES |
Cells--
Human peripheral blood mononuclear cells were
isolated on a Ficoll-Paque (Amersham Pharmacia Biotech, Baie D'Urfé,
Quebec, Canada) density gradient. To purify resting T cells, monocytes and B lymphocytes were removed by adhering them to a nylon wool column
for ~1 h and then eluting the T cells, which were then placed in
culture medium containing RPMI 1640 with L-glutamine (Life
Technologies, Inc., Burlington, Ontario, Canada) supplemented with 10%
fetal bovine serum (Sigma, Mississauga, Ontario, Canada) and 50 µg/ml
gentamicin sulfate (Life Technologies, Inc.). Mitogenically activated T
lymphocytes (lymphoblasts) were prepared by treating the mixed
mononuclear cells with 7 µg/ml phytohemagglutinin (PHA-P, Sigma) and
growing them in culture medium in a humidified 37 °C incubator with
5% CO2 for 3-4 days. PHA selectively stimulates T cell
proliferation, and monocytes die during prolonged culture.
Cloning and expression of hSK4 in CHO cells was as described previously
(7). The full-length hSK4 clone was isolated from a human placental
cDNA library, subcloned into pcDNA3 (Invitrogen, Carlsbad, CA),
and transfected, using LipofectAMINE (Life Technologies, Inc.), into
Chinese hamster ovary (CHO) cells that had been grown to ~50%
confluency. Stable transfectants were selected using Geneticin (Life
Technologies, Inc.) and subsequently separated into clonal populations
by single-cell sorting (FACSIV, Yale Cell-sorting Facility). Two cell
lines with similar current densities were selected for
electrophysiological characterization. These were grown in Iscove's
modified Dulbecco's medium, supplemented with 10% fetal bovine serum,
HT supplement, antibiotic/antimycotic, and 1 mg/ml Geneticin (all
reagents from Life Technologies, Inc.).
Preparing RNA Probes--
Total RNA was isolated from resting
and activated human T lymphocytes, rat lung, and human placenta using
the guanidinium isothiocyanate method (18) and subjected to DNase I
digestion (0.1 units/ml, 15 min, 37 °C; Amersham Pharmacia Biotech,
Toronto, Ontario, Canada) to eliminate genomic contamination. First
strand cDNA was synthesized according to the manufacturer's
instructions (Amersham Pharmacia Biotech) using an oligo(dT)-based
primer. The cDNA was then used as a template for PCR reactions
using the following primers: Kv1.3 (GenBankTM accession
number M30312) forward primer 5'
AATGAGTACTTCTTCGACCGCAACAGACCCAGCTTCGA 3' and reverse primer 5'
CCAATGAAAAGGAAAATGAGCAGCCCCAG 3'; hSK4 (GenBankTM accession
number AF000972) forward primer 5' GTGCGTGCAGGATTTAGG 3' and reverse
primer 5' TGCTAAGCAGCTCAGTCAGGG 3'. The PCR reaction was conducted with
1.5 mM MgCl2, 0.5 µM forward and
reverse primers, and 10% of the cDNA reaction mixture, using a
Minicycler system (MJ Research, Watertown, MA). After incubating the
mixture at 85 °C for 1 min, 1.25 units of Taq DNA
polymerase (Sangon Ltd., Toronto, Ontario, Canada) was added and heated
to 94 °C for 3 min, followed by 30 cycles through a 1-min
denaturation step at 94 °C, a 1-min annealing step at 50 °C, and
a 3-min extension step at 72 °C. A final extension for 5 min at
72 °C was followed by incubation at 4 °C until further processing
of samples. After PCR, the DNA products were resolved in 2% agarose
gels containing 0.5 mg/ml ethidium bromide. The identities of
PCR-amplified fragments of the predicted sizes (856 bp for hSK4 and 790 bp for Kv1.3) were confirmed by restriction endonuclease digestion,
which yielded the correct size bands on an agarose gel. By using
additional hSK4 PCR primers, we then amplified cDNA encoding the
entire open reading frame (1284 bp). Cloning and subsequent sequencing
revealed 100% homology with the published amino acid sequences of
hSK4/hIK1/hKCa4 (7-9).
Ribonuclease Protection Assays--
RNase protection assays on T
cells, lung, and placenta were performed using the amplified cDNA
fragments for hSK4 and Kv1.3 as probes. To obtain RNA probes, these
fragments were linearized with SmaI (for Kv1.3) or
MluI (for hSK4) and in vitro transcribed with
[
-32P]dUTP and T7 RNA polymerase, yielding RNA
transcripts of 579 (Kv1.3) and 254 bp (hSK4). However, since about 66 bp of non-hybridizing vector sequence was included in each probe, the
protected channel fragments should be 513 (Kv1.3) and 188 bp (hSK4). A
control plasmid (mouse
-actin cDNA), purchased from Ambion and
transcribed according to the supplier's instructions, should yield an
RNase-protected fragment of 250 bp. Because of the great abundance of
-actin mRNA in these cells, the
-actin probe was labeled to a
low specific activity (~600-fold lower than hSK4 and Kv1.3) to allow
simultaneous quantitation. For each experimental data point, 10 µg of
total RNA was used, with 5 µg of yeast tRNA as a negative control for probe self-protection bands. Following hybridization and RNase digestion, the samples were electrophoresed in polyacrylamide gels,
dried, and exposed overnight to x-ray film (X-Omat, Eastman Kodak Co.).
Specific signals were quantified by densitometic analysis of the
developed film using a Bio-Rad model Gs-670 densitometer, and results
are expressed as mean ± S.D. with statistical analyses using the
Student's t test.
hSK4-Flag DNA Constructs--
An XhoI site was added
by PCR to the C terminus of the full-length hSK4, and then a pair of
Flag-encoding oligonucleotides was spliced into this site. This
construct was used to create a second proto-construct in which all
transmembrane domains were eliminated, and a consensus Kozak sequence
was added to the beginning of the cytoplasmic C terminus. Morph
mutagenesis (5 Prime
3 Prime, Inc., Arapaho, CO) was used to add a
second, silent EcoRV site and a second XhoI site
to the C termini of each proto-construct. Derivatives of these
proto-constructs (for details, see text accompanying Fig. 7) were
created by cutting out the fragments flanked by either EcoRV
or XhoI and religating the larger fragment.
Calmodulin Affinity Chromatography--
CHO cells stably
expressing Flag-tagged hSK4 constructs were grown to confluency in
100-mm Petri dishes. To assess binding of hSK4 protein to
calmodulin-Sepharose, we used methods modified from Chapin et
al. (19). The dishes were washed three times in cold
phosphate-buffered saline containing calcium and magnesium. Then, 1 ml
of ice-cold solubilization buffer was added, which contained 50 mM Tris-HCl, pH 7.4, 150 mM NaCl, 1 mM MgCl2, 1 mM CaCl2,
1% Triton X-114 (Sigma), and protease inhibitors (5 µg/ml pepstatin,
10 µg/ml chymostatin, 5 µg/ml leupeptin, 10 µg/ml antipain, 500 µM benzamidine, 0.1% Trasylol), and the dishes were
rotated for 15 min at 4 °C. The lysate was triturated with a
23.5-gauge needle, and after removing insoluble material by
centrifugation (15 min, 4 °C) the supernatant was transferred to a
new tube to which 6 µl of 0.5 M EGTA was added (final
concentration, 3 mM) to chelate the calcium. To promote
Triton X-114 phase partitioning, this solution was warmed to 37 °C
for 3 min and then centrifuged (5 min, room temperature) at full speed
in a microcentrifuge. The detergent phase (bottom) was resuspended (5 min, 4 °C) in 900 µg of cold calcium-free wash buffer (50 mM Tris, pH 7.4, 150 mM NaCl, 1 mM
MgCl2, 3 mM EGTA). The entire phase-separation
procedure was repeated, and then the detergent phase was diluted 9:1 in an ice-cold "binding" buffer containing calcium (50 mM
Tris, pH 7.4, 150 mM NaCl, 1 mM
MgCl2, 0.5 mM CaCl2) and incubated
for 5 min at 4 °C. The protein samples were pre-cleared (1-3 h,
4 °C) with 50 µl of Sepharose Cl-2B.
The pre-cleared sample (500 µl) plus binding buffer (450 µl) was
added to a 20% slurry (50 µl) of calmodulin-conjugated agarose beads
(Sigma) that had been equilibrated through a series of buffers, ending
with binding buffer. The entire sample in binding buffer was incubated
overnight at 4 °C on a rotator. For CaM antagonist competition
experiments, 1000 nM calmidazolium was added to the sample
slurry before the overnight incubation. The beads were pelleted by a
brief microcentrifuge spin (~5 s, 4 °C) and washed with 1 ml of
binding buffer containing the detergent, Triton X-100, first at 0.5%
and then at 0.05%. This was followed by a 5-min incubation at 4 °C
in the 0.05% Triton X-100 solution, with or without 3 mM
EGTA. The beads were harvested as before and washed once with
detergent-free binding buffer with or without EGTA. The proteins were
dissociated from the calmodulin-conjugated agarose beads by boiling
them 5 min in 60 µl of Laemmli buffer. The resulting protein sample
was analyzed by SDS-polyacrylamide gel electrophoresis and Western
blotting using a biotinylated anti-Flag antibody and an
avidin-horseradish peroxidase-coupled secondary antibody (both from Sigma).
Patch Clamp Electrophysiology, T Cells--
Activated T
lymphoblasts were used 3-4 days after PHA stimulation, at which time
IK current amplitudes are much larger than in resting T cells at the
same cytoplasmic Ca2+ concentration (5). Whole-cell
currents were measured using an Axopatch 200 amplifier, with 8-12 M
pipettes. During data acquisition, capacitive currents were canceled by
analogue subtraction, 50-70% series resistance compensation was used,
and all currents were filtered at 2 kHz. All voltages were corrected to
account for the junction potential between bath and pipette solutions. The bath contained the following (in mM): 145 sodium
aspartate, 5 KCl, 1 MgCl2, 1 CaCl2, 5 HEPES,
adjusted with NaOH to pH 7.4, 270-283 mosmol. The K+
selectivity was verified and current rectification examined by replacing the sodium aspartate with 80 mM sodium aspartate,
65 mM potassium aspartate (70 mM K+
solution), or 145 mM potassium aspartate (150 K+ solution). To activate fully the IK channels (5), we
used a high Ca2+ (1.1 µM) pipette solution,
consisting of the following (in mM): 140 potassium
aspartate, 1 K4BAPTA, 2 K2ATP, 0.9 CaCl2, 1 MgCl2, 5 HEPES adjusted to pH 7.2 with
KOH, 260-270 mosmol. Contributions from volume-sensitive
Cl
currents (20) were small since aspartate was used in
the bath and pipette, and the internal solution was slightly
hypo-osmotic. Fresh K2ATP (Sigma) was always added to
pipette solutions just before use to help maintain channel and
second-messenger activity during whole-cell recording.
CHO Cells--
Stably transfected CHO cells were passaged every
3-4 days using trypsin/EDTA (Life Technologies, Inc.). Recordings were
made 1-2 days after replating, using an Axopatch 1D amplifier (Axon Instruments). Pipettes had resistances of 3-5 M
, on-line
capacitance compensation, and 60-80% series resistance compensation
were used, and data were filtered at 2 kHz. The bath contained (in
mM) 140 NaCl, 5 KCl, 1 CaCl2, 29 glucose, 25 HEPES, pH 7.4, and the pipette contained 32.5 KCl, 97.5 potassium
gluconate, 5 MgATP, 4.3 CaCl2 (free Ca2+, 1 µM), 5 EGTA, 20 HEPES, pH 7.2. Conventional voltage clamp recordings were used to measure whole-cell K+ currents and
current clamp recordings used to measure membrane potential.
Free Ca2+ concentrations were calculated assuming a
dissociation constant (Kd) of 10
11 for
the EGTA4-·2 Ca2+ complex and
10
7 M for the BAPTA4-·2
Ca2+ complex at pH 7.2, and allowing for the weak calcium
binding by ATP. When calmodulin (CaM) was added to the pipette,
CaCl2 was increased to 0.91 mM (rather than
0.90 mM) to maintain the free Ca2+
concentration (Ca2+i) at 1.1 µM,
according to the following chemical information. Ca2+ binds
to two globular domains in CaM, each with two Ca2+-binding
sites (E-F hands) for a total of four binding sites (21). In principle,
CaM can reduce free Ca2+i, but the effect of adding
50 µM CaM will be very small, as follows. At
physiological KCl concentrations (K+ competes with
Ca2+ binding to some extent), the affinity of CaM for
Ca2+ is ~100-fold lower than that of BAPTA, which is
present at 1 mM in our pipette solutions. Since the mean of
log Kd is 5 for CaM (22) versus 7 for
BAPTA, the effect of 50 µM CaM on free
Ca2+i is equivalent to adding a further 0.5 µM BAPTA.
Data analysis was performed using pCLAMP (version 5.5 or 6, Axon
Instruments, Foster City, CA), SigmaPlot (version 2, Jandel, San
Rafael, CA), and Origin (version 4.1, Microcal, Northampton, MA). Where
appropriate, data are presented as mean ± S.E., with paired
Student's t tests (when each cell acted as its own control) or unpaired t tests used to determine the statistical
significance of differences (95% confidence interval). All recordings
were made at room temperature (19-22 °C), except when KN-62 and
KN-04 were used at 37 °C.
Cell Proliferation--
Activated lymphoblasts (stimulated for 3 days) and resting (naive) T cells were prepared as above and then
seeded at 2 × 104 cells/well in 96-well plates
(Corning Glass, Corning, NY) in culture medium. Cells were first
incubated with or without K+ channel blockers for 10 min,
and then 7 µg/ml PHA-P was added to either initiate or re-stimulate
proliferation. After a further 3 days growth, the plates were spun
(1,200 rpm for 15 min at room temperature); the supernatant was
removed, and the plates were frozen at
80 °C (to aid subsequent
cell lysis). Proliferation was calculated from the change in total
nucleic acid content in each well using an assay (CyQUANT kit,
Molecular Probes, Eugene, OR) that measures the signal generated by
binding of a fluorescent dye to nucleic acids, a signal that increases
in proportion to the number of cells. Fluorescence signals were read
from a 96-well plate using a plate-reader (CytoFluor II, PerSeptive
Biosystems Inc., Framingham, MA) with excitation at 485 nm and emission
at 530 nm. For each treatment, the average fluorescence from control wells (lymphocytes without mitogen or channel blocker) was subtracted from the average final fluorescence from treated wells to yield the
change in fluorescence (nucleic acid content) during 3 days of
proliferation. Results are expressed as the mean increase in fluorescence ± S.D. (n = 4 experiments, 4 replicates/experiment), and a Bonferroni multiple comparison test was
used for statistical analysis.
Flow Cytometric Analysis of Regulatory Volume Decrease
(RVD)--
Changes in cell volume were measured using a flow cytometer
(FACScan, Becton Dickinson, CA) by monitoring changes in right angle
light scattering (side scatter) as an index of cell volume (23). To
indicate percent change from the initial volume, we calculated a
swelling index as (1/SSCh)/(1/SSCi) × 100, where SSCh is the average side scatter at each time after
exposure to hypotonic medium and SSCi is the average value
in isotonic medium. The flow cytometer was set to exclude dead cells
and debris by omitting cells that stained with propidium iodide (0.04 µg/ml), a nuclear marker excluded by live cells.
For control volume measurements, resting or activated T cells were
suspended in isotonic medium which contained the following (in
mM): 140 NaCl, 5 KCl, 1 MgCl2, 1 CaCl2, 10 HEPES, pH 7.4, 285 mosmol. Initial measurements
were taken from 5000 live cells, and then aliquots of the same cells
were exposed to a hypotonic medium (56% of normal osmolarity) with or
without K+ channel blockers. The hypotonic medium contained
the following (in mM): 70 NaCl, 5 KCl, 1 MgCl2,
1 CaCl2, 10 HEPES, pH 7.4, 159 mosmol. Side scatter was
recorded from 5000 live cells every 30 s for the first 3 min and
then at 6 min after the hypotonic shock. Whenever K+
channel blockers were used, the initial volume was measured in the
presence of drug before exposing the cells to hypotonic medium. Data
were analyzed using CellQuest software (version 3.0.1f, Becton Dickinson). Values are presented as mean ± S.D. of at least 4 experiments per treatment, and a Bonferroni multiple comparison test
was used for statistical analysis. Experiments were performed at room
temperature (21-23 °C).
Chemicals--
We used the K+ channel blockers,
charybdotoxin and iberiotoxin (purchased from Peptides International,
Louisville, KY, or a gift from V. Gribkoff and S. Dworetzky,
Bristol-Myers Squibb Co.), margatoxin and agitoxin-2 (Alomone
Laboratories, Jerusalem, Israel), and apamin,
d-tubocurarine, and clotrimazole (Sigma). The calmodulin antagonists, trifluoperazine and W-7
(N-(6-aminohexyl)-5-chloro-1-sulfonamide) were from Sigma.
Bovine brain calmodulin, rat brain CaMKII, calmidazolium (compound
R24571), and KN-62
(1-[N,O-bis(5-isoquinolinesulfonyl)-N-methyl-L-tyrosyl]-4-phenylpiperazine) were from Calbiochem. KN-04
(N-(1-[N-methyl-p-(5-isoquinolinesulfonyl)benzyl]-2-(4-phenylpiperazine)ethyl]-5-isoquinolinesulfonamide), an analogue of KN-62 without CaM kinase inhibitor activity, was purchased from Seikagaku America (Rockville, MD). These reagents (except KN-62 and KN-04) were prepared in bath saline, frozen in
aliquots, and thawed just before use. KN-62 and KN-04 stock solutions
were prepared in Me2SO and diluted in bath saline before use. The Me2SO concentration in the final bathing solution
was
0.5% for T cells and
1% for CHO cells. Whenever cells were
treated with KN-62, control cells were incubated in the same
concentration of Me2SO for the same duration.
 |
RESULTS |
Up-regulation of hSK4 and Kv1.3 mRNA Expression in Activated T
Lymphocytes--
We amplified DNA corresponding to full-length
transcripts of hSK4 and Kv1.3 channels from activated human T
lymphocytes. The open reading frame of the full-length hSK4 clone was
1284 bp, which encodes a protein of 428 amino acids that is 100%
identical to the recently cloned hSK4 (7), hIK1 (8), and hKCa4 (9). RNase protection assays (Fig. 1,
A-C) on total RNA from resting and activated human T cells
were used to determine if changes in mRNA expression correlate with
previously observed increases in amplitude of the two K+
currents in lymphoblasts. Human placenta and rat lung were used as
positive controls, and yeast tRNA was used as a negative control. In
lymphoblasts, there was a 14.6-fold increase in hSK4 mRNA
(p < 0.001, n = 4) and a 1.3-fold
increase in Kv1.3 mRNA (p < 0.001, n = 4) 3-4 days after mitogenic stimulation.

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Fig. 1.
Expression of hSK4 and Kv1.3 channels.
A and B, hSK4, Kv1.3 and actin mRNA
expression. RNase protection assays showed protected fragments of the
correct size in resting and activated human T cells, with rat lung and
human placental tissue as positive controls. Activated T cells were
used on days 3-4 after mitogenic stimulation. Total RNA (10 µg) from
the indicated cells and tissues was hybridized to 6 × 104 cpm of 32P-labeled hSK4 or Kv1.3 probe and
1 × 103 cpm of actin probe. Left-hand
panels show the full-length probe and self-protected bands
determined using 5 µg of yeast tRNA as a negative control. See
"Experimental Procedures" for details and predicted band sizes.
C, histogram showing substantial up-regulation of hSK4
mRNA (left-hand bars) in activated T lymphocytes
(14.6-fold, p < 0.001) and a small increase in Kv1.3
mRNA (right-hand bars, 1.3-fold, p < 0.001). Values are densitometer readings (mean ± S.E., four
independent mRNA samples) normalized to -actin counts in the
same samples. D, co-existing IK and Kv1.3 currents in
activated human T lymphocytes: typical currents for a cell 3 days after
PHA stimulation. The pipette solution was 150 mM potassium
aspartate solution, buffered to 1.1 µM free
Ca2+, and the holding potential was 95 mV after
correcting for junction potentials (see "Experimental Procedures").
Upper panel, total current obtained by applying 300-ms steps
at 20-mV intervals between 65 and +15 mV. Control current was
measured 5 min after attaining the whole-cell configuration, and then
margatoxin (MgTx, 2.5 nM) was added to the bath at 15 min
to block Kv1.3, and charybdotoxin (ChTx, 15 nM) was added
at 30 min to block KCa, leaving the "leak" (anion)
current. Lower panel, current versus voltage
(I-V) relations obtained from the same cell using a voltage ramp
protocol, before and after addition of the peptide toxins, as above.
All voltage ramps began with a step to 125 mV, followed immediately
by a 225- or 300-ms ramp to +40 mV.
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Biophysical and Pharmacological Properties of the Lymphoblast IK
Are the Same as hSK4 Expressed in CHO Cells--
Charybdotoxin (ChTx)
is an effective blocker of native IK currents in lymphocytes (3, 5) and
of expressed hSK4/hIK1/hKCa4 channels (7-9). In lymphoblasts, with
free Ca2+ in the pipette buffered to 1.1 µM
to maximally activate IK, typical current responses to voltage steps or
ramps were resolved into the following three components: Kv1.3, IK, and
an anion current (Fig. 1D). Separation of the currents was
achieved by first blocking Kv1.3 with margatoxin (MgTx, 2.5 nM) and then adding ChTx (15 nM) to block IK,
leaving the anion current (Cl
Nernst potential about
22
mV) which we have previously characterized (20). For subsequent IK
measurements we blocked Kv1.3 with 2.5-5 nM MgTx, measured
the remaining current, and then blocked IK with 10-20 nM
ChTx and subtracted the remaining anion current. (This procedure was
not necessary for stably transfected CHO cells, wherein endogenous
currents were negligible compared with the large hSK4 currents.)
The ChTx-sensitive currents (Fig.
2A) were
K+-selective, as seen from the intersection of their
current versus voltage (I-V) curves with the anion current
(at
78 mV; Nernst potential, EK =
86 mV). We
further confirmed their K+ selectivity using high
K+ bathing solutions, wherein the reversal potentials of
both Kv1.3 and IK were commensurate with changes in the external
K+ concentration: at 70 mM, reversal potential,
Erev =
15 to
17 mV, and at 150 mM Erev =
2 to 0 mV (after
junction potential corrections). Expressed hSK4 and native IK currents
were not time- or voltage-dependent, even during long
voltage clamp steps (Figs. 1D and 2, B and
C) (5). Their whole-cell I-V relations (Fig. 2A)
were nearly linear over a wide voltage range under physiological Na+/K+ gradients (also Fig. 1D) but
rectified inwardly with high K+ concentrations in the bath
and pipette. These biophysical features are consistent with previous
studies of native IK in lymphocytes (5, 24), and inward rectification
in high K+ was observed for the channels expressed in
Xenopus oocytes (hIK1 (8)) and in HEK cells (hKCa4 (9)). For
the remaining experiments, whole-cell recordings from both cell types
were made with high concentrations of external Na+ and
internal K+.

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Fig. 2.
Comparison of hSK4 stably expressed in CHO
cells, with native IK current in activated T lymphocytes. For
this, and all subsequent figures showing IK currents from lymphoblasts,
5 nM MgTx was present to block Kv1.3 current. A,
hSK4 whole-cell I-V relations were plotted from currents in response to
voltage steps, whereas native IK currents were from voltage ramps (as
in Fig. 1D). The bath K+ concentrations were 5 or 145 mM for CHO cells, 5 or 150 mM for
lymphoblasts, yielding calculated EK values of
86 mV and ~0 mV. B, charybdotoxin blocks hSK4 and IK
currents. Currents in response to voltage clamp steps between 120 and
+20 mV show profound block by 20 nM ChTx, with a slight
relief over time at positive potentials in CHO cells. C,
clotrimazole blocks both hSK4 and IK in a voltage- and time-independent
manner. Currents in response to voltage clamp steps as in B.
D, dose-response curves show fraction of current remaining
(mean ± S.E.).
|
|
For comparison with the literature we tested ChTx (Fig. 2, B
and D) and found it to be a potent blocker of hSK4 in CHO
cells (IC50 = 1.7 nM, n = 7)
and of IK in T lymphoblasts (IC50 = 6 nM, n = 3). For expressed hSK4 the dose dependence was
calculated from the initial block during each voltage clamp step, to
avoid the time- and voltage-dependent relief of block at
depolarized potentials (Fig. 2B). Clotrimazole blocks
Ca2+-activated K+ fluxes in thymocytes and red
blood cells (IC50 50 nM (25)) and hIK1/hKCa4
channels expressed in Xenopus oocytes (Kd ~25 nM (8)) and HEK cells (Kd 387 nM (9)). Clotrimazole effectively blocked IK in
lymphoblasts (IC50 = 40 nM) and hSK4 in CHO
cells (IC50 = 56 nM), with no time or voltage
dependence (Fig. 2, C and D). Although some brain
SK channels are potently blocked by apamin and
d-tubocurarine (26, 27), neither hSK4 nor the lymphoblast IK
current was sensitive to these drugs (apamin, IC50
100
nM; d-tubocurarine, IC50
250
µM). Iberiotoxin, a potent inhibitor of large conductance
KCa channels, did not block hSK4 or IK (IC50
>200 nM, data not shown).
For functional studies of proliferation and volume regulation in
lymphocytes, we avoided ChTx since it blocks both IK and Kv1.3 currents
with similar potencies. Margatoxin is much more selective for Kv1.3 but
can reduce IK at high concentrations (>10 nM, data not
shown); thus we used agitoxin-2, which is more potent (Kd ~200 pM) and does not block IK
(28). IK was selectively blocked with clotrimazole. Drug concentrations
were chosen to block different amounts of the two currents. To
eliminate essentially all Kv1.3, we used 5 nM AgTx-2 (~25
Kd), whereas to allow us to test for additive
effects of IK block, we used 250 nM clotrimazole (~6
Kd for IK). Peptide toxins are especially useful since they are not membrane-permeant, so are unlikely to affect other
intracellular processes. Although clotrimazole is membrane-permeant and
also inhibits cytochrome P-450 (25), there is no evidence for P-450
involvement in lymphocyte proliferation or volume regulation.
Proliferation Is Inhibited by Combined IK and Kv1.3 Channel
Block--
We first stimulated freshly isolated (naive) human T cells
with the mitogen, phytohemagglutinin (PHA-P), and then measured proliferation after 3 days. Aliquots of these stimulated cells (lymphoblasts) were re-exposed to the mitogen for a further 3 days. In
control experiments, 3 days after mitogen treatment the total nucleic
acid content increased by 3.1- and 3.3-fold for naive and re-stimulated
cells, respectively. Neither AgTx-2 nor clotrimazole alone
significantly inhibited proliferation of naive cells, whereas the
combined drugs were effective, inhibiting by 36.5% (Fig.
3). Consistent with our prediction,
proliferation of activated lymphoblasts was inhibited to a much greater
degree by blocking IK. That is clotrimazole (250 nM, ~6
Kd) inhibited by 65.0%, whereas AgTx-2 (5 nM, ~25 Kd) inhibited by 18.4%. The
two drugs were essentially additive, inhibiting by 86.8%. In parallel
experiments using trypan blue exclusion, we determined that cell
viability at the end of the 3-day proliferation period was not reduced
by the highest concentrations of channel blockers used but remained at
>98%. Thus, reduced nucleic acid content reflects reduced
proliferation and not cell death.

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Fig. 3.
Comparison of the ability of K+
channel blockers to inhibit proliferation of naive T cells
(A) and previously activated T lymphoblasts
(B). The term, naive, is used to indicate cells
that were stimulated from an initial resting state. Each well of a
96-well plate was seeded with 2 × 104 resting cells
or lymphoblasts and incubated with or without channel blockers for 10 min (5 nM agitoxin-2, AgTx-2; 250 nM
clotrimazole, CLT). Then PHA-P (7 µg/ml) was added to
initiate or restimulate proliferation. After 72 h, the CyQUANT
assay was used to measure a change in fluorescence that is proportional
to the change in cell number (see "Experimental Procedures"). Data
are expressed as mean ± S.D. of four independent experiments
(four replicates each). A Bonferroni multiple comparison test was used
to assess each combination of treatments. Values that differ
significantly from controls are indicated (***, p < 0.001), as are significant differences between drug treatments (  ,
p < 0.001).
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Regulatory Volume Decrease (RVD) Is Inhibited by K+
Channel Blockers--
When T lymphocytes are swollen by exposure to
hypotonic solution, they undergo a regulatory volume decrease (RVD) by
loss of K+ and Cl
through ion channels,
followed by osmotically obliged water loss (see "Discussion"). To
assess the relative contributions of IK and Kv1.3 to RVD, we exposed
cells to a standard hypotonic shock (56% of normal osmolarity) in the
presence or absence of 5 nM AgTx-2, 250 nM
clotrimazole, or both drugs. There were no changes in cell volume (with
or without blockers) measured for up to 15 min in isotonic solutions.
Following hypotonic shock, maximal swelling for each blocker was
expressed as a percent increase over the control volume in the same
blocker. Both resting and activated T cells swelled to a maximal volume
within 2 min and then recovered to varying degrees depending on which
K+ channels were blocked. Blocking IK (clotrimazole), Kv1.3
(AgTx-2), or both channels increased the maximal swelling of resting T
cells (Fig. 4A). For
lymphoblasts, blocking IK was as effective as blocking both channels,
whereas blocking Kv1.3 alone did not significantly increase the maximal
volume. Hence, in lymphoblasts, IK appears to play a greater role than
Kv1.3 during the initial swelling phase.

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Fig. 4.
Inhibition of the RVD by K+
channel blockers in resting T cells compared with activated
lymphoblasts. Flow cytometric analysis of right angle light
scattering was used to measure cell swelling (see "Experimental
Procedures"). A, maximal swelling, which occurred within 2 min after the hypotonic shock, is expressed as the percent increase
above the control value. Control values for each drug treatment were
measured in isotonic solution containing the drug, that is 5 nM AgTx-2 to block Kv1.3, 250 nM clotrimazole
to block IK, or both drugs. Open bars, resting T cells;
hatched bars, activated T lymphoblasts 3 days after
mitogenic stimulation. B, the percent recovery at 6 min
after hypotonic shock. Data are presented as mean ± S.D. from at
least 4 experiments. Bonferroni multiple comparison tests were used to
assess all differences. Significant differences between control and
drug-treated cells are indicated for resting T cells (*,
p < 0.05; **, p < 0.01; ***,
p < 0.001) or lymphoblasts ( , p < 0.05;  , p < 0.01;   , p < 0.001).
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We determined the effects of channel blockers on RVD within the first 6 min after the hypotonic shock (Fig. 4B). By this time, in
control hypotonic solution both cell types had almost fully recovered
from swelling: by 81.1 ± 6.8% (n = 6) in resting
cells and 88.5 ± 5.5% (n = 6) in lymphoblasts.
K+ channel blockers attenuated this recovery. Blocking
Kv1.3 reduced volume recovery to 62.3 ± 9.0% (n = 4) in resting cells and 80.9 ± 5.9% (n = 4) in
activated lymphoblasts, a significantly greater effect in resting
cells. IK block was more effective than Kv1.3 block for both cell
types: recovery was 39.6 ± 6.5% (n = 5) in resting cells and 28.8 ± 5.3% (n = 6) in
lymphoblasts, a significantly greater effect in lymphoblasts. When both
channels were blocked, RVD was further decreased in resting cells
(30.7 ± 6.1% recovery, n = 5) but not in
lymphoblasts (29.8 ± 6.8% recovery, n = 5). Our
data show that both K+ channels contribute to RVD; IK is
generally more important and, as predicted from its up-regulated
expression, IK plays a greater role than Kv1.3 in lymphoblasts.
Calmodulin Antagonists Inhibit Native IK and Expressed hSK4
Currents--
Three structurally unrelated drugs were used to inhibit
the Ca2+-calmodulin complex: trifluoperazine (TFP,
Kd ~1 µM (29)), W-7
(Kd ~12 µM (30)), and calmidazolium
(Kd ~50 nM (31)). None of the drugs
were toxic at the concentrations used; i.e. the cells did
not become leaky even after a 30-60-min preincubation. Representative
current traces are shown in Fig. 5,
A
C, and the voltage dependence is summarized in Fig.
5D by plotting the current amplitude at the end of each step
as a function of membrane potential. For hSK4 current in stably
transfected CHO cells, TFP and W-7 effects were steeply
voltage-dependent, with increased current at negative
potentials (36% for TFP and 11% for W-7 at
100 mV). The inhibition
at positive potentials (75% for TFP and 71% for W-7 at +40 mV) showed
a pronounced time dependence (Fig. 5, A and B).
Calmidazolium was not voltage-dependent; i.e.
500 nM inhibited hSK4 by ~35% at all voltages. For
native IK current we were particularly interested in drug effects at negative membrane potentials typical of a non-excitable lymphocyte. In
contrast to hSK4, the lymphoblast IK current was significantly decreased at negative potentials by all three calmodulin antagonists. IK inhibition by 10 µM TFP was mildly
voltage-dependent (a 55% decrease at
100 mV
versus 75% at +40 mV), and this was due to a
time-dependent reduction at depolarized potentials (Fig.
5A). Effects of 25 µM W-7 were qualitatively
similar, with 42% inhibition at
100 mV and 65% at +40 mV. Again,
inhibition by calmidazolium was not voltage-dependent and
500 nM inhibited IK by ~70% (Fig. 5, C and
D). Thus, calmidazolium more effectively inhibited native IK
than hSK4, and at negative potentials TFP and W-7 were only effective
in reducing the native IK. This suggests two mechanisms of action, a
physiologically relevant reduction of native IK at negative potentials
and a time- and voltage-dependent reduction of both IK and
expressed hSK4 at positive potentials.

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Fig. 5.
Calmodulin antagonists reduce hSK4 stably
expressed in CHO cells and native IK current in activated
lymphoblasts. A-C, representative currents in control
saline or with 10 µM trifluoperazine (TFP), 25 µM W-7, or 500 nM calmidazolium added to the
bath. D, average current in CHO cells ( ) and lymphoblasts
( ) expressed as current at the end of each voltage clamp step as a
fraction of control current (IDrug/Icontrol) at
each membrane potential (mean ± S.E., 2-6 cells). Lymphoblast IK
currents were omitted near the reversal potential (about 86 mV) when
they were too small to construct accurate ratios. Some error
bars are smaller than the symbol. All values
marked * are significantly lower for IK than respective values for hSK4
currents (p < 0.05).
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To investigate further IK inhibition by CaM antagonists in
lymphoblasts, we used voltage ramps or steps (see Figs. 1D
and 2) and measured IK at a sufficiently negative potential to elicit large IK currents without contamination by Kv1.3. The current amplitude
was measured at
120 mV from each cell before (control) and after
adding a calmodulin antagonist to the bath. At the end of each
experiment the anion current was recorded (after blocking IK and Kv1.3
with 20 nM ChTx) and subtracted from each total current at
120 mV to calculate the IK amplitude. Both ramp and step protocols yielded the same results. As summarized in Fig.
6, all three CaM antagonists
dose-dependently inhibited IK (n = 4-6
cells unless otherwise indicated). When bath-applied during a
recording, TFP reduced IK by 61.8 ± 11.9% at 5 µM
(p < 0.025) and by 46.1 ± 2.0% at 10 µM (p < 0.03) compared with control
currents in the same cells. W-7 inhibited IK by 70.1 ± 8.5%
(p < 0.05) at 5 µM, by 84.2 ± 5.2% at 10 µM (p < 0.01, n = 10), and by 96.6 ± 3.6% (p < 0.01) at 25 µM.

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Fig. 6.
Inhibition of lymphoblast IK by calmodulin
antagonists. IK amplitude at 120 mV was calculated from
whole-cell currents in response to voltage ramps as explained in the
text. The bars show mean ± S.E. for 4-6 cells unless
otherwise indicated under "Results." A, after the onset
of each recording, trifluoperazine was added to the bath in increasing
concentrations (1, 2.5, and 5 µM) at 10-min intervals. A
separate set of cells was used to control for current rundown by adding
10 µM trifluoperazine to the bath immediately after the
recording had stabilized. Each remaining steady-state current was
calculated as a percent of the control value for that cell (* indicates
a significant reduction of IK, see text). B, increasing
doses of W-7 were added to the bath with each cell serving as its own
control (*, significant reduction of IK). In a separate series of
cells, 50 µM CaM was present in the pipette, and then 10 µM W-7 was applied to the bath 15-20 min after a
whole-cell recording was established. Intracellular CaM significantly
abrogated the effect of W-7 ( ). C, after measuring control
currents, increasing concentrations of calmidazolium were added to the
bath at 10-min intervals (2nd to 4th bars). In
separate experiments cells were preincubated with 500 nM
calmidazolium for 10-15 min before recording for 10-15 min in the
continued presence of the drug. indicates significantly greater
inhibition following preincubation.
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We used a competition experiment in which excess CaM (50 µM with 1.1 µM free Ca2+, see
"Experimental Procedures") was added to the pipette solution, followed by 10 µM W-7 addition to the bath, in the
expectation that CaM would bind to internal W-7 and relieve inhibition.
CaM addition did not significantly increase IK compared with the
current in the same cells during the first 2 min of recording; however, the variability between cells was high (IK increased by 47 ± 26%, p > 0.34, n = 6). Nevertheless,
as expected, excess CaM reduced the W-7-induced IK inhibition (from
84.2 ± 5.2% to 11.3 ± 29.1%, n = 12, p < 0.05), consistent with CaM-antagonist competition at an intracellular site. This result also rules out drug effects at
external sites on the channel at negative potentials (see
"Discussion").
When calmidazolium was bath-applied after a recording was begun, it
dose-dependently inhibited native IK current within 5-10 min, by 54.2 ± 4.5% at 100 nM (p < 0.05) and by 69.0 ± 3.1% (p < 0.01) at 500 nM. Inhibition was more effective when lymphoblasts were
preincubated for 10-15 min before recording (94.1 ± 5.8% inhibition, p < 0.01). In contrast, a 1-2-h
preincubation with 500 nM calmidazolium did increase the
inhibition of hSK4 in CHO cells (34%, n = 8). Thus,
slow drug permeation, which is a potential limiting factor, does not
explain the difference in sensitivities between native IK and expressed
hSK4 currents.
Calmodulin Binds to a Proximal Portion of the C Terminus of
hSK4--
One possibility is that CaM antagonists affect this
KCa current by interfering with interactions between CaM
and the channel protein. If so, by analogy with properties of CaM
binding to CaMK, one might expect CaM-hSK4 channel binding to be
Ca2+-dependent and competitively inhibited by
CaM antagonists. We tagged hSK4 with the Flag epitope and expressed
this construct stably in CHO cells and then determined whether the
expressed protein bound to CaM-conjugated agarose beads. This assay
favors detection of proteins that reversibly bind to CaM. That is if hSK4 channels had already bound to CaM in situ (as reported
for brain SK channels (15)), they would be unable to interact with CaM
on the agarose beads. The membrane proteins were then incubated with
CaM-agarose beads in the presence or absence of Ca2+ or the
CaM antagonist, calmidazolium.
After pre-clearing the samples and washing the beads to remove
nonspecific binding, Ponceau-stained Western blots indicated that
several proteins bound to the beads (data not shown). However, in
samples containing the expressed full-length Flag-tagged hSK4 protein,
only a single band was labeled by the anti-Flag antibody. Fig.
7A shows that binding of the
wild-type full-length hSK4 protein was
Ca2+-dependent (i.e. greatly reduced
by EGTA) but not competed by 1,000 nM of the CaM
antagonist, calmidazolium. To map the region of the channel that binds
to CaM, we made several constructs by deleting the following: the
distal C terminus (leucine zipper region, which we call "N-M-Ct1"
since it contains the N terminus, membrane-spanning, and C terminus 1 regions), the entire N terminus and transmembrane domains (Ct1-Ct2),
all but the proximal C terminus (Ct1), or the proximal C-terminal tail
(N-M-Ct2). All but one channel construct bound to CaM in a
Ca2+-dependent manner that was not competitive
with calmidazolium, i.e. the construct lacking the proximal
C terminus (Ct1) did not detectably bind to CaM.

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Fig. 7.
Calmodulin binds to the proximal portion of
the C terminus of expressed hSK4; this region is important in channel
function. A, left panel, channel constructs
were made to delete the following: the distal C-terminal leucine-zipper
region (called N-M-Ct1), entire N terminus, and
transmembrane domains (Ct1-Ct2), all but the proximal C
terminus region (Ct1), and just the proximal C-terminal tail
(N-M-Ct2). Right panel, Western blot analysis of
binding of Flag-tagged hSK4 to CaM-agarose beads. Left lane,
in the presence of 0.5 mM CaCl2; middle
lane, calcium was chelated with 3 mM EGTA; right
lane, in the presence of the CaM antagonist, 1000 nM
calmidazolium with 0.5 mM CaCl2. All lanes
contained ~20 µg of protein. B, membrane potentials (in
mV) of CHO cells expressing various hSK4 constructs. Whole-cell
recordings were made in the current clamp mode with high (~1.1
µM) intracellular Ca2+. Data are expressed as
mean ± S.D with the number of cells in parentheses.
Significant differences between hSK4.Flag (i.e. wild type,
WT) and N-M-Ct1 or N-M-Ct-2 are indicated (***,
p < 0.001).
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C-terminal Deletions Have Functional Consequences--
We used the
membrane potential (Vm) to monitor the expression of
functional hSK4 channels of various constructs in transfected CHO cells
(Fig. 7B). Whole-cell patch clamp recordings were made with
1 µM free Ca2+ in the pipette (as for all
recordings of currents), and the membrane potential was recorded in the
current clamp mode. The Vm of CHO cells transfected
with wild-type hSK4 channels was
67 ± 4 mV (n = 5) indicating a significant K+ permeability (Nernst
potential for K+,
82 mV), and hSK4 with a C-terminal Flag
tag produced the same Vm (
68 ± 4 mV,
n = 5). The Vm of control cells transfected with vector alone (pcDNA3) was
10 ± 8 mV,
indicating a very low background K+ permeability. The hSK4
construct lacking the CaM-binding region (N-M-Ct2) did not produce a
negative membrane potential despite the high intracellular
Ca2+; Vm was significantly less negative
than for the wild-type construct (
2 ± 1 mV, n = 5, p < 0.001). Interestingly, Vm was also close to 0 (
5 ± 2 mV, n = 6) for the
construct lacking the distal C terminus (N-M-Ct1).
The Lymphoblast IK Current Is Reduced by a CaM Kinase
Antagonist--
KN-62 is a membrane-permeant CaM kinase antagonist,
whereas KN-04 is an inactive analogue used as a negative control. KN-62 reduced IK in a time- and voltage-independent manner (Fig.
8A). Fig. 8B
summarizes effects of KN-62 alone or in combination with the CaM
antagonist, W-7. IK was measured in lymphoblasts at
120 mV after
subtracting the small anion current (as for Fig. 6). We first tested
intact cells at 37 °C in an attempt to maintain kinase and
phosphatase activity and turnover of protein phosphorylation. Preincubation with 10 µM KN-62 alone (30 min, 37 °C)
decreased the current by 55.5 ± 11.2% (p < 0.02, n = 4), whereas KN-04 had no effect (97 ± 11% of the control value, n = 4, p > 0.8). After KN-62 preincubation and with the drug present throughout
whole-cell recordings, adding W-7 to the bath further reduced the
current, i.e. by 80.2 ± 10.1% reduction
(n = 4, p < 0.05) with the combined drugs. Acute effects of bath-applied KN-62 were tested during whole-cell recordings at ~37 °C. IK was substantially reduced by
KN-62 (by 67.9 ± 11.7%; n = 4, p < 0.02) but not by KN-04. Under drug-free conditions, IK in
lymphoblasts was not obviously temperature-dependent; the
specific conductance was 606 ± 210 pS/picofarads at room
temperature and 552 ± 13 pS/picofarads when the same cells were
warmed to >33 °C (n = 6, p > 0.7).
Interestingly, the variability in current amplitude was greatly reduced
at the higher temperature. Unlike the native channels in lymphoblasts, hSK4 stably expressed in CHO cells was not inhibited by 10 µM KN-62 and remained at 94.3 ± 2.5%
(n = 6, p > 0.2) of the control value
measured with solvent alone (1% Me2SO).

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Fig. 8.
Native IK current in T cells is inhibited by
the CaM kinase inhibitor, KN-62. A, representative
current traces during 800-ms voltage clamp steps between 120 and +20
mV before and 10 min after adding 10 µM KN-62. All
recordings were at 37 °C and included 5 nM MgTx to block
Kv1.3. B, average current amplitudes (±S.E.,
n = 4-5 cells from 2 to 3 batches) calculated at 120
mV (as in Fig. 4) as a percent of control values. For each batch of
cells, 1 aliquot was used to measure control IK currents at room
temperature. A 2nd aliquot was preincubated with KN-62 (10 µM, 30 min at 37 °C), and IK was measured at room
temperature 10-15 min after beginning a recording. For some cells from
the same KN-62-preincubated aliquots, 10 µM W-7 was added
15-20 min into a recording. Values that differ from control (room
temperature) currents are indicated (*, p < 0.05; **,
p < 0.01). A 3rd aliquot of cells was used at 37 °C
for control recordings (> 10 min), followed by KN-62 or KN-04 addition
for 10-20 min. A significant inhibition was seen with KN-62 only ( ,
p < 0.05). All control cells were treated with 0.5%
Me2SO, the maximum solvent concentration used for KN-62 and
KN-04.
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DISCUSSION |
Comparison of the Cloned hSK4 with IK Current in Activated T
Cells
The present results are entirely consistent with the IK current in
T lymphocytes being the product of the hSK4/hIK1/hKCa4 gene,
which was recently cloned from cDNA libraries from human placenta
(7), pancreas (8), and lymph node (9). Since the product we cloned is
100% identical to hSK4/hIK1/hKCa4, differences in
properties of the native lymphocyte IK and exogenously expressed hSK4
channels are not expected unless such properties are determined by
something other than the
subunit of the channel. In principle, differences could arise if the channel forms heteromultimers with another protein, if alternative splice variants exist, or if the channel interacts with accessory molecules. It is intriguing that multiple transcript sizes are commonly seen for this channel, i.e. 2.6 and 3.8 kb (7), ~2.1 kb, and at least one larger
band (8), 2.2 kb, with two larger bands (9), and a prominent 2.2-kb
band with a weaker 2.6-kb band (present study, data not shown).
Biophysical properties of the lymphocyte IK current have been described
at the single channel (2, 5, 24) and whole-cell level (5). Whereas
channel gating is independent of voltage, it is highly sensitive to
intracellular free Ca2+, activating at <200 nM
in T and B cells, reaching half-maximal activation at about 450 nM, and maximal activation at ~1 µM (2, 5).
Both the single channel and whole-cell current versus
voltage (I-V) relations are inwardly rectifying with symmetrical
K+ concentrations on both sides of the membrane (5, 6, 24). The single channel I-V relation is linear under physiological Na+/K+ gradients, which, together with the
voltage-independent gating, results in a whole-cell current that is
nearly linear (5, 24). Expressed hSK4/hIK1/hKCa4 currents (Refs. 7-9
and present study) have the following features in common with the
lymphocyte IK current (Refs. 2, 3, 5, 7, and 24 and present study);
activation by sub-micromolar free Ca2+, time- and
voltage-independent gating, inwardly rectified single channel I-V
relations in symmetrical K+ (10-35 pS), and nearly linear
I-Vs in physiological Na+/K+ gradients (~10
pS).
The pharmacological profiles of native IK and hSK4/hIK1/hKCa4 are also
similar. Native IK in lymphoblasts and hSK4 expressed in CHO cells were
blocked by ChTx (IC50 2-10 nM) but very poorly by iberiotoxin (IC50 >200 nM), margatoxin
(IC50 >100 nM), or tetraethylammonium (IC50 30-40 mM). Clotrimazole showed a similar
potency for blocking IK in lymphoblasts (present study) and for
heterologously expressed hSK4 (IC50 25-60 nM
(Ref. 8 and present study)). The hSK4/hIK1/hKCa4 channel is expected to
be insensitive to both apamin and d-tubocurarine since it
lacks two necessary amino acids in the putative pore (27) and, as
expected, neither the lymphoblast IK nor the expressed hSK4 current
were significantly inhibited by apamin (IC50 >100 nM (Refs. 5 and 7-9 and present study) or
d-tubocurarine (IC50 >250 µM;
present study).
Increased Role for hSK4 in Lymphoblast Proliferation
K+ channel activity is important during the early
activation phase of naive T cells, especially for maintaining a
hyperpolarized membrane potential, promoting a rise in intracellular
Ca2+, and permitting a cascade of events that culminates in
interleukin-2 production (1, 32, 33). In the first few hours after
mitogenic stimulation, precisely when Ca2+ elevation is
necessary (34, 35), K+ channel blockers, or other means of
depolarizing T cells (high external K+, voltage clamp),
inhibit T cell activation (36) by compromising Ca2+ influx
and the resulting rise in Ca2+. Early studies using
non-selective K+ channel blockers (e.g.
quinidine, 4-aminopyridine) were later substantiated by more selective
peptide toxins including charybdotoxin, which blocks both IK
(Kd ~2-6 nM (Refs. 3 and 5 and
present study)) and Kv1.3 channels (Kd ~1
nM (1, 5, 16, 17, 24)), and margatoxin or noxiustoxin which block Kv1.3 but not IK channels (1, 16, 17, 28, 37). From the limited
functional studies using blockers that discriminate between Kv1.3 and
other K+ channels, Kv1.3 appears to be important for
activation of naive T cells through pathways that are
Ca2+-dependent (1, 32).
The contribution of KCa channels to T cell activation and
proliferation is still poorly understood. Since we previously found that mitogens activate KCa channels in cell-attached
patches from naive human T cells, we proposed that they also play a
role in T cell activation (14). In the present study we observed a
14.6-fold increase in hSK4 transcripts by 3-4 days after mitogenic
stimulation. During the same period, mRNA levels for Kv1.3
increased only 1.3-fold. These changes are consistent with previous
patch clamp studies showing an approximate doubling in Kv1.3 current
(see Ref. 1), a 30-fold increase in the ChTx-sensitive KCa
current (5) and an increase in hKCa4 mRNA (9). The prediction that
IK will be increasingly important for the secondary immune response
(e.g. proliferation of previously activated lymphoblasts) is
supported by the present results. There is also a recent report that
Ca2+ signaling and proliferation were more strongly
inhibited by ChTx in lymphoblasts than in naive T cells (38); however,
ChTx does not discriminate between IK and Kv1.3 channels. To separate
better the contributions of Kv1.3 and IK to T cell function, we used AgTx-2 to block Kv1.3 and clotrimazole to block IK. Consistent with our
expectations, IK block more effectively inhibited proliferation of
lymphoblasts than naive T cells. Furthermore, despite the greater Kv1.3
channel block (AgTx-2 at ~25 Kd) than IK block (clotrimazole at ~6 Kd), IK block was more
effective in inhibiting lymphoblast proliferation, i.e. by
65.0% compared with 18.4% for Kv1.3 block. Blocking both channels
(AgTx-2 + clotrimazole) was approximately additive, reducing
lymphoblast proliferation by 86.8%. Proliferation of naive T cells was
also sensitive to blocking both channels (36.5% inhibition),
consistent with previous reports of reduced proliferation when IK + Kv1.3 were blocked with ChTx (1, 32, 33).
How might both Kv1.3 and IK channels contribute to T cell
proliferation? Within seconds after stimulating the T cell receptor, tyrosine-kinase mediated activation of phospholipase C produces inositol 1,4,5-trisphosphate and quickly triggers Ca2+
release from internal stores. A plasma membrane channel (the Ca2+ release-activated Ca2+ channel) then opens
to allow Ca2+ influx, which is required for several hours.
Ca2+ release-activated Ca2+ channel opening is
not voltage-dependent, but Ca2+ influx is
strongly driven by the membrane potential. Thus, any means of
increasing the K+ conductance and hyperpolarizing the cell
will facilitate Ca2+ entry. Kv1.3 is voltage-gated,
activated by depolarization, and its steady-state activity is maximal
between
50 and
30 mV in resting T cells, depending on
post-translational modulation (37). Hence it is likely to play a role
only when the membrane is moderately depolarized. In contrast, gating
of the IK/hSK4 channel is voltage-independent but exquisitely sensitive
to internal Ca2+; thus, it is well designed to open
whenever Ca2+ rises marginally above the resting level.
Rather than Ca2+ changing in a sustained manner after T
cell receptor stimulation, Ca2+ and membrane potential can
oscillate (1, 38, 39). Collectively, these complementary properties
would allow the cell alternately to use IK channels when
Ca2+ is high, even if the membrane is hyperpolarized, and
Kv1.3 channels during periods of low Ca2+ and/or depolarization.
Role of hSK4 and Kv1.3 in Volume Regulation
Volume regulation in leukocytes and other mammalian cells has been
extensively reviewed (10-12). The RVD involves ion efflux through
separate K+ and anion channels. Its rate and extent depend
on the combined ion conductances; thus if either the K+ or
Cl
current is small (or blocked pharmacologically) it
will limit volume recovery. Identifying the particular K+
channel(s) that underlie RVD has been problematic, largely due to the
lack of selective blockers and uncertainty over whether swelling evokes
a rise in intracellular Ca2+. For instance, early evidence
of a role for Kv1.3 was not convincing since it first relied on
nonspecific K+ channel blockers and then on ChTx (10, 11,
40) which we now know blocks both Kv1.3 and IK. Kv1.3 can confer RVD
when transfected into a mouse T cell line that lacks voltage-gated
K+ currents (13); however, that study did not address the
presence or role of KCa channels. To determine whether
intracellular Ca2+ is elevated in human T cells during RVD,
we previously designed a perfused cuvette system for fluorometric
measurements (41). We found that hypotonic shock elicits a rapid,
biphasic rise in Ca2+ (14) which comprises release from
internal stores and influx across the plasma membrane. Ca2+
reached a peak in 1-2 min and remained elevated for at least 10 min,
and the entire pattern was indistinguishable from Ca2+
signaling during T cell activation. Thus, we proposed (14) that both
KCa and Kv1.3 channels will contribute to RVD, with Ca2+ and voltage oscillations alternately opening each type
of K+ channel.
Previous studies of RVD in T cells have been restricted to resting
cells and show a stereotypical response (10, 11, 13); within 1-2 min
after a hypotonic shock, T cells swelled to ~120% of their original
volume and then returned to their original volume within 5-15 min. Our
results, using right angle light scattering to measure RVD, are in
excellent agreement both in the extent of swelling (~120%) and in
the time course, with maximal swelling within 1-2 min and nearly full
recovery within 6 min for both resting T cells and lymphoblasts. We
have now assessed the relative contributions of IK and Kv1.3 channels
to RVD in resting T cells compared with activated lymphoblasts. Owing
to the dramatic increase in IK and small increase in Kv1.3 in
lymphoblasts, we expected RVD to depend more on IK than Kv1.3 channels
in lymphoblasts.
We examined the role of each channel type in the initial swelling to
maximal volume and in the degree of recovery by 6 min after a 56%
hypotonic shock. For resting T cells, blocking Kv1.3 or IK, or both
channels simultaneously, increased the maximal volume, implying that
K+ efflux through both channels occurs even during the
initial swelling phase. Consistent with this conclusion, in previous
studies T cells swelled less than predicted for a passive osmometer
(10-12). We had anticipated a role for IK channels in resting cells,
since IK is expressed (Refs. 6 and 39 and present study), and
intracellular Ca2+ rises after a hypotonic shock (14). For
lymphoblasts, IK block was very effective in increasing the maximal
volume, whereas Kv1.3 block had no effect. Thus, in lymphoblasts volume
regulation also proceeds during the swelling phase but IK plays a much
greater role than Kv1.3 current at this time.
The extent of recovery after the maximal volume is reached reflects
both the K+ and Cl
conductances during the
RVD phase. Substantial recovery occurred within 6 min in both cell
types, and the slightly greater recovery in lymphoblasts is consistent
with up-regulation of IK/hSK4 expression. RVD was significantly
inhibited by blocking Kv1.3 channels in both resting T cells and
lymphoblasts but was more effective in resting cells. For both cell
types IK block was more effective than Kv1.3 block; moreover,
inhibition of RVD was greater in lymphoblasts. Thus, both
K+ channels contribute to RVD, but their relative
importance is opposite as follows: Kv1.3 plays a greater role in
resting cells, and IK is more important in lymphoblasts. Not only are
these results predicted from the up-regulated expression of IK/hSK4 in
lymphoblasts but IK activation implies that hypotonic shock elicits an
early and sustained rise in Ca2+ in activated lymphoblasts,
as we have previously shown for resting T cells (14).
Although there is insufficient information to calculate K+
fluxes through Kv1.3 and IK channels during RVD, some predictions can
be made by considering their expression and biophysical properties. Flux through each channel type is proportional to the number of channels (n), their open probability
(Po), their single channel conductance (
),
and the driving force, which is the same at a given voltage. For
resting T cells, the number of Kv1.3 channels per cell is much larger
than IK/hSK4 channels, and
is similar (~10 pS in a normal Na/K
gradient). So, for IK channels to play a substantial role in RVD, their
Po must be much larger than the Po of Kv1.3 channels in resting cells. Kv1.3
contribution will be controlled by the membrane potential since these
channels require moderate depolarization to be tonically active (37,
42, 43). A simple model is that Ca2+ remains elevated
thereby activating IK, and the membrane potential remains
hyperpolarized, thereby reducing the opening of Kv1.3 channels. IK/hSK4
expression is much higher in lymphoblasts than in resting T cells, and
since IK gating is voltage-independent, this channel is expected to
contribute more whether or not the membrane potential fluctuates,
provided Ca2+ remains modestly elevated.
Calmodulin-dependent Modulation, Evidence for More Than
One Mechanism
Our electrophysiological results implicate calmodulin (CaM) in
regulating native IK channels in lymphoblasts. In principle, CaM
antagonists could act by interfering with interactions between CaM and
the channel protein, by interacting with accessory CaM-binding molecules (e.g. CaM kinases or channel
subunits, if they
exist), or by directly interfering with the channel protein. Some
predictions can be made from the mechanism by which the antagonists
inhibit CaM (44-46). In cell-free systems, CaM changes conformation
when at least two of its four Ca2+-binding domains are
saturated (Kd ~2.4 µM
Ca2+). CaM antagonists can then bind reversibly to a newly
exposed hydrophobic site (47) thereby preventing interactions between CaM and target proteins. Thus, it is expected that excess CaM will
competitively reduce inhibition by titrating the amount of drug
available for inhibition.
Direct interactions of some CaM antagonists in the pore of some
K+ channels have been proposed when their potency for CaM
inhibition differed from that of channel inhibition (48-50), or the
drugs were effective even when Ca2+ was not elevated (50),
or exogenously added CaM did not compete with the antagonists (49). We
found that trifluoperazine and W-7 produced time- and
voltage-dependent decreases in both native IK and expressed
hSK4 current at positive potentials, which may reflect a direct drug
interaction with the channel protein. Of greater physiological
relevance is the inhibition of native IK channels we observed at
negative membrane potentials. In this case the potency of inhibition by
W-7 and TFP was consistent with effects on CaM, and as expected for
competitive drug binding, excess internal CaM significantly relieved
the inhibition by W-7. This result also rules out significant channel
block by W-7 from the outside at negative potentials. CaM antagonists
may affect the lymphoblast IK current through interactions between CaM
or other CaM-binding molecules and the channel protein. Such
interactions must differ for hSK4 channels stably expressed in CHO
cells since, at negative potentials, these currents were not inhibited
by TFP or W-7, and calmidazolium was less effective than on IK
currents. As discussed below, differences in actions on native IK and
hSK4 channels may reflect multiple sites of action.
Direct Interactions between CaM and IK/hSK4 Channels--
Despite
the exquisite Ca2+ sensitivity of IK/hSK4 gating, the
primary amino acid sequence of the
subunit contains no known Ca2+-binding sites, that is no E-F hands, C2 domains (51),
or Ca2+ "bowls" (52). We found that channels made from
wild-type, full-length hSK4
subunits bind to calmodulin. Although
binding was greatly inhibited when Ca2+ was chelated, some
binding remained. Deletion mutants of several cytoplasmic regions that
are relatively conserved between hSK4 and brain SK channels showed that
CaM binding was restricted to the proximal C-terminal tail of hSK4 (a
region we call "Ct1," see Fig. 7). Deleting the Ct1 region
prevented the expression of functional Ca2+-gated hSK4
channels. When wild-type hSK4 was expressed and whole-cell membrane
potential (Vm) recordings were made with micromolar intracellular Ca2+ to maximally activate hSK4,
Vm became highly negative owing to the
hyperpolarizing K+ conductance. In contrast, the
Ct1-deleted channel failed to produce a hyperpolarizing K+
conductance, and Vm remained essentially at zero.
There are two possibilities as follows: without CaM binding the
channels did not open in response to high Ca2+ (as is the
case for SK2 channels (15)) or the mutant channels did not assemble
properly in the cell membrane. In the future it will be useful to
examine the assembly and trafficking of mutant hSK4 channels,
particularly since the Ct2-deletion mutant (lacking the leucine zipper
region) also failed to produce a hyperpolarizing K+
conductance. The
subunits of brain SK channels (SK1-3) also bind
to CaM in the proximal part of the cytoplasmic C terminus, and CaM
apparently serves as the Ca2+-binding gate (15). Most of
the C terminus of SK2 channels (4
helices, A-D) bound to CaM,
whereas a "post-D" C-terminal tail (corresponding to our Ct2,
leucine-zipper region) did not. If helices A-D were all present,
binding was independent of Ca2+, whereas helices B and C
and B-D conferred Ca2+-dependent binding to
CaM.
Our results on hSK4 share several features with SK2 (15), but they also
differ in ways that are consistent with additional sites of interaction
or modulation of the native channel in lymphocytes. Although brain SK
channels have only modest overall homology (~40%) to hSK4 (7), some
regions are more homologous. The 95-amino acid CaM-binding domain (Ct1)
that we identified in hSK4 is the same channel region as helices A-C
in SK2. Helix A in SK2 is highly homologous to the corresponding region
of hSK4 (79% identical), whereas regions B and C have much lower
homology (20% identical). For SK2, the Ca2+-independent
CaM binding and patch clamp studies in which calmidazolium failed to
inhibit the expressed channels were taken as evidence that CaM binds
constitutively and irreversibly to brain SK channels (15). We found
that most of the CaM binding to hSK4 was
Ca2+-dependent; however, as explained earlier,
the binding assay we used would not detect channels irreversibly bound
to CaM. Several possible explanations for these differences will
require further study. For instance, there may be more than one
CaM-binding site with different affinities, as is the case for cation
channels in retinal rods (53), with a lower affinity site that is
reversible and Ca2+-dependent, perhaps in Ct1
of hSK4 (helices B-D in SK2). CaM antagonists inhibited the native
current in lymphoblasts, with little or no inhibition of expressed hSK4
currents, a result that is consistent with the failure of 1000 nM calmidazolium to prevent CaM binding to the expressed
hSK4 protein. A further possibility is that weaker CaM channel binding
in lymphoblasts, perhaps a result of other protein-channel interactions
(see below), allows more effective competition by CaM antagonists.
Evidence for Accessory Molecules in Lymphocytes--
The striking
differences in inhibition at negative potentials of native IK
versus expressed hSK4 channels provide the first evidence
that accessory molecules (other than CaM) modulate a member of the SK
channel family. Candidate molecules include CaMK, calcineurin, and
subunits analogous to those interacting with voltage-gated
K+ channels. Although
subunits have not been identified
for SK channels, there is evidence that apamin-sensitive SK channels form hetero-oligomers. In a variety of cells expressing SK channels, apamin binds to both high (59 or 86 kDa) and low (30 or 33 kDa) molecular mass polypeptides that are integral membrane proteins (54).
It is unlikely that such accessory molecules are essential for channel
activity since all known members of the SK family, with the exception
of rSK1 (7), are functional in expression systems, including
Xenopus oocytes, HEK, and CHO cells. This observation also
implies that the Ca2+-binding site, which is thought to be
CaM bound to the channel (15), functions normally in these cells.
Since the CaM kinase antagonist, KN-62, inhibited native IK current but
had no effect on hSK4, it is necessary to consider how CaM kinase might
selectively modulate the current in lymphocytes. hSK4 contains a
potential phosphorylation site in the C-terminal domain that should
accommodate either CaM kinase II (or protein kinase A) (45); however,
this site is not necessarily phosphorylated. It may be that CHO cells
have insufficient CaM kinase (T cells express high levels of CaM kinase
II and IV (55)) or that the site of phosphorylation is not on the
channel protein itself, but rather on a
subunit or other unknown
accessory molecule. Interestingly, in lymphoblasts the increased
inhibition by W-7 in the presence of KN-62 is consistent with dual
modulation by CaM binding and CaM kinase.
Potential modulation of lymphocyte IK/hSK4 channels by CaM and CaM
kinase is of broader importance. Early in T cell activation or
lymphoblast re-activation, there is a rise in intracellular Ca2+ that activates CaM-dependent enzymes.
These include CaM kinases II and IV (55) and calcineurin (protein
phosphatase 2B), which is highly expressed in lymphocytes and crucial
for T cell proliferation (34). CaM antagonists can inhibit some
lymphocyte functions that either trigger conductive K+
fluxes or are sensitive to the membrane potential of the cell (which
depends on K+ channels). T cell activation (56),
cell-mediated cytotoxicity (57), and volume regulation (40) are
inhibited both by K+ channel blockers and by CaM
antagonists. Our present results provide new evidence that a specific
K+ channel (IK/hSK4) that is important for at least two of
these functions (proliferation and volume regulation) is susceptible to
CaM antagonists, thus providing a link between CaM and K+
channels in regulating lymphocyte function.