Structural Compatibility between the Putative Voltage Sensor of
Voltage-gated K+ Channels and the Prokaryotic KcsA
Channel*
Marco
Caprini
§,
Stefano
Ferroni
,
Rosa
Planells-Cases§,
Joaquín
Rueda¶,
Carmela
Rapisarda
,
Antonio
Ferrer-Montiel§
, and
Mauricio
Montal**
From the
Department of Human and General Physiology,
University of Bologna, Via San Donato 19/2, 40127 Bologna, Italy, the
§ Centro de Biología Molecular y Celular and the
¶ Departamento de Histología, Universidad Miguel
Hernández, 03202 Elche Alicante, Spain, and the ** Department of
Biology, University of California at San Diego,
La Jolla, California 92093
Received for publication, January 18, 2001, and in revised form, March 21, 2001
 |
ABSTRACT |
Sequence similarity among and
electrophysiological studies of known potassium channels, along with
the three-dimensional structure of the Streptomyces
lividans K+ channel (KcsA), support the tenet that
voltage-gated K+ channels (Kv channels) consist of two
distinct modules: the "voltage sensor" module comprising the
N-terminal portion of the channel up to and including the S4
transmembrane segment and the "pore" module encompassing the
C-terminal portion from the S5 transmembrane segment onward. To
substantiate this modular design, we investigated whether the pore
module of Kv channels may be replaced with the pore module of the
prokaryotic KcsA channel. Biochemical and immunocytochemical studies
showed that chimeric channels were expressed on the cell surface of
Xenopus oocytes, demonstrating that they were properly synthesized, glycosylated, folded, assembled, and delivered to the
plasma membrane. Unexpectedly, surface-expressed homomeric chimeras did
not exhibit detectable voltage-dependent channel activity
upon both hyperpolarization and depolarization regardless of the
expression system used. Chimeras were, however, strongly dominant-negative when coexpressed with wild-type Kv channels, as
evidenced by the complete suppression of wild-type channel activity.
Notably, the dominant-negative phenotype correlated well with the
formation of stable, glycosylated, nonfunctional, heteromeric channels.
Collectively, these findings imply a structural compatibility between
the prokaryotic pore module and the eukaryotic voltage sensor domain
that leads to the biogenesis of non-responsive channels. Our results
lend support to the notion that voltage-dependent channel
gating depends on the precise coupling between both protein domains,
probably through a localized interaction surface.
 |
INTRODUCTION |
Ion channels are multisubunit membrane proteins involved in action
potential propagation, neurotransmitter release, and
excitation-contraction coupling in excitable tissues. Protein sequence
information obtained from recombinant DNA technology has revealed that
voltage-gated ion channels form a large superfamily of related proteins
that include Na+, Ca2+, and K+
channels. Voltage-gated K+ channels are involved in a host
of cellular processes, from setting the resting membrane potential and
shaping action potential waveform and frequency to controlling synaptic
strength (1). The first K+ channel cloned from the
Drosophila Shaker locus (2) seemed to code for a unit
similar to one of the four internal repeats of the more complex
Na+ and Ca2+ channels, consisting of six
-helical transmembrane segments (S1-S6) and a pore-forming loop
(P-loop) (3, 4).
The sequence similarity between voltage-gated K+ channels
and voltage-dependent Na+ and Ca2+
channels suggested a modular architecture of the voltage-gated ion
channel family. In this modular context, the N-terminal portion of the
eukaryotic voltage-gated proteins up to and including the S4 segment
may represent a sensor module, responsible for detecting changes in
transmembrane potential (5). This notion appears warranted since work
in several laboratories combining mutagenesis and biophysics has
demonstrated that perturbation of this domain selectively affects
channel gating without altering the permeation properties (6-13). The
S5-P-S6 region of voltage-gated channels may represent a "pore"
module within the larger protein. Indeed, conduction can be abolished
by a pore mutation without affecting channel gating (14).
Examination of the sequence of subsequently cloned potassium channels
from diverse sources also substantiates the existence of modularity
within the voltage-gated potassium channels (15-17). The genes coding
for the inward rectifier class of potassium channels code for a protein
analogous to the carboxyl-terminal portion of the voltage-gated
potassium channels, the S5-P-S6 region (18). These genes are
sufficient to form potassium-selective pores, but with poor intrinsic
voltage sensitivity. Further evidence for the modular organization of
the potassium channels was provided by the identification of a class of
channels consisting of two pore modules, with (19) or without
(20, 21) an attached "sensor" module. More recently, a novel
structural class of mammalian potassium channels has been discovered,
with four transmembrane segments and two pore regions (TWIK-1, TREK-1,
TRAAK, TASK, and TASK-2), and activated by different kind of stimuli
such as membrane stretch and modification of pH (22).
The identification of the KcsA channel from
Streptomyces lividans has shown that this channel has the
closest kinship to the S5-P-S6 region of the Kv channel family.
Moreover, KcsA is most distantly related to eukaryotic inwardly
rectifying channels with two putative predicted transmembrane segments
(23). Single-channel recording, flux measurements, and ligand binding
assays have shown KcsA to be a high-conductance, tetrameric,
K+-selective channel with an externally located receptor
site for charybdotoxin family peptides (24-26). In addition, the
recent crystallization of the KcsA protein has provided a structure for such a pore module, enlightening our knowledge of the molecular basis
of ion permeation (27). The sequence similarity between KcsA and Kv
channels has led to the notion that the prokaryotic channel may be the
bacterial ancestor of the pore module present in eukaryotic channels
(23).
Thus, it appears reasonable that voltage-gated potassium channels are
structurally modular. It has been recently demonstrated, in channel
proteins consisting of the sensor module from mouse Kv1.1
(mKv1.1)1 and the pore module
from fly Shaker with inactivation ball removed channel (Shaker), and vice versa, that these putative modules can
operate outside their native context (28). To further substantiate the
modular design of the Kv channel family, we examined whether it would
be feasible to confer voltage sensitivity to the
voltage-insensitive prokaryotic ancestor KcsA channel by linking its
pore domain to the voltage sensor of eukaryotic Kv channels. Thus, we
constructed chimeric channels by replacing the pore domain of mKv1.1
and Shaker with the pore module of the KcsA channel (23). Chimeric Kv
channels appear to proceed via appropriate biogenesis pathways, as
evidenced by their normal translation, glycosylation, folding, and
delivery to the plasma membrane of the injected oocytes. Heterologously expressed chimeras did not show detectable voltage-gated ionic current,
although they behaved as strong dominant-negative subunits, completely
inhibiting the channel activity of wild-type mKv1.1 and Shaker
channels. Taken together, these results lend support to the notion of
structural compatibility of both protein domains. Our findings also
suggest a higher degree of molecular adaptability for functional
coupling of both protein modules.
 |
MATERIALS AND METHODS |
Molecular Biology of Chimeric Design
Standard molecular biological techniques were as described (29).
Shaker (30) was a gift of L. Toro (UCLA), and KcsA was from S. Choe
(Salk Institute). Three versions of KcsA-containing chimeras were
designed, and a hemagglutinin (HA) peptide tag was included at the
carboxyl-terminal end of mKv1.1 and all chimeric coding regions for
immunodetection of the expressed protein.
The mKv1.1(S1-S4,5)-KcsA chimera was constructed by replacing the
region of mKv1.1 encompassing the S5 segment to the C-terminal end
(amino acids 321-481) with the corresponding residues of KcsA (amino
acids 27-160). Polymerase chain reaction was used to introduce a
silent ClaI site in mKv1.1 at amino acids 321 and 322. Two
oligonucleotide primers were also used to amplify KcsA from amino acid
27 through the stop codon while simultaneously introducing a
ClaI site at amino acid 27 and a BamHI at amino
acid 160, subsequently used to subclone KcsA into pGEMHE/mKv1.1. Note
that this cloning strategy left a 14-amino acid segment from mKv1.1
(amino acids 482-495) at the C-terminal end of KcsA. The
mKv1.1(S1-S4)-KcsA chimera was constructed by replacing the region
encompassed by the S4-S5 loop up to the C-terminal end of mKv1.1
(amino acids 311-481) with the complete KCSA gene.
Polymerase chain reaction was used to introduce a silent
PstI site in mKv1.1 at amino acids 311 and 312. KcsA was
amplified by polymerase chain reaction, inserting PstI and
BamHI sites at amino acids 1 and 160, respectively.
The amplified fragment was then subcloned into a silent
PstI site and the BamHI site into pGEMHE/mKv1.1.
The Shaker(S1-S4,5)-KcsA chimera was constructed by replacing the
region of Shaker from amino acid 351 onward with KcsA. A silent
ClaI site was introduced in Shaker at amino acids 350 and
351. A fragment of KcsA from the first chimera cut at the
ClaI and EcoRV sites 3' of the stop codon was
subcloned into pBluescript/Shaker between the ClaI site and
a blunted XhoI site 3' of the stop codon. The sequences of the transferred segments were verified by both restriction analysis and
dideoxy sequencing (31). For in vitro transcription,
chimeric and wild-type channel clones were linearized and used as
templates with the mMESSAGE mMACHINE kit (Ambion Inc., Austin, TX). For expression in COS-7 mammalian cells, mKv1.1(S1-S4,5)-KcsA,
Shaker(S1-S4,5)-KcsA, and wild-type mKv1.1 were subcloned into the
mammalian expression vector pCIneo (Promega). Enhanced green
fluorescent protein (GFP; gift of R. Tsien, University of California,
San Diego) was also subcloned into pCIneo.
Cell Culture Methodology
COS-7 cells (gift of M. Canossa, University of Bologna) were
cultured as described (32). The day prior to transfection, COS-7 cells
were replated in 35-mm Petri dishes at a density of 2-5 × 104 cells/dish and maintained in supplemented Dulbecco's
modified Eagle's medium. COS-7 cells were transfected with the
constructs and GFP by the DEAE-dextran method and assayed for
electrophysiological measurements 48-72 h post-transfection (33).
Protein Immunoprecipitation and Immunoblotting
Protein analysis for the different clones was carried out by
Western immunoblotting. Oocytes (8-10/sample) were collected and lysed
as described (34).
Soluble materials from homogenized oocytes were separated on an
SDS-polyacrylamide gel and transferred to a nitrocellulose membrane.
The nitrocellulose was blocked and probed with 4 µg/µl anti-HA
monoclonal antibody (mAb) (Roche Molecular Biochemicals, Mannheim,
Germany), anti-Shaker polyclonal antibody (gift of F. Tejedor, Consejo
Superior de Investigaciones Científicas, Universidad Miguel
Hernández), or anti-mKv1.1 antibody (Alomone), and proteins were detected with an alkaline phosphatase-conjugated antibody (Sigma). The bands were later visualized using the alkaline phosphatase conjugate substrate kit (Bio-Rad). Immunoprecipitation was carried out
with an anti-HA monoclonal antibody (2.5 µg/ml) as described (35).
Samples were then boiled in SDS-polyacrylamide gel electrophoresis loading buffer and electrophoresed as described above.
Immunocytochemical Labeling of Channel Protein
Xenopus oocytes injected with transcripts of
mKv1.1-HA or HA-tagged chimeras or water were selected 48 and 72 h
post-injection, embedded in optimal cutting temperature resin
(ProSciTech, Thuringowa, Australia), and quickly frozen.
12-µm-thick sections were cut and fixed in 2% formaldehyde in
phosphate-buffered saline for 1 h at room temperature.
Immunocytochemical labeling was carried out using an indirect alkaline
phosphatase method. After blocking overnight 4 °C in
phosphate-buffered saline containing 2% bovine serum albumin and 0.2%
Triton X-100, sections were incubated with 2.5 µg/µl anti-HA mAb
for 90 min at room temperature. After washing in phosphate-buffered
saline, sections were incubated for 90 min at room with an alkaline
phosphatase-conjugated anti-mouse secondary antibody (diluted 1:2000;
Roche Molecular Biochemicals) in phosphate-buffered saline. The
reaction was detected using the Bio-Rad alkaline phosphatase kit. The
coverslips were mounted in Eukitt (O. Kindler GmbH & Co., Freiburg,
Germany), analyzed, and photographed using a Leica DMRB microscope.
Electrophysiological Recordings
Xenopus Oocytes--
In vitro transcribed RNA was
injected into Xenopus oocytes (~10 ng/oocyte) as described
(36). Ionic currents were recorded 2-4 days after injection using a
two-electrode voltage clamp (TEC 10CD, NPI Electronic, Tamm, Germany)
and PULSE Version 8.09 acquisition software (Heka Electronic,
Lambrecht, Germany). The oocytes were continually perfused in
barium-containing Ringer's solution (3.8 mM
K+, 114.2 mM Na+, 2 mM
Ba2+, and 10 mM TES, pH 7.4). Electrodes were
pulled from Corning 7052 glass (Garner Glass, Clairemont, CA) on a P-97
puller (Sutter Instrument Co., Novato, CA). Electrodes were filled with
1 M KCl buffered with 10 mM TES and typically
had a resistance of <500 kilo-ohms. The currents were sampled at 4-5
kHz after filtering at 1 kHz. Leak subtraction was accomplished with
two inverted quarter amplitude prepulses that were scaled and
subtracted from the test pulse. All recordings were made at room
temperature (~21 °C).
COS-7 Cells--
Membrane currents were recorded in the
whole-cell configuration as described in detail (37). Patch pipettes
were made from borosilicate glass capillaries (Clark Electromedical,
Pangbourne, United Kingdom) using a horizontal puller (P-87,
Sutter Instrument Co.) and heat-polished (MF-83, Narishige,
Tokyo, Japan) to have a resistance of 2-4 megaohms when filled
with the standard internal solution (144 mM KCl, 2 mM MgCl2, 10 mM TES, and 5 mM EGTA, buffered with KOH to pH 7.2). All experiments were
performed with an external solution containing 140 mM NaCl,
4 mM KCl, 2 mM CaCl2, 2 mM MgCl2, 10 mM TES, and 5 mM glucose, buffered with NaOH to pH 7.4. Voltage stimulation and current recordings were obtained with a patch-clamp amplifier (Jens Meyer, Munich, Germany) interfaced (Labmaster TL-1,
Axon Instruments, Inc., Foster City, CA) with a microcomputer equipped
with pClamp Version 5.5.1 software (Axon Instruments, Inc.). The
currents were low pass-filtered at 3 kHz (
3 dB) and acquired at
different sampling rates according to the stimulation protocols.
Capacitive transients and series resistance were minimized with the
analog circuits of the amplifier. In some experiments, membrane
currents were leak-subtracted by using a P/4 protocol (38). An agar
bridge electrode, filled with 150 mM NaCl, was used as the
reference electrode. All experiments were performed at room temperature
(~21 °C).
 |
RESULTS |
Design of Chimeric Channels--
Given that KcsA exhibits
significant sequence similarity to the S5-P-S6 region of Kv channels,
it was hypothesized that a chimeric construct consisting of the
ancestor KcsA linked to the putative eukaryotic voltage sensor would
exhibit voltage-dependent gating properties. For this task,
we constructed chimeric channels that combined the S1-S4 domain of
mKv1.1 or Shaker and the S5-P-S6 domain of the KcsA channel (Fig.
1, A-C, left
panels). These chimeras are referred to as mKv1.1(S1-S4)-KcsA,
mKv1.1(S1-S4,5)-KcsA, and Shaker(S1-S4,5)-KcsA, where S1-S4 denotes
the N-terminal domain of the Kv channels up to and including the S4
transmembrane segment and S1-S4,5 additionally incorporates the S4-S5
loop from the eukaryotic channels. We considered the intracellular
S4-S5 loop because of its contribution to pore properties (39). The
C-terminal end of all chimeras was tagged with a hemagglutinin epitope
to facilitate the biochemical and immunological analyses.

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Fig. 1.
A-C, topological models of mKv1.1,
Shaker, and chimeras (left panels) and
voltage-dependent channel activities of mKv1.1, Shaker, and
chimeras (right panels), respectively, in Xenopus
oocytes injected with the corresponding transcripts. Oocytes were held
at 80 mV and depolarized up to +100 mV in voltage steps of 10 mV.
D, molecular design of chimeras containing mKv1.1 and KcsA
as voltage sensor and pore module, respectively. In
mKv1.1(S1-S4,5)-KcsA, the S4-S5 loop is from mKv1.1, whereas in
mKv1.1(S1-S4)-KcsA, it is absent, and KcsA is full-length.
Shaker(S1-S4,5)-KcsA is formed by Shaker as voltage module and KcsA as
pore module, identical to the mKv1.1(S1-S4,5)-KcsA chimera. The
S4-S5 loop arrow and the vertical lines indicate
the approximate location of the chimeric joint. Scale
bar = 200 base pairs. E, immunochemical analysis
of protein expression in Xenopus oocytes. After cRNA
injection in oocytes, proteins were extracted and blotted as total
fractions, or immunoprecipitates (IP) were generated with
the anti-hemagglutinin antibody. Non-injected oocytes were used as a
negative control, and mKv1.1-injected oocytes were used as a positive
control. Lane 1-3, mKv1.1, mKv1.1(S1-S4,5)-KcsA, and
Shaker(S1-S4,5)-KcsA total proteins, respectively; lanes
5-7, immunoprecipitated fractions. Lane 4 was
non-injected.
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Designed Chimeras Do Not Exhibit Voltage-dependent
Channel Activity in Xenopus Oocytes or Mammalian Cells--
We first
assessed whether the designed chimeras exhibit
voltage-dependent channel gating. As illustrated in Fig.
1(A-C, right panels), at variance with wild-type
mKv1.1 and Shaker, heterologous expression of the
mKv1.1(S1-S4,5)-KcsA, mKv1.1(S1-S4)-KcsA, and Shaker(S1-S4,5)-KcsA
chimeras did not elicit voltage-activated outward currents from oocytes
held at
80 mV and depolarized from
70 to +100 mV (n = 32). The lack of functional expression was not overcome by larger
hyper- or depolarizing steps, by injecting increasing amounts of cRNA,
or by co-injecting the
- and
-subunits (data not shown). At
variance with homomeric wild-type Shaker-expressing oocytes,
chimera-injected cells did not display gating currents (data not
shown). The absence of functional expression was not due to lack of
protein synthesis, as evidenced by immunodetection of the
heterologously expressed chimeras (Fig. 1E). The immunoblot displays the presence of two bands with molecular masses of ~60 and
~62 kDa in oocytes injected with mKv1.1(S1-S4,5)-KcsA (Fig. 1E, lanes 2 and 6) and
Shaker(S1-S4,5)-KcsA (lanes 3 and 7) chimeras, respectively. mKv1.1-injected oocytes exhibited a band of ~67 kDa
(Fig. 1E, lanes 1 and 5). These
proteins were selectively immunopurified with anti-HA mAb (Fig.
1E, lanes 5-7)
We next addressed the question of whether the heterologous expression
system was inadequate for expression of chimeras. cDNAs encoding
chimeric channels were then cotransfected in COS-7 cells with GFP to
facilitate detection of transfected cells. Fluorescent cells were
selected for electrophysiological measurements (Fig. 2A). As for oocytes,
transfection of chimeras did not result in the expression of
voltage-dependent ionic currents in response to either
depolarizing steps up to +60 mV or hyperpolarization to
130 mV
(n = 23) (Fig. 2, B and C). In
contrast, cells transfected with mKv1.1 or Shaker channels expressed
sustained outward currents with kinetic properties overlapping
those obtained in oocytes (data not shown).

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Fig. 2.
Chimeric constructs expressed in COS-7 cells.
A, shown is a fluorescence photomicrograph of COS-7 cells
cotransfected with GFP and the mKv1.1(S1-S4,5)-KcsA chimera and
visualized 72 h post-transfection. Scale bar = 60 µm. B, a voltage stimulation protocol was employed to
activate tight-seal, whole-cell currents in COS-7 cells and consisted
of hyperpolarizing and depolarizing steps from 130 to +60 mV in 10-mV
increments. The holding potential was 60 mV. C, none of
the chimeric constructs produced voltage-gated currents. Shown is a
representative example of a COS-7 cell transfected with
mKv1.1(S1-S4,5)-KcsA at extracellular pH 7.4. D, in the
same cell, no currents were elicited by lowering the extracellular pH
to 4.0 for 2 min.
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Since strong acidification (pH < 5) favors KcsA channel gating (25),
we investigated whether extra- and intracellular acidification (pH 4)
could promote the appearance of voltage-gated channel activity in
chimera-transfected cells. The data indicate that neither extracellular (n = 4) (Fig. 2D) nor intracellular
(n = 5) (data not shown) acidification evoked channel
activity in response to voltage steps. These observations indicate that
the linkage of the putative voltage sensor of Kv channels to KcsA does
not endow the prokaryotic channel with voltage-dependent channel gating activity.
KcsA-containing Chimeras Exhibit a Dominant-negative
Phenotype--
To investigate if the translated chimeric subunits have
the ability to interact with wild-type channel subunits, we examined whether the chimeras disturbed the channel activity exhibited by
wild-type mKv1.1 and Shaker channels in a coexpression experiment. Xenopus oocytes were co-injected with both types of
subunits, and the channel activity of the presumed heteromeric proteins was compared with that displayed by homomeric proteins. Total cRNA
injected was constant in all samples. As illustrated in Fig. 3A, oocytes injected with
wild-type subunits showed robust voltage-dependent channel
activity. By contrast, coexpression of wild-type and chimeric subunits resulted in a virtually complete suppression of wild type-like
channel activity. This negative dominance of chimeric subunits was more
effective with wild-type mKv1.1 subunits than with wild-type Shaker
subunits, as evidenced by the lower amount of chimeric subunit required
to inhibit the functional activity of mKv1.1 channels (Fig.
3A). The reduction of wild-type channel activity was not due
to injection of lower wild-type subunit cRNA amounts since the
magnitude of voltage-elicited ionic currents remained fairly invariant
as the cRNA injected was decreased from 20 ng (8.2 ± 2.1 µA for
mKv1.1 (n = 4) and 12.2 ± 3.0 µA for Shaker (n = 4)) to 5 ng (6.1 ± 1.9 µA for mKv1.1
(n = 4) and 11.2 ± 2.7 µA for Shaker
(n = 4)). Therefore, these data indicate a
dominant-negative phenotype of the chimeras on wild-type channels.

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Fig. 3.
A, chimeric subunits exerted a
dominant-negative effect on Shaker and mKv1.1 expression. Wild-type
(WT) Shaker or mKv1.1 was coexpressed with the indicated
chimeric constructs (Chi) at 4:0, 3:1, 2:2, 1:3, and 0:4
molar ratios. Xenopus oocytes were clamped at 80 mV and
depolarized to 0 mV, and current amplitudes were normalized.
B, wild-type mKv1.1 and mKv1.1(S1-S4,5)-KcsA subunit
mRNAs were co-injected in Xenopus oocytes,
immunoprecipitated, and subjected to immunoblot analysis. The anti-HA
epitope antibody was used for immunoprecipitation (IP), and
the immunoblot (IB) was probed with an anti-mKv1.1 antibody
raised against a C-terminal peptide sequence present only in the
wild-type subunits. The molecular mass standards (MS)
are shown in the first lane. The approximate positions of
the protein constructs are indicated on the right. NI,
Non-injected. C, Kv channels and chimeric constructs
were coexpressed in Xenopus oocytes, immunoprecipitated, and
subjected to electrophoresis gel. The anti-HA epitope antibody, present in the C terminus of all chimeric
constructs, was used for immunoprecipitation, and the blot was probed
with anti-Shaker polyclonal antibody. Both mature (upper
band) and immature (lower band) forms of the Shaker
protein are visible, as indicated. D, immunoprecipitates
obtained from samples generated at 3:1 and 0:4 molar ratios of
wild-type Shaker and Shaker(S1-S4,5)-KcsA were subjected to
N-glycosidase F digestion. The glycosidase hydrolyzed all
N-glycosidated chains from either the homomeric or
heteromeric channels.
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The occurrence of a dominant-negative phenotype suggests the formation
of stable, nonfunctional, heteromeric channels composed of chimeric and
wild-type subunits. We used an immunology-based strategy to evaluate
this hypothesis. Oocytes co-injected with different molar ratios of
wild-type (mKv1.1 or Shaker) and chimeric (mKv1.1(S1-S4,5)-KcsA or
Shaker(S1-S4,5)-KcsA) subunits were lysed 72 h post-injection,
and chimeric subunits were immunoprecipitated with anti-HA mAb,
followed by SDS-polyacrylamide gel electrophoresis. Coprecipitation of
mKv1.1 and Shaker subunits was revealed by immunoblotting using either
an anti-mKv1.1 antibody raised against the C-terminal domain or an
anti-Shaker antibody raised against the N-terminal portion,
respectively. As shown in Fig. 3B (upper panel),
immunoblots probed with anti-Kv1.1 mAb revealed a band of >62 kDa
corresponding to mKv1.1 in oocytes co-injected with mKv1.1 and
mKv1.1(S1-S4,5)-KcsA. A band of
62 kDa was also evident as the ratio
of mKv1.1(S1-S4,5)-KcsA was increased, indicating that the chimeric
channels were also recognized by anti-mKv1.1 antibody, consistent with
the presence of a small portion of the epitope in the chimeric subunit
(Fig. 1D). As expected, these protein bands were absent in
mKv1.1-injected and non-injected oocytes. The intensity of the
coprecipitated mKv1.1 subunit declined as the amount of
mKv1.1(S1-S4,5)-KcsA increased. In contrast, probing the same
immunoblots with anti-HA mAb exposed a band of
62 kDa only in oocytes
injected with mKv1.1(S1-S4,5)-KcsA, which was augmented as the
ratio of the chimeric subunit was increased (Fig. 3B,
lower panel). These data are consistent with the formation of stable hetero-oligomers composed of wild-type mKv1.1 and the mKv1.1(S1-S4,5)-KcsA chimera.
Similarly, immunoblots probed with an anti-Shaker antibody raised
against the N-terminal domain unmasked the presence of two major bands
of ~75 kDa corresponding to Shaker in oocytes injected exclusively
with Shaker and Shaker(S1-S4,5)-KcsA subunits and a band of ~62 kDa
corresponding to Shaker(S1-S4,5)-KcsA in oocytes containing chimeric
subunits (Fig. 3C). The intensity of the ~75 kDa band
became more faint while that of the ~62 kDa band increased as the
amount of chimeric transcripts was augmented in the coexpression system, indicating that the composition of the assembled heteromeric channels is a function of the subunit concentration. The presence of
higher molecular mass bands (
115 kDa) in the immunoprecipitates (Fig.
3C), which may correspond to fully N-glycosylated
subunits in agreement with other reports (40, 41), is also noteworthy. Indeed, treatment of immunoprecipitates with N-glycosidase F
resulted in the complete and selective disappearance of the higher
molecular mass bands for both homomeric Shaker(S1-S4,5)-KcsA and
heteromeric Shaker·Shaker(S1-S4,5)-KcsA complexes (Fig.
3D). Accordingly, as for mKv1.1-containing chimeras, these
findings indicate the formation of stable heteromers between Shaker and
Shaker(S1-S4,5)-KcsA subunits. Furthermore, it appears that the
heteromeric complexes are N-glycosylated, suggesting that
they may fold correctly and even be distributed to the plasma membrane.
Homomeric Chimeras Are Delivered to the Plasma Membrane--
The
dominant-negative strategy suggests that chimeric subunits could
assemble as stable oligomers and be delivered to the plasma membrane,
where they form nonfunctional channels. However, the retention of these
complexes in the endoplasmic reticulum cannot be ruled out. To
distinguish between both possibilities, we examined the surface
expression of homomeric chimeras in the plasma membrane of
Xenopus oocytes by immunolabeling microscopy. Injected
oocytes were collected 72 h post-injection, and frozen sections
were probed by an indirect alkaline phosphatase method using anti-HA
primary antibody. As depicted in Fig. 4,
whereas non-injected oocytes did not show significant labeling of the plasma membrane, a clear distinct brown ring was evident surrounding wild-type mKv1.1- and chimeric mKv1.1(S1-S4,5)-KcsA- and
Shaker(S1-S4,5)-KcsA-injected oocytes. Therefore, this assay indicates
that a population of both mKv1.1(S1-S4,5)-KcsA and
Shaker(S1-S4,5)-KcsA chimeric oligomers was processed and not retained
in the intracellular compartments. Homo-oligomeric proteins assembled
in the plasma membrane appear to be nonfunctional channels.

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Fig. 4.
Surface expression of wild-type mKv1.1-HA or
homomeric chimeras in the plasma membrane of Xenopus
oocytes detected by immunolabeling microscopy. Non-injected
oocytes did not show significant labeling of the plasma membrane
(A), whereas a clear distinct brown ring (arrows)
surrounded wild-type mKv1.1-injected (B) and chimeric
mKv1.1(S1-S4,5)-KcsA-injected (C) and
Shaker(S1-S4,5)-KcsA-injected (D) oocytes. Scale
bar = 50 µ m.
|
|
 |
DISCUSSION |
The remarkable sequence similarity between the prokaryotic KcsA
channel and the pore domain of Kv channels suggests that KcsA may be a
bacterial ancestor of these eukaryotic channel proteins. Our objective
here was to evaluate this model by examining whether KcsA could
structurally and functionally replace the pore domain (S5-P-S6) of
voltage-gated channels; and additionally, we explored the possibility
of endowing the prokaryotic KcsA channel with voltage-dependent gating activity. Replacement of the pore
domain of mouse brain (mKv1.1(S1-S4,5)-KcsA) or that of Shaker
(Shaker(S1-S4,5)-KcsA) created chimeric channels that failed to
express voltage-dependent channel activity or gating
currents in Xenopus oocytes and a cell line, although they
displayed a conspicuous dominant-negative phenotype when coexpressed
with wild-type mKv1.1 or Shaker subunits (Fig. 1). The lack of
functional expression appeared not to arise from the synthesis of
incomplete proteins, as evidenced by the presence of proteins of the
expected size in both heterologous expression systems (Fig. 2). A
plausible explanation for the nonfunctional phenotype is the retention
of chimeric channels in the endoplasmic reticulum because of a
misfolding of the subunits that prevents normal trafficking. However,
analysis of the pathway of channel biogenesis suggested that chimeras
were properly folded and targeted to the cell surface. First, the
dominant-negative phenotype exhibited by both chimeric subunits
correlates with the formation of stable hetero-oligomers with wild-type
subunits (Fig. 3). Second, a significant population of wild-type and
chimeric subunits appeared to be heavily glycosylated, presumably at
the two glycosylation sites located in the loop connecting the first
and second transmembrane segments (40). Although glycosylation is not
an essential requirement for the assembly of functional channels at the
cell membrane, the presence of two glycosylated forms in the homo- and
hetero-oligomer subunits suggests correct folding, assembly, and
endoplasmic reticulum/Golgi trafficking of at least a portion of
these channel proteins to the plasma membrane. Indeed,
immunocytochemical analysis using an anti-HA mAb showed that
homo-oligomers of chimeric channel subunits were efficiently expressed
on the oocyte plasma membrane, exhibiting a level of expression similar
to that of wild-type Kv subunits (Fig. 4). This is significant since
the endoplasmic reticulum contains a stringent quality control system
that retains misfolded, incomplete, or incorrectly assembled proteins,
and only fully assembled channels are transported to the Golgi for further processing and delivery to their functional location (41, 42).
Collectively, these findings demonstrate that replacement of the pore
module of Kv channel subunits such as mKv1.1 and Shaker with the pore
domain of KcsA gives rise to chimeric subunits that fold and assemble
into stable complexes that are delivered to the cell surface.
Therefore, the prokaryotic pore module represented by KcsA appears to
be structurally compatible with the voltage sensor of eukaryotic Kv
channels. However, the molecular linkage of two distinct protein
modules is not sufficient to express functional coupling between them.
A central question arises: why do chimeric channels fail to
display channel activity? The suppression of channel activity by
replacement of the pore domain of Kv channels with KcsA implies that
the precise coupling interaction between the voltage sensor module and
the putative docking site on the pore domain is not effectively
restored in the chimeric channels. Presumably, a discrete set of
specific and critical protein-protein interaction surfaces between both
modules are necessary to couple the movement of the S4 segment to
channel opening, but are not essential for correct protein folding.
Disruption or perturbation of these interactions would specifically
lead to a partial or complete uncoupling of both domains, thus giving
rise to the biogenesis of correctly assembled nonfunctional channels
(43). In support of this notion, fluorescence scanning studies have led
to the identification of protein rearrangements that correlate with
voltage-dependent gating and the postulation of the
existence of a docking site on the pore domain for accommodating the
sensor module (44). Furthermore, a tryptophan scanning strategy has
identified an interaction surface for voltage-sensing domains on the
pore domain of Shaker near the interface between adjacent pore
domain subunits (45). Mutation of residues at the cytoplasmic one-third
of the pore results in significant changes in
voltage-dependent gating (45). Interestingly, the lowest
sequence similarity between the pore modules of Kv channels and KcsA is
constrained to the internal domain, where the activation gate may be
located (43, 46-49), suggesting a plausible structural divergence in
this region. Recent data from blocker protection in the pore of
voltage-gated Kv channels are also consistent with the presence of a
kink at the level of the highly conserved PXP motif present
in the S6 helices of Kv channels (50). A kink on the cytoplasmic side
of the S6 helix may provide the structural flexibility required to
couple voltage sensor movements to the activation gate in Kv channels.
This structural relaxation is absent in KcsA because this pore lacks
the PXP motif in the second transmembrane domain (43, 48,
50). The gating mechanism of KcsA appears to involve rigid body motions
of both transmembrane helices (48). Accordingly, it is plausible that
KcsA-containing chimeras are not functional because the rigidity of the
KcsA pore imposes higher activation energies to couple the
voltage-sensing machinery to the channel gate. Should this hypothesis
be valid, the reconstitution of the interaction surface of the voltage
sensor in the pore domain of KcsA may endow the chimeric channels with voltage-dependent channel activity.
In conclusion, our results indicate that the structural compatibility
of the voltage sensor and pore modules produces stable, folded
channels, but does not necessarily lead to channel activity. Functional
coupling of protein modules appears to depend on a constellation of
interactions that probably tune the energetic requirements for
efficient voltage gating. Further experimental work is necessary to
understand the intricacies underlying coupling of both channel modules.
 |
ACKNOWLEDGEMENTS |
We thank the members of the Ferrer-Montiel
laboratory for perceptive comments and helpful suggestions and
discussion. We are grateful to Reme Torres for technical assistance
with cRNA preparation and oocyte manipulation and injection, Alessia
Minardi for technical assistance with cell culturing, and Carolina
Garcia-Martinez for comments on the manuscript. We are indebted to L. Toro for the Shaker clone, S. Choe for the KcsA clone, F. Tejedor for
the anti-Shaker antibody, M. Canossa for the COS-7 cells, and R. Tsien
for the enhanced GFP clone.
 |
FOOTNOTES |
*
This work was supported by grants from the
Comisión Interministerial de Ciencia y Tecnología and
European Commission Fondos Europeos Desarrollo Regional grant
1FD97-0662-C02-01, and La Fundació la Caixa grant 98/027-00 (to
A. F.-M.). Research carried out at the University of California was
supported by United States Public Health Service Grant GM-49711.The costs of publication of this article were defrayed in part by the
payment of page charges. The article
must therefore be hereby marked
"advertisement" in accordance with 18 U.S.C. Section
1734 solely to indicate this fact.
To whom correspondence should be addressed: Centro de
Biología Molecular y Celular, Universidad Miguel
Hernández, Avda Ferrocarril s/n, 03202 Elche Alicante, Spain.
Tel.: 34-96-665-8727; Fax: 34-96-665-8758; E-mail:
aferrer@umh.es.
Published, JBC Papers in Press, March 26, 2001, DOI 10.1074/jbc.M100487200
 |
ABBREVIATIONS |
The abbreviations used are:
mKv1.1, mouse Kv1.1;
HA, hemagglutinin;
GFP, green fluorescent protein;
mAb, monoclonal
antibody;
TES, 2-{[2-hydroxy-1,1-bis(hydroxymethyl)ethyl]amino}ethanesulfonic
acid.
 |
REFERENCES |
1.
|
Rudy, B.
(1988)
Neuroscience
25,
729-749[CrossRef][Medline]
[Order article via Infotrieve]
|
2.
|
Tempel, B.,
Papazian, D.,
Schwarz, T.,
Jan, Y.,
and Jan, L.
(1987)
Science
237,
770-775[Medline]
[Order article via Infotrieve]
|
3.
|
Noda, M.,
Shimizu, S.,
Tanabe, T.,
Takai, T.,
Kayano, T.,
Ikeda, T.,
Takahashi, H.,
Nakayama, H.,
Kanaoka, Y.,
Minamino, N.,
Kangawa, K.,
Matsuo, H.,
Raftery, M. A.,
Hirose, T.,
Inayama, S.,
Hayashida, H.,
Miyata, T.,
and Numa, S.
(1984)
Nature
312,
121-127[Medline]
[Order article via Infotrieve]
|
4.
|
Tanabe, T.,
Takeshima, H.,
Mikami, A.,
Flokerzi, V.,
Takahashi, H.,
Kangawa, K.,
Kojima, M.,
Matsuo, H.,
Hirose, T.,
and Numa, S.
(1987)
Nature
328,
313-318[CrossRef][Medline]
[Order article via Infotrieve]
|
5.
|
Greenblatt, R.,
Blatt, Y.,
and Montal, M.
(1985)
FEBS Lett.
193,
125-134[CrossRef][Medline]
[Order article via Infotrieve]
|
6.
|
Jan, L.,
and Jan, Y.
(1997)
Annu. Rev. Neurosci.
20,
91-123[CrossRef][Medline]
[Order article via Infotrieve]
|
7.
|
Heginbotham, L.,
Lu, Z.,
Abramson, T.,
and MacKinnon, R.
(1994)
Biophys. J.
66,
1061-1067[Abstract]
|
8.
|
Papazian, D.,
Timpe, L.,
Jan, Y.,
and Jan, L.
(1991)
Nature
349,
305-310[CrossRef][Medline]
[Order article via Infotrieve]
|
9.
|
McCormack, K.,
Tanouye, M. A.,
Iverson, L. E.,
Lin, J. W.,
Ramaswami, M.,
McCormack, T.,
Campanelli, J. T.,
Mathew, M. K.,
and Rudy, B.
(1991)
Proc. Natl. Acad. Sci. U. S. A.
88,
2931-2935[Abstract]
|
10.
|
Lopez, G. A.,
Jan, Y.,
and Jan, L.
(1991)
Neuron
7,
327-336[CrossRef][Medline]
[Order article via Infotrieve]
|
11.
|
Perozo, E.,
Santacruz-Tolosa, L.,
Stefani, E.,
Bezanilla, F.,
and Papazian, D.
(1994)
Biophys. J.
66,
345-354[Abstract]
|
12.
|
Planells-Cases, R.,
Ferrer-Montiel, A.,
Patten, C.,
and Montal, M.
(1995)
Proc. Natl. Acad. Sci. U. S. A.
92,
9422-9426[Abstract]
|
13.
|
Seoh, S.,
Sigg, D.,
Papazian, D. M.,
and Bezanilla, F.
(1996)
Neuron
16,
1159-1167[Medline]
[Order article via Infotrieve]
|
14.
|
Perozo, E.,
MacKinnon, R.,
Bezanilla, F.,
and Stefani, E.
(1993)
Neuron
11,
353-358[Medline]
[Order article via Infotrieve]
|
15.
|
Montal, M.
(1995)
Annu. Rev. Biophys. Biomol. Struct.
24,
31-57[CrossRef][Medline]
[Order article via Infotrieve]
|
16.
|
Montal, M.
(1996)
Curr. Opin. Struct. Biol.
6,
499-510[CrossRef][Medline]
[Order article via Infotrieve]
|
17.
|
Nelson, R.,
Kuan, G.,
Saier, M.,
and Montal, M.
(2000)
J. Mol. Microbiol. Biotechnol.
1,
281-287
|
18.
|
Ho, K.,
Nichols, C.,
Lederer, W.,
Lytton, J.,
Vassilev, P.,
Kanazirska, V.,
and Hebert, S.
(1993)
Nature
362,
31-38[CrossRef][Medline]
[Order article via Infotrieve]
|
19.
|
Ketchum, K.,
Joiner, W.,
Sellers, A.,
Kaczmarek, L.,
and Goldstein, S.
(1995)
Nature
376,
690-695[CrossRef][Medline]
[Order article via Infotrieve]
|
20.
|
Lesage, F.,
Guillermare, E.,
Fink, M.,
Duprat, F.,
Lazdunski, M.,
Romey, G.,
and Barhanin, J.
(1996)
J. Biol. Chem.
271,
4183-4187[Abstract/Free Full Text]
|
21.
|
Lesage, F.,
Guillermare, E.,
Fink, M.,
Duprat, F.,
Lazdunski, M.,
Romey, G.,
and Barhanin, J.
(1996)
EMBO J.
18,
1004-1011
|
22.
|
Maingret, F.,
Patel, A. J.,
Lesage, F.,
Lazdunski, M.,
and Honore, E.
(1999)
J. Biol. Chem.
274,
26691-26696[Abstract/Free Full Text]
|
23.
|
Schrempf, H.,
Schmidt, O.,
Kümmerlen, R.,
Hinnah, S.,
Müller, D.,
Betzler, M.,
Steinkamp, T.,
and Wagner, R.
(1995)
EMBO J.
14,
5170-5178[Abstract]
|
24.
|
Cortes, M.,
and Perozo, E.
(1997)
Biochemistry
36,
10343-10352[CrossRef][Medline]
[Order article via Infotrieve]
|
25.
|
Heginbotham, L.,
Le Masurier, M.,
Kolmakova-Partensky, L.,
and Miller, C.
(1999)
J. Gen. Physiol.
114,
551-560[Abstract/Free Full Text]
|
26.
|
MacKinnon, R.,
Cohen, S.,
Kuo, A.,
Lee, A.,
and Chait, B.
(1998)
Science
280,
106-109[Abstract/Free Full Text]
|
27.
|
Doyle, D.,
Cabral, J.,
Pfuetzner, R.,
Kuo, A.,
Gulbis, J.,
Cohen, S.,
Chait, B.,
and MacKinnon, R.
(1998)
Science
280,
69-77[Abstract/Free Full Text]
|
28.
|
Patten, C.,
Caprini, M.,
Plannells-Cases, R.,
and Montal, M.
(1999)
FEBS Lett.
17,
375-381
|
29.
|
Sambrook, J.,
Fritsch, E.,
and Maniatis, T.
(1989)
Molecular Cloning: A Laboratory Manual
, 2nd Ed.
, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY
|
30.
|
Stefani, E.,
Toro, L.,
Perozo, E.,
and Bezanilla, F.
(1994)
Biophys. J.
66,
996-1010[Abstract]
|
31.
|
Sanger, F.,
Nicklen, S.,
and Coulson, A.
(1977)
Proc. Natl. Acad. Sci. U. S. A.
74,
5463-5467[Abstract]
|
32.
|
Ferroni, S.,
Planells-Cases, R.,
Ahmed, C. M. I.,
and Montal, M.
(1992)
Eur. Biophys. J.
21,
185-191[Medline]
[Order article via Infotrieve]
|
33.
|
Luthman, H.,
and Magnusson, G.
(1983)
Nucleic Acids Res.
11,
1295-1308[Abstract]
|
34.
|
Sun, W.,
Ferrer-Montiel, A.,
Schinder, A.,
McPherson, J.,
Evans, G.,
and Montal, M.
(1992)
Proc. Natl. Acad. Sci. U. S. A.
89,
1443-1447[Abstract]
|
35.
|
Ferrer-Montiel, A.,
Canaves, J.,
DasGupta, B.,
Wilson, M.,
and Montal, M.
(1996)
J. Biol. Chem.
271,
18322-18325[Abstract/Free Full Text]
|
36.
|
Ferrer-Montiel, A.,
and Montal, M.
(1994)
Methods Companion Methods Enzymol.
6,
60-69[CrossRef]
|
37.
|
Hamill, P.,
Marty, A.,
Neher, E.,
Sakmann, B.,
and Sigworth, F.
(1981)
Pfluegers Arch. Eur. J. Physiol.
391,
85-100[Medline]
[Order article via Infotrieve]
|
38.
|
Armstrong, C.,
and Bezanilla, F.
(1977)
J. Gen. Physiol.
70,
567-590[Abstract]
|
39.
|
Slesinger, P.,
Jan, Y.,
and Jan, L.
(1993)
Neuron
11,
739-749[Medline]
[Order article via Infotrieve]
|
40.
|
Santacruz-Tolosa, L.,
Huang, Y.,
Scott, J.,
and Papazian, D.
(1994)
Biochemistry
33,
5607-5613[Medline]
[Order article via Infotrieve]
|
41.
|
Schulteis, C.,
Nagaya, N.,
and Papazian, D.
(1998)
J. Biol. Chem.
273,
26210-26217[Abstract/Free Full Text]
|
42.
|
Zerangue, N.,
Schwappach, B.,
Jan, Y.,
and Jan, Y.
(1999)
Neuron
22,
537-548[Medline]
[Order article via Infotrieve]
|
43.
|
Durrel, S.,
Hao, Y.,
and Guy, R.
(1998)
J. Struct. Biol.
121,
263-284[CrossRef][Medline]
[Order article via Infotrieve]
|
44.
|
Gandhi, C.,
Loots, E.,
and Isacoff, E.
(2000)
Neuron
27,
585-595[Medline]
[Order article via Infotrieve]
|
45.
|
Li-Smerin, Y.,
Hackos, D.,
and Swartz, K. J.
(2000)
Neuron
25,
411-423[Medline]
[Order article via Infotrieve]
|
46.
|
Holmgren, M.,
Shin, K.,
and Yellen, G.
(1998)
Neuron
2,
1617-1621
|
47.
|
Liu, Y.,
Holmgren, M.,
Jurman, E.,
and Yellen, G.
(1997)
Neuron
19,
175-184[Medline]
[Order article via Infotrieve]
|
48.
|
Perozo, E.,
Cortes, D.,
and Cuello, L.
(1999)
Science
285,
73-78[Abstract/Free Full Text]
|
49.
|
Tatulian, S.,
Cortes, M.,
and Perozo, E.
(1998)
FEBS Lett.
423,
205-212[CrossRef][Medline]
[Order article via Infotrieve]
|
50.
|
del Camino, D.,
Holmgren, M.,
Liu, Y.,
and Yellen, G.
(2000)
Nature
403,
321-325[CrossRef][Medline]
[Order article via Infotrieve]
|
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