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N-Glycosylation-dependent Control of Functional Expression of Background Potassium Channels K2P3.1 and K2P9.1

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Two-pore domain potassium (K(2P)) channels play fundamental roles in cellular processes by enabling a constitutive leak of potassium from cells in which they are expressed, thus influencing cellular membrane potential and activity. Hence, regulation of these channels is of critical importance to cellular function. A key regulatory mechanism of K(2P) channels is the control of their cell surface expression. Membrane protein delivery to and retrieval from the cell surface is controlled by their passage through the secretory and endocytic pathways, and post-translational modifications regulate their progression through these pathways. All but one of the K(2P) channels possess consensus N-linked glycosylation sites, and here we demonstrate that the conserved putative N-glycosylation site in K(2P)3.1 and K(2P)9.1 is a glycan acceptor site. Patch clamp analysis revealed that disruption of channel glycosylation reduced K(2P)3.1 current, and flow cytometry was instrumental in attributing this to a decreased number of channels on the cell surface. Similar findings were observed when cells were cultured in reduced glucose concentrations. Disruption of N-linked glycosylation has less of an effect on K(2P)9.1, with a small reduction in number of channels on the surface observed, but no functional implications detected. Because nonglycosylated channels appear to pass through the secretory pathway in a manner comparable with glycosylated channels, the evidence presented here suggests that the decreased number of nonglycosylated K(2P)3.1 channels on the cell surface may be due to their decreased stability.
rK 2P 9.1 cell surface expression is less sensitive than rK 2P 3.1 to channel glycosylation state. A, currents evoked by applied membrane potential pulses from 100 to 90 mV in HEK293 cells expressing rK 2P 9.1 (WT) or rK 2P 9.1 N53Q (N53Q) at pH 7.8. B, average current-voltage relationship for untransfected (UT) HEK293 cells, or HEK cells expressing rK 2P 9.1 (WT) or rK 2P 9.1 N53Q (N53Q) at pH 7.8. C, flow cytometric analysis of intact COS-7 cells expressing GFPrK 2P 9.1-HA (WT) or GFP-rK 2P 9.1 N53Q HA (HA). The cells were stained with either a monoclonal antibody against the external HA tag or an isotype control antibody, followed by goat anti-mouse F(ab) 2 fragment conjugated to Alexa Fluor 647. Solid gray curve, cells expressing GFP-rK 2P 9.1-HA, stained with the isotype control. Solid black line, cells expressing GFP-rK 2P 9.1-HA, stained with anti-HA tag antibody. Solid gray line, GFP-rK 2P 9.1 N53Q HA, anti-HA-tag. Dashed gray line, GFP-rK 2P 9.1 N53Q HA, isotype control. Bar chart, summary of three independent experiments to determine the surface expression of GFP-rK 2P 9.1 N53Q HA (N53Q) to GFP-rK 2P 9.1-HA (WT). The results are expressed relative to WT (100%). The error bar shows S.E. D, immunofluorescence images of the COS-7 cells described above, fixed and stained to detect external HA tag (red fluorescence). Upper panels, GFP-rK 2P 9.1-HA; lower panels, GFP-rK 2P 9.1 N53Q HA (green fluorescence ). Merge, superimposed images of channel and HA tag fluorescence. The scale bar represents 10 m.
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N-Glycosylation-dependent Control of Functional Expression
of Background Potassium Channels K
2P
3.1 and K
2P
9.1
*
S
Received for publication, July 30, 2012, and in revised form, December 11, 2012 Published, JBC Papers in Press, December 18, 2012, DOI 10.1074/jbc.M112.405167
Alexandra Mant
, Sarah Williams
‡1
, Laura Roncoroni
‡1
, Eleanor Lowry
§
, Daniel Johnson
§
, and Ita O’Kelly
‡2
From the
Human Development and Health, Centre for Human Development, Stem Cells and Regeneration, Faculty of Medicine,
University of Southampton, Southampton SO16 6YD, United Kingdom and the
§
Faculty of Life Sciences, University of Manchester,
Manchester M13 9PT, United Kingdom
Background: N-Glycosylation regulates the function of many membrane proteins.
Results: K
2P
3.1 and K
2P
9.1 possess functional glycosylation sites, and a lack of glycosylation results in fewer channels on the cell
surface.
Conclusions: N-Linked glycosylation has a critical role in K
2P
3.1 and a modulatory role in K
2P
9.1 cell surface expression.
Significance: This defines a direct link between background potassium channel function and metabolic status.
Two-pore domain potassium (K
2P
) channels play fundamen-
tal roles in cellular processes by enabling a constitutive leak of
potassium from cells in which they are expressed, thus influenc-
ing cellular membrane potential and activity. Hence, regulation
of these channels is of critical importance to cellular function. A
key regulatory mechanism of K
2P
channels is the control of their
cell surface expression. Membrane protein delivery to and
retrieval from the cell surface is controlled by their passage
through the secretory and endocytic pathways, and post-trans-
lational modifications regulate their progression through these
pathways. All but one of the K
2P
channels possess consensus
N-linked glycosylation sites, and here we demonstrate that the
conserved putative N-glycosylation site in K
2P
3.1 and K
2P
9.1 is a
glycan acceptor site. Patch clamp analysis revealed that disrup-
tion of channel glycosylation reduced K
2P
3.1 current, and flow
cytometry was instrumental in attributing this to a decreased
number of channels on the cell surface. Similar findings were
observed when cells were cultured in reduced glucose concen-
trations. Disruption of N-linked glycosylation has less of an
effect on K
2P
9.1, with a small reduction in number of channels
on the surface observed, but no functional implications
detected. Because nonglycosylated channels appear to pass
through the secretory pathway in a manner comparable with
glycosylated channels, the evidence presented here suggests that
the decreased number of nonglycosylated K
2P
3.1 channels on
the cell surface may be due to their decreased stability.
Cellular membrane potential influences the function of both
excitable and nonexcitable cells. The two-pore domain potas-
sium (K
2P
) channels are a family of background channels that
regulate the membrane potential of cells in which they are
expressed. Both structurally and functionally dissimilar to
other potassium channel families, K
2P
channels display little
voltage or time dependence, are active at resting membrane
potentials, and allow a constitutive leak of K
from cells (1, 2).
The acid-sensitive K
2P
subgroup (TASK channels) includes two
well characterized members, K
2P
3.1 (TASK-1) and K
2P
9.1
(TASK-3), and a third proposed member (K
2P
15.1 or TASK-5),
which remains uncharacterized to date. Because of their cellu-
lar localization and sensitivity to physiological stimuli (extra-
cellular acidification and hypoxia) TASK channels have been
implicated in an array of physiological processes including reg-
ulatory roles in cell proliferation (and oncogenesis), activation
of T-cells, chemoreception, and neuroprotective roles in
response to ischemia and inflammation (3–5). TASK channels
are also molecular targets for both local anesthetics and endo-
cannabinoids (3, 6). These channels show constitutive activity
once expressed on the cell surface; hence the control of TASK
channel surface expression is of critical importance, because
any change in channel number at the plasma membrane
impacts the electrical properties of the cell in which these chan-
nels are expressed.
Numerous in-built processes within the secretory pathway
are employed to regulate delivery of correctly folded membrane
proteins, in appropriate number, to the cell surface. We and
others have previously shown that TASK channel phosphory-
lation and association with cytosolic adaptor protein 14-3-3 is
critical to K
2P
3.1 and K
2P
9.1 export from the endoplasmic retic-
ulum (ER)
3
and hence cell surface expression (7–10).
Understanding of the quality control processes newly syn-
thesized proteins undergo en route to the cell surface is still
developing. Key steps include retention of nascent proteins
within the ER until correctly folded and assembled, removal of
persistently misfolded proteins, and transport of correctly
folded proteins to the Golgi complex (GC). Further quality con-
trol together with protein maturation occurs within the endo-
plasmic reticulum-Golgi intermediate compartment (ERGIC)
*This work was supported by Biotechnology and Biological Sciences
Research Council Grant BB/J008168/1 (to I. O.).
S
This article contains supplemental Fig. S1.
1
Supported by the Gerald Kerkut Charitable Trust.
2
To whom correspondence should be addressed: Human Development and
Health, Duthie Building, Mail Point 808, University of Southampton,
Southampton SO16 6YD, UK. Tel.: 44-23-8079-6421; Fax: 44-23-8079-4264;
E-mail: I.M.O’Kelly@southampton.ac.uk.
3
The abbreviations used are: ER, endoplasmic reticulum; GC, Golgi complex;
ERGIC, ER-golgi intermediate compartment; r, rat; MFI, mean fluorescence
intensity; EEA1, early endosome antigen 1; HCN, hyperpolarization-acti-
vated cyclic nucleotide-gated; TASK, TWIK-related Acid Sensitive Potas-
sium channels.
THE JOURNAL OF BIOLOGICAL CHEMISTRY VOL. 288, NO. 5, pp. 3251–3264, February 1, 2013
© 2013 by The American Society for Biochemistry and Molecular Biology, Inc. Published in the U.S.A.
FEBRUARY 1, 2013VOLUME 288NUMBER 5 JOURNAL OF BIOLOGICAL CHEMISTRY 3251
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and GC that ultimately leads to delivery of mature membrane
proteins to the plasma membrane or removal of misfolded pro-
teins to the endosomes and lysosomes (11, 12).
Protein glycosylation can play a key role in these processes
and has previously been shown to be a critical modulator of ion
channel gating, trafficking, and stability (13–16). N-Linked gly-
cosylation occurs within the ER and undergoes further modifi-
cations within the GC (17). A large, preformed oligosaccharide
precursor is added to the nascent protein within the ER. Trim-
ming of specific glycans signals that the glycoprotein is ready
for transport to the GC for further processing. If the glycosy-
lated protein is unfolded or misfolded, a glucose residue is
added back to the initial oligosaccharide, preventing its export
to the GC (18, 19). Correct conformation of the protein triggers
removal of this glucose residue and protein release from the ER.
Similarly, within the GC, sugar moieties are rearranged. The
final glycan composition and number regulates glycoprotein
trafficking and stability (18, 20).
N-Glycosylation occurs on Asn in NX(S/T) motifs. Both
K
2P
3.1 and K
2P
9.1 carry a conserved, predicted glycosylation
site at position 53. We sought, therefore, to determine whether
these channels are glycosylated in vivo and whether glycosyla-
tion has a regulatory role in channel function.
EXPERIMENTAL PROCEDURES
Molecular Biology—HA (YPYDVPDYA)-tagged rat (r)
K
2P
3.1 has been described previously (10). Similarly, the HA tag
was introduced into GFP-rK
2P
9.1 between Ala-213 and Leu-
214, using Pfu Ultra DNA polymerase (Agilent Technologies
UK Ltd., Stockport, UK). A conserved, putative glycosylated
asparagine, Asn-53, was altered to glutamine in rK
2P
3.1 and
rK
2P
9.1, GFP-rK
2P
3.1, and GFP-rK
2P
9.1 and the HA-tagged
GFP-rK
2P
3.1 and GFP-rK
2P
9.1 (Table 1), also using Pfu Ultra
DNA polymerase. The DNA constructs were fully sequenced
before use.
Western Blotting—COS-7 cells were plated at 5 10
5
cells/
10-cm dish in DMEM with 10% FCS and then transiently trans-
fected with 10
g of DNA encoding GFP-rK
2P
3.1-HA, GFP-
rK
2P
3.1
N53Q
HA, GFP-rK
2P
9.1-HA, or GFP-rK
2P
9.1
N53Q
HA,
using jetPEI transfection reagent, according to the supplier’s
instructions (Polyplus; Source Bioscience Autogen, Notting-
ham, UK). DNA-transfection complexes were removed from
cells after 4 h and replaced with fresh DMEM with 10% FCS.
Transfected cells were allowed to recover for 1 h, and then
tunicamycin (or an equivalent volume of Me
2
SO) was added to
a final concentration of 1.0
g/ml. Control and tunicamycin-
treated samples were incubated for 16 h overnight at 37 °C, 5%
CO
2
. The cells were harvested by scraping on ice in PBS sup-
plemented with a protease inhibitor mixture (Thermo Fisher
Scientific and Perbio Science UK, Ltd., Cramlington, UK),
washed in PBS, and then lysed for 30 min on ice in 200
l of lysis
buffer (10 mMTris, pH 7.5, 150 mMNaCl, 0.5% Nonidet P-40)
supplemented with protease inhibitors. The lysates were cen-
trifuged at 5000 gfor 5 min at 4 °C, and the post-nuclear
supernatant was mixed with protein sample buffer containing
100 mMDTT (final) for K
2P
9.1 and 200 mMDTT (final) for
K
2P
3.1 and incubated for 30 min at room temperature. Samples
were separated by SDS-PAGE and transferred to nitrocellulose
membranes. The membranes were probed with either 1/1000
rabbit anti-GFP antibody (for K
2P
3.1, ab290; Abcam, Cam-
bridge, UK) or 1/1000 dilution anti-HA tag antibody (for
K
2P
9.1, mouse clone 16B12; Covance, Leeds, UK) and then a
horseradish peroxidase-conjugated anti-rabbit or anti-mouse
secondary antibody (Dako UK Ltd., Ely, UK), followed by detec-
tion using Pierce Super Signal West (Thermo Fisher Scientific).
Whole Cell Patch Clamping Recordings—HEK293 cells were
plated on 22-mm sterile coverslips in 6-well plates at 10
5
cells/
well. After 3 h, the cells were transiently transfected with either 1.5
g of untagged, full-length rK
2P
3.1 or rK
2P
3.1
N53Q
, or 0.1– 0.25
g
of rK
2P
9.1 or rK
2P
9.1
N53Q
in pcDNA3.1 and 0.75
g of eGFP-C1/
well of a 6-well plate (Clontech), as described above. Controls were
non-green fluorescent cells in the transfection wells.
Green fluorescent cells were selected for whole cell patch
clamp analysis 24 h post-transfection. Pipette solution was
K
-rich and contained 150 mMKCl, 1 mMMgCl
2
,10mM
HEPES, 2 mMEGTA, pH 7.2, with KOH; free [Ca
2
]27 nM.
Bath solution was Na
-rich and contained 135 mMNaCl, 5 mM
KCl, 1 mMMgCl
2
,10mMHEPES, 1 mMCaCl
2
, pH 7.8, with
NaOH. All of the experiments were carried out at room tem-
perature. Patch pipettes were manufactured from standard
walled borosilicate glass capillary tubing (1B150-4; World Pre-
cision Instruments) on a two-stage Narishige PC-10 pipette
puller (Narishige Scientific Instrument Laboratory, Kasuya,
Tokyo, Japan), were heat-polished on a Narishige microforge,
and had measured tip resistances of 2–5 M(when filled with
TABLE 1
Summary of K
2P
constructs used in this study
Channels were tagged with eGFP fused to the N terminus, with or without an HA epitope tag in the external loop of the second pore-forming domain.
Tag Mutation Experiments
K
2P
3.1 rat channel
rK
2P
3.1 None Wild type Electrophysiology
rK
2P
3.1
N53Q
None N53Q
GFP-rK
2P
3.1 eGFP Wild type Immunofluorescence
GFP-rK
2P
3.1
N53Q
eGFP N53Q
GFP-rK
2P
3.1-HA eGFP and HA Wild type Immunofluorescence, flow cytometry, and immunoblotting
GFP-rK
2P
3.1
N53Q
HA eGFP and HA N53Q
K
2P
9.1 rat channel
rK
2P
9.1 None Wild type Electrophysiology
rK
2P
9.1
N53Q
None N53Q
GFP-rK
2P
9.1 eGFP Wild type Immunofluorescence
GFP-rK
2P
9.1
N53Q
eGFP N53Q
GFP-rK
2P
9.1-HA eGFP and HA Wild type Immunofluorescence, flow cytometry, and immunoblotting
GFP-rK
2P
9.1
N53Q
HA eGFP and HA N53Q
Glycosylation of TASK Channels
3252 JOURNAL OF BIOLOGICAL CHEMISTRY VOLUME 288 •NUMBER 5 •FEBRUARY 1, 2013
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K
-rich pipette solution). Resistive feedback voltage clamp was
achieved using an Axopatch 200 B amplifier (Axon Instru-
ments, Foster City, CA). Voltage protocols were generated, and
currents were recorded using Clampex 10.2 employing Digi-
data 1400A (Axon Instruments). The data were filtered (four-
pole Bessel) at 1 kHz and digitized at 5 kHz. Following successful
transition to the whole cell recording mode, capacitance transients
were compensated for and measured. To evoke ionic currents
voltage step protocol (100 to 90 mV in 10 mV increments, 100
ms) was employed, and current-voltage relationships were con-
structed from the plateau stage of each 100-ms step.
Flow Cytometry—COS-7 cells were plated in 10 cm dishes,
510
5
/dish and transfected transiently with 10
g of plasmid
DNA encoding GFP-rK
2P
3.1-HA, GFP-rK
2P
3.1
N53Q
HA, eGFP
alone, or empty pcDNA3.1 (Invitrogen), as described above.
In tunicamycin-treated samples, the transfected cells were
allowed to recover for 1 h before addition of the antibiotic (final
concentration, 1.0, 0.1, or 0.01
g/ml) or Me
2
SO alone. After
overnight incubation (16 h), the cells were harvested using tryp-
sin and then stained at room temperature, with occasional gen-
tle agitation, for 1 h with anti-HA tag antibody (Covance) at
1/400 dilution or an isotype control (IgG1; Invitrogen), fol-
lowed by goat anti-mouse F(ab)
2
fragment conjugated to Alexa
Fluor 647 (Invitrogen;1hatroom temperature; darkness;
1/1000 dilution). Immediately prior to analysis, the cells were
stained with SYTOX AADvanced Dead Cell Stain (Invitrogen)
to exclude damaged cells from the subsequent flow cytometric
analyses (FACSCanto; BD Biosciences, Oxford, UK). Surface
expression for each sample was calculated as the mean fluores-
cence intensity (MFI) of HA tag-stained cells minus the MFI of
the corresponding isotype control. For low glucose experi-
ments, COS-7 cells were grown in DMEM with 10% FCS con-
taining 1 g/liter glucose (“low”) or 4.5 g/liter glucose (“high”) for
3–10 days, before transfection and analysis, as described above.
Microscopy—COS-7 cells were transfected transiently
with DNA constructs encoding GFP-rK
2P
3.1-HA, GFP-
rK
2P
3.1
N53Q
HA, GFP-rK
2P
9.1, or GFP-rK
2P
9.1
N53Q
on cover-
slips as described above, cultured overnight, then fixed with 4%
(w/v) formaldehyde in PBS for 7 min at room temperature,
blocked with 3% (w/v) BSA in PBS, and stained with anti-HA
tag antibody and goat anti-mouse F(ab)
2
fragment conjugated
to Alexa Fluor 647 or with biotinylated anti-mouse (Vector
Laboratories, Burlingame, CA) and streptavidin-Texas Red
(Vector Laboratories) as described above. For co-localization
studies, fixed cells were permeabilized with 0.1% Triton-X-100
in PBS before the blocking step. The primary antibodies were
mouse the anti-58-kDa Golgi protein (1/1000 dilution, clone
58K-9, Abcam), rabbit anti-ERGIC-53 (1/100 dilution, Sigma),
and goat anti-EEA1 (1/100 dilution, C-15, Santa Cruz).
For the recycling assay, COS-7 cells were transfected tran-
siently with GFP-rK
2P
3.1-HA or GFP-rK
2P
3.1
N53Q
HA, cul-
tured overnight, and then incubated for2hinDMEM without
FCS. After 90 min, cycloheximide (Sigma) was added to a final
concentration of 100
g/ml. At 2 h, the cells were moved to a
cold room (4 °C), placed on ice for 45 min, and then washed
three times in ice-cold PBS. Surface proteins were biotinylated
for 45 min on ice using 0.5 mg of EZ-Link NHS-SS-Biotin
(Thermo Fisher Scientific) per well of a 6-well plate. The cells
were then returned to DMEM with 10% FCS and 100
g/ml
cycloheximide and incubated for 0 and 20 min. At each time
point, coverslips were transferred to wells containing ice-cold
biotin stripping buffer (50 mMsodium methanethiolate, 50 mM
Tris-HCl, pH 8.6, 100 mMNaCl, 1 mMMgCl
2
, 0.1 mMCaCl
2
),
where they were incubated on ice for 15 min, the buffer was
exchanged, and then the incubation was repeated. The strip-
ping step was to remove noninternalized biotin from the
plasma membrane. The coverslips were then rinsed with ice-
cold PBS, fixed with 4% w/v formaldehyde as described above,
and stained for EEA1 and biotin. Biotin was detected with a
streptavidin-Alexa Fluor 546 conjugate (Invitrogen). Triple-
stained vesicles were identified by drawing transects through
regions of interest and looking for areas of signal overlap in all
three dimensions. Quantitative analysis was carried out using
Imaris 7.5.2 software. The coverslips were mounted and visual-
ized using either a Zeiss Axio Observer D1 or using a Leica TCS
SP5 confocal scanning microscope in the University of South-
ampton Biomedical Imaging Unit.
Statistics—Graph and statistical analysis software SigmaPlot
11.0 (Systat Software, Chicago, IL) was used to plot electrophys-
iology data and perform significance tests. The data were first
subjected to the Shapiro-Wilk test for normality. Normal pop-
ulations were analyzed using Student’s ttest. Non-normal pop-
ulations were analyzed using the Mann-Whitney rank sum test.
RESULTS
Glycosylation of K
2P
Channels—Protein sequences of human,
mouse, and rat K
2P
channels were analyzed for the presence of
N-glycosylation consensus sites using the NetNGlyc 1.0 server
(21). Sites with a glycosylation potential 0.5 were cross-
checked for predicted external domains. Thirteen of the fifteen
mammalian K
2P
channels sequenced to date contain at least
one predicted glycosylation site within at least one of the three
species examined (Table 2). Notably, using a threshold glyco-
sylation potential 0.5, neither K
2P
1.1 (TWIK) nor K
2P
15.1
TABLE 2
Predicted N-glycosylation sites in K
2P
channels
Protein sequences were analyzed using the NetNGlyc Server 1.0, hosted by the
Center for Biological Sequence Analysis at the Danish Technical University.
Channel Human Mouse Rat
K
2P
1.1 (TWIK-1) Asn-95 Asn-95 Asn-95
K
2P
2.1 (TREK-1) Asn-110 Asn-110 Asn-95
Asn-134 Asn-134 Asn-119
K
2P
3.1 (TASK-1) Asn-53 Asn-53 Asn-53
K
2P
4.1 (TRAAK) Asn-78
Asn-82 Asn-81 Asn-81
K
2P
5.1 (TASK-2) Asn-77 Asn-77 Asn-77
K
2P
6.1 (TWIK-2) Asn-85 Asn-83 Asn-79
Asn-85
K
2P
7.1 Asn-83 Asn-83
K
2P
9.1 (TASK-3) Asn-53 Asn-53 Asn-53
K
2P
10.1 (TREK-2) Asn-144 Asn-144
Asn-148 Asn-148
K
2P
12.1 (THIK-2) Asn-78 Asn-78 Asn-78
K
2P
13.1 (THIK-1) Asn-59 Asn-59 Asn-59
Asn-65 Asn-65 Asn-65
K
2P
15.1 (TASK-5) No sites No sites No sites
K
2P
16.1 (TALK-1) Asn-57 Asn-57 Asn-57
Asn-86 Asn-86 Asn-86
K
2P
17.1 (TALK-2) Asn-65
Asn-94
K
2P
18.1 (TRESK) Asn-70
Asn-96
a
Asn-94 Asn-83
a
This site is predicted, but only Asn-70 is glycosylated in human K
2P
18.1 (27).
Glycosylation of TASK Channels
FEBRUARY 1, 2013VOLUME 288NUMBER 5 JOURNAL OF BIOLOGICAL CHEMISTRY 3253
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were predicted to be glycosylated. However, K
2P
1.1 does con-
tain a glycosylation consensus sequence at positions 95–97
(conserved between human and rodent) with Asn-95 as a
potential glycan acceptor. Lesage et al. (22) reported K
2P
1.1 to
be a glycoprotein, because of altered band sizes following sep-
aration by PAGE of glycopeptidase-treated channels. Signifi-
cantly, K
2P
15.1, which to date has failed to show functional
expression, is the only K
2P
channel that lacks an N-glycosyla-
tion consensus site (Table 2).
Both K
2P
3.1 and K
2P
9.1 possess a single N-glycosylation site
at position 53 (Asn-53; rK
2P
3.1 position potential 0.689;
rK
2P
9.1 0.802). Asn-53 is located within the first external
domain adjacent to the first pore and is conserved within all
species examined (Fig. 1, Aand B).
To determine whether Asn-53 is in fact a target for glycan
attachment, we abolished the predicted glycosylation site by
substitution of glutamine for asparagine, creating the mutant
channels GFP-rK
2P
3.1
N53Q
HA and GFP-rK
2P
9.1
N53Q
HA. We
examined whether the wild-type and glycosylation mutant
channels were glycosylated when expressed in COS-7 cells.
COS-7 cells transiently expressing GFP-rK
2P
3.1-HA, GFP-
rK
2P
3.1
N53Q
HA, GFP-rK
2P
9.1-HA, and GFP-rK
2P
9.1
N53Q
HA
were treated either with 1.0
g/ml tunicamycin, which blocks
synthesis of all N-glycans, or with its vehicle Me
2
SO alone. The
relative mobility of the tagged channels from total cell lysates
was compared by SDS-PAGE, followed by Western blotting.
For K
2P
3.1, two distinct bands are detected in the nontreated
wild-type lane; on treatment with tunicamycin or in cells trans-
fected with GFP-rK
2P
3.1
N53Q
HA, the intensity of the upper
band is decreased, whereas that of the lower band increases,
suggesting a higher proportion of higher mobility channels (Fig.
1C). Untreated, wild-type K
2P
9.1 channels migrate as a single
band to the predicted weight. A proportion of K
2P
9.1 from the
tunicamycin-treated cells migrated further than their
FIGURE 1. K
2P
3.1 and K
2P
9.1 are glycoproteins. A, partial amino acid sequence alignment of mouse, rat, and human K
2P
3.1 and K
2P
9.1 with N-linked
glycosylation consensus site and channel pore selectivity sequence highlighted.B, schematic membrane topology of TASK channels subunit with four trans-
membrane domains (dark gray), pore-forming domains (medium gray), and external domains (light gray) with the location of the putative N-glycosylation site
depicted (N). C, immunoblot showing differences in mobility of WT and glycosylation mutant (N53Q) TASK channels with or without prior treatment with
tunicamycin to prevent the addition of N-linked glycans. Post-nuclear supernatants from untreated () or tunicamycin-treated () COS-7 cells transfected with
GFP- and HA-tagged K
2P
3.1 (upper panel) and K
2P
9.1 (lower panel) were separated by SDS-PAGE and transferred to nitrocellulose. The membranes were probed
with anti-GFP antibody (K
2P
3.1) or anti-HA tag antibody (K
2P
9.1). The positions of size standards are indicated on the left.Dotted arrows indicate the middle of
each band.
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untreated controls, whereas glycosylation mutant channels
show mobility similar to the tunicamycin-treated wild-type
channel and higher than the glycosylated wild-type channel
(Fig. 1C).
Disruption of K
2P
3.1 Glycosylation Prevents Channel Func-
tional Expression—To determine whether channel glycosyla-
tion impacts channel function, we investigated the electro-
physiological properties of wild-type and N53Q mutant rK
2P
3.1
in HEK293 cells. The cells expressing wild-type channel show a
current-voltage (I-V) relationship typical of K
2P
potassium leak
channels, with a reversal potential of 60.88 mV 3.20, (n8;
Fig. 2A). As predicted for TASK family members, rK
2P
3.1 cur-
rents were inhibited by exposure to acidic external solutions
(pH 6.5) with currents at 60 mV reduced from 0.97 0.17 nA at
pH 7.8 to 0.45 0.08 nA at pH 6.5, which is not significantly
different from currents produced by untransfected cells (0.39
0.09 nA at pH 7.8, n8; (p0.63); Fig. 2, Aand B). HEK293
cells transiently expressing rK
2P
3.1
N53Q
channels failed to pro-
duce current significantly different from untransfected cells at
both pH 7.8 and pH 6.5 (0.41 0.08 nA at pH 7.8, n7): p
0.81; 0.43 0.10 nA at pH 6.5 n9) and p0.79 (Fig. 2, Aand
B). This decrease in rK
2P
3.1
N53Q
current compared with wild-
type rK
2P
3.1 current is coupled with a more positive reversal
potential (31.22 6.01 mV), which is predicted if background
K
flux is reduced.
To determine whether the observed decreased flux is due to
altered channel function or a reduced number of channels on
the cell surface, channel cellular localization and expression on
the cell surface was investigated in COS-7 cells transiently
expressing either the wild-type or glycosylation mutant K
2P
3.1
channels. Channel constructs contained an internal N-terminal
GFP tag plus a noninterfering HA tag incorporated into an
external loop within the second pore-forming domain of the
channel. The GFP tag made it possible to monitor total expres-
sion (internal and cell surface) of the wild-type and mutant
channel. The external HA tag enabled quantitative comparison
of the relative amount of each channel expressed at the cell
surface by means of staining nonpermeablized cells with
anti-HA antibodies, followed by flow cytometry (Fig. 2, Cand
D).
In a typical experiment, the mean surface expression level of
double-tagged rK
2P
3.1
N53Q
in live, intact cells was only 9% rel-
ative to the wild-type channel, as measured by staining exter-
nally with anti-HA tag (Fig. 2C; MFI values as follows: GFP-
rK
2P
3.1-HA, 294; GFP-rK
2P
3.1
N53Q
HA, 97; isotype control,
78). Significantly, when cells expressing double-tagged rK
2P
3.1
channels were treated with 1.0
g/ml tunicamycin, a reduction
in HA tag fluorescence was observed (MFI 101), comparable
with cells expressing the mutant channel. Over a series of
experiments, summarized in Fig. 2D, disruption of glycosyla-
tion by mutation of the channel always resulted in very low cell
surface expression of K
2P
3.1
N53Q
(mean 11.3% relative to wild-
type K
2P
3.1; S.E. 1.1%, n4). Chemical inhibition of glycosyl-
ation by tunicamycin resulted in an average surface expression
of 19.8% relative to wild-type K
2P
3.1 (S.E. 4.6%, n3).
Images from immunofluorescence experiments comparing
cellular localization of tunicamycin-treated and nontreated
wild-type rK
2P
3.1 channels together with mutant channels sup-
ported the flow cytometry data. When wild-type channels bear-
ing both GFP and external HA tag (GFP-rK
2P
3.1-HA) were
transiently expressed in COS-7 cells, total channel protein
expression was detected by GFP fluorescence (Fig. 2E,upper
panels,green), whereas cell surface channel was detected using
anti-HA tag antibody (red). GFP fluorescence is visible in all
four samples: untreated and tunicamycin-treated cells express-
ing either wild-type and N53Q mutant K
2P
3.1. Anti-HA stain-
ing, however, was only detected on the surface of untreated cells
expressing wild-type channel (Fig. 2E,lower panels). Taken
together these data support the conclusion that disruption of
channel glycosylation has a negative effect on cell surface
expression of rK
2P
3.1.
External glucose concentration has been reported to impact
protein glycosylation (23). We wanted to test whether reducing
external glucose concentration might modulate the surface
expression of rK
2P
3.1. Cells transiently expressing GFP-
rK
2P
3.1-HA were cultured under normal cell culture glucose
concentrations (4.5 g/liter or 25 mM) and reduced glucose (1.0
g/liter or 5.6 mM), and then flow cytometry (Fig. 3A) and con-
focal microscopy (Fig. 3B) were used to probe cell surface
expression, detected by anti-HA staining. Both methods dem-
onstrate a modest but consistent reduction in channel cell sur-
face expression. In a typical flow cytometry experiment (Fig.
3A), cell surface HA tag fluorescence in cells cultured in low
external glucose was 84% relative to control glucose concentra-
tions (4.5 g/liter); over six experiments, the mean value was
88.2 3.8% of the control. This reduced cell surface expression
in response to reduced external glucose was not detected for the
glycosylation mutant channel rK
2P
3.1
N53Q
(Fig. 3). These data
suggest that K
2P
3.1 cell surface expression is linked to glucose
concentration.
K
2P
9.1 Cell Surface Expression Is Less Sensitive to Channel
Glycosylation State—The impact of disrupting rK
2P
9.1 glyco-
sylation was investigated by examining the functional expres-
sion of both the wild-type and glycosylation mutant rK
2P
9.1
channels by patch clamp analysis, together with examining cell
surface expression of the channels by flow cytometry and
immunofluorescence. Currents evoked from HEK293 cells
transiently expressing either wild-type or glycosylation mutant
channels showed no significant difference in channel kinetics,
current amplitude, or reversal potential (Fig. 4, Aand B). Rever-
sal potential of HEK293 cells expressing rK
2P
9.1 was 67.23
1.31 mV (n13) compared with 60.00 5.78 mV (n9) for
rK
2P
9.1
N53Q
. At 60 mV test potential maximum current at pH
7.8 for wild-type channels was 2.36 0.47 nA compared with
2.91 0.56 nA for mutant channels (p0.46).
The mean surface expression level of GFP- and HA-tagged
rK
2P
9.1
N53Q
from three independent experiments was 63.6%,
S.E. 1.7, relative to the wild-type channel, when measured by
flow cytometry (Fig. 4C; MFI values as follows: GFP-rK
2P
9.1-
HA, 433; isotype control, 53; GFP-rK
2P
9.1
N53Q
HA, 287; isotype
control, 52). Although there was a clear difference in MFI, there
was considerable overlap between the range of fluorescence
intensities of cells expressing wild-type and N53Q mutant
channels. Immunofluorescence microscopy (Fig. 4D) also indi-
cated a reduction in the amount of cell surface-localized GFP-
rK
2P
9.1
N53Q
HA, compared with wild-type channel, but this
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FIGURE 2. Disruption of rK
2P
3.1 glycosylation prevents channel expression. A, electrophysiological properties of HEK293 cells transiently expressing
rK
2P
3.1 or rK
2P
3.1
N53Q
and eGFP on separate plasmids. The traces were derived from at least seven cells. , wild-type rK
2P
3.1, external pH 7.8; E, wild-type
rK
2P
3.1, external pH 6.5; ,rK
2P
3.1
N53Q
, external pH 7.8; ,rK
2P
3.1
N53Q
, external pH 6.5; f, untransfected cells, pH 7.8. B, comparison of the currents from Aat
60 mV. C, representative flow cytometric analysis of intact COS-7 cells expressing GFP-rK
2P
3.1-HA or GFP-rK
2P
3.1
N53Q
-HA. The cells were stained with either a
monoclonal antibody against the external HA tag or an isotype control antibody, followed by goat anti-mouse F(ab)
2
fragment conjugated to Alexa Fluor 647.
Solid gray curve, cells expressing GFP-rK
2P
3.1-HA, stained with the isotype control; black solid line, cells expressing GFP-rK
2P
3.1-HA and stained with anti-HA;
black dashed line, cells expressing GFP-rK
2P
3.1-HA, stained with anti-HA and treated with 1
g/ml tunicamycin; gray solid line, cells expressing GFP-
rK
2P
3.1
N53Q
HA and stained with anti-HA; gray dashed line, cells expressing GFP-rK
2P
3.1
N53Q
HA stained with anti-HA and treated with 1
g/ml tunicamycin.
D, comparison of the surface expression of GFP-rK
2P
3.1-HA and GFP-rK
2P
3.1
N53Q
HA with or without tunicamycin treatment, from a several flow cytometric
experiments. The values are percentages relative to wild-type channel in untreated cells. E, immunofluorescence images of fixed, nonpermeabilized COS-7
cells expressing GFP-rK
2P
3.1-HA (WT) or GFP-rK
2P
3.1
N53Q
HA (N53Q), with or without tunicamycin treatment. Upper panels, merged GFP (green) and anti-HA
tag-conjugated fluorescence (red). Lower panels, anti-HA tag fluorescence alone. The scale bar represents 10
m.
FIGURE 3. Surface expression of rK
2P
3.1 responds to a reduction in the concentration of glucose in the culture medium. A, flow cytometric analysis
of intact COS-7 cells expressing GFP-rK
2P
3.1-HA or GFP-rK
2P
3.1
N53Q
HA. Cells were stained with either a monoclonal antibody against the external HA tag
or an isotype control antibody, followed by goat anti-mouse F(ab)
2
fragment conjugated to Alexa Fluor 647. Solid gray curve, cells expressing GFP-
rK
2P
3.1-HA, cultured in 4.5 g/liter glucose, stained with the isotype control; black solid line, GFP-rK
2P
3.1-HA, 4.5 g/liter glucose, anti-HA; black dashed line,
GFP-rK
2P
3.1-HA, 1.0 g/liter glucose anti-HA; gray solid line: GFP-rK
2P
3.1
N53Q
HA, 4.5 g/liter glucose, anti-HA; gray dashed line: GFP-rK
2P
3.1
N53Q
HA, 1.0
g/liter glucose, anti-HA. B, confocal microscopic z-stack images of the COS-7 cells described in A, fixed and stained against HA tag (red). WT, GFP-rK
2P
3.1-
HA; N53Q, GFP-rK
2P
3.1
N53Q
HA (green fluorescence); Merge, superimposed images of total channel expression (green) and surface-exposed HA tag (red).
The scale bar represents 10
m.
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reduction was less marked in comparison with GFP-
rK
2P
3.1
N53Q
HA (Fig. 2E).
Cellular Localization of Wild-type and Glycosylation Mutant
Channels—Because the nonglycosylated rK
2P
3.1
N53Q
channel
shows marked reduction in cell surface expression compared
with the wild-type channel and because nonglycosylated pro-
teins are often retarded through the secretory pathway, we
asked whether the glycosylation mutant channel experienced
altered trafficking to the cell surface. Previous studies have
reported an ER distribution for TASK channels (7, 24). We
FIGURE 4. rK
2P
9.1 cell surface expression is less sensitive than rK
2P
3.1 to channel glycosylation state. A, currents evoked by applied membrane potential
pulses from 100 to 90 mV in HEK293 cells expressing rK
2P
9.1 (WT)orrK
2P
9.1
N53Q
(N53Q) at pH 7.8. B, average current-voltage relationship for untransfected
(UT) HEK293 cells, or HEK cells expressing rK
2P
9.1 (WT)orrK
2P
9.1
N53Q
(N53Q) at pH 7.8. C, flow cytometric analysis of intact COS-7 cells expressing GFP-
rK
2P
9.1-HA (WT) or GFP-rK
2P
9.1
N53Q
HA (HA). The cells were stained with either a monoclonal antibody against the external HA tag or an isotype control
antibody, followed by goat anti-mouse F(ab)
2
fragment conjugated to Alexa Fluor 647. Solid gray curve, cells expressing GFP-rK
2P
9.1-HA, stained with the
isotype control. Solid black line, cells expressing GFP-rK
2P
9.1-HA, stained with anti-HA tag antibody. Solid gray line, GFP-rK
2P
9.1
N53Q
HA, anti-HA-tag. Dashed gray
line, GFP-rK
2P
9.1
N53Q
HA, isotype control. Bar chart, summary of three independent experiments to determine the surface expression of GFP-rK
2P
9.1
N53Q
HA
(N53Q) to GFP-rK
2P
9.1-HA (WT). The results are expressed relative to WT (100%). The error bar shows S.E. D, immunofluorescence images of the COS-7 cells
described above, fixed and stained to detect external HA tag (red fluorescence). Upper panels, GFP-rK
2P
9.1-HA; lower panels, GFP-rK
2P
9.1
N53Q
HA (green fluores-
cence). Merge, superimposed images of channel and HA tag fluorescence. The scale bar represents 10
m.
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consistently observe a K
2P
3.1 and K
2P
9.1 perinuclear accumu-
lation in transfected cells, as well as a generalized ER staining
pattern, which co-localizes with an ER resident, protein disul-
fide isomerase. The perinuclear staining was more frequent and
pronounced in the N53Q channel mutants. To characterize the
subcellular localization of both the wild-type and glycosylation
mutant K
2P
3.1, we used markers for ERGIC (ERGIC-53) and
GC (58-kDa Golgi protein). COS-7 cells transiently express-
ing either GFP-rK
2P
3.1 or GFP-rK
2P
3.1
N53Q
showed overlap
between channel signal and each of the compartments exam-
ined (Fig. 5). To determine whether signal from the GFP-tagged
channel localized specifically with signal for each of the subcel-
lular compartments examined, the intensity of both signals
(channel, green; and compartment, red) were quantified and
compared along defined transects. This analysis revealed par-
tial overlap with the 58-kDa Golgi protein for both wild-type
and mutant channels (Fig. 5A), whereas substantial signal over-
lap was observed for both channels with ERGIC-53 (Fig. 5B).
Comparable experiments were performed for rK
2P
9.1 and
rK
2P
9.1
N53Q
with equivalent results (supplemental Fig. S1). Sig-
nificantly, both the wild-type and glycosylation mutants for
both channels were observed beyond the ER and detected in
both the ERGIC and GC compartments, providing evidence
that the lack of channel glycosylation does not completely block
forward transport of these channels.
Does Channel Stability Contribute to Reduced Function of
Glycosylation Mutant rK
2P
3.1
N53Q
?—The total expression
(both intracellular and cell surface) of GFP-tagged wild-type
rK
2P
3.1 could be compared with the GFP-tagged N53Q mutant
channel by quantitation of GFP fluorescence. In six flow cytom-
etry experiments, the total expression of GFP-rK
2P
3.1
N53Q
-HA
was reproducibly lower than the wild-type channel. In a typical
experiment (Fig. 6A), the MFI (GFP fluorescence) of cells
expressing GFP-rK
2P
3.1-HA was 24,900, whereas that of GFP-
rK
2P
3.1
N53Q
HA was 17,740. Tunicamycin treatment reduced
the MFI of cells expressing wild-type channel to 18,400, com-
pared with 16,460 for the N53Q mutant. We reasoned that
lower total amounts of nonglycosylated channel may arise
because of enhanced turnover, whether the mutant channel is
targeted for degradation direct from the ER and/or is very rap-
idly retrieved from the plasma membrane, given that
rK
2P
3.1
N53Q
is not detected at the cell surface but does proceed
through the secretory system.
Adopting a sensitive plasma membrane biotinylation
method (25), we tested whether nonglycosylated rK
2P
3.1
could be detected in newly formed endocytic vesicles. COS-7
cells expressing GFP-rK
2P
3.1 (Fig. 6B,WT panel) or GFP-
rK
2P
3.1
N53Q
(Fig. 6C,N53Q panel) underwent cell surface
biotinylation followed by a period of endocytosis. Numerous
endocytic vesicles containing biotin were visible after a
20-min incubation of surface-labeled transfected cells (Fig.
6, Band C,biotin panels). Many of the vesicles containing
biotinylated cell surface material also stained positive for the
early endosome marker EEA1 (Fig. 6, Band C,EEA1 panels).
A further subset of these vesicles was triple-stained, contain-
ing GFP-tagged wild-type or N53Q mutant channel, biotin,
and EEA1 (Fig. 6, Band C,merge panels). The number of
triple-stained vesicles for each channel was quantified, in
cells transfected with wild-type K
2P
3.1, 67 18 (n4 fields
of view) triple-stained vesicles were identified. A similar
number (77 19; n7) of triple-stained vesicles were iden-
tified in cells transfected with the glycosylation mutant
channel. Four example transects for GFP-rK
2P
3.1 and GFP-
rK
2P
3.1
N53Q
illustrate co-localization of channel, biotin, and
EEA1 in individual triple-stained vesicles (numbered 1– 4 in
Fig. 6, Band C,merge panels). These results indicate that
rK
2P
3.1
N53Q
does reach the plasma membrane and, like wild-
type, rK
2P
3.1 is retrieved in endocytic vesicles.
DISCUSSION
Glycosylation of membrane proteins is a common post-
translational modification with a variable role in the processing
and function of glycoproteins (26). In this study we examined
the prospect and impact of N-glycosylation on members of the
K
2P
family of background potassium channels with particular
focus on TASK channels (K
2P
3.1 and K
2P
9.1) and demonstrate
a conserved N-linked glycosylation site at Asn-53. Although
membrane proteins may possess an N-glycosylation consensus
sequence, it is worth noting that not all predicted sites undergo
glycan modification. This was recently demonstrated for
K
2P
18.1 (or TRESK), when two sites were predicted to undergo
N-linked glycosylation, but only one was shown to accept gly-
cans (27). The best estimate for the proportion of all proteins
glycosylated has been revised substantially downwards from
over 50% to under 20% (28), underlining the importance of
experimentally verifying predicted glycosylation sites. There-
fore, TASK channel glycosylation was verified by electrophore-
sis mobility shift assays and revealed that prevention of accept-
ance of glycosylation alters the molecular weight and hence the
mobility of these channels.
Channel glycosylation was shown to be critical for cell sur-
face expression and hence function of K
2P
3.1. Patch clamp anal-
ysis, flow cytometry, and immunofluorescence studies all verify
that K
2P
3.1
N53Q
targeting to the cell surface was disrupted. Sim-
ilarly, cells transiently expressing wild-type channels treated
with tunicamycin (which inhibits N-linked glycosylation)
showed cell surface expression comparable with cells express-
ing the glycosylation mutant channels (K
2P
3.1
N53Q
), support-
ing a regulatory role for channel glycosylation in K
2P
3.1 surface
expression and validating that the altered surface expression is due
to removal of the glycan tree rather than substitution of the Asn.
Furthermore, when cells expressing wild-type K
2P
3.1 were cul-
tured in glucose concentrations lower than standard cell culture
conditions, a 12% reduction in cell surface expression of K
2P
3.1
was detected by flow cytometry, with reduced surface expression
of the channel detected by immunofluorescence. Together, these
data provide evidence that K
2P
3.1 surface expression and function
are sensitive to the glycosylation state of the channel.
Channel glycosylation has less impact on the surface expres-
sion and function of K
2P
9.1. K
2P
9.1
N53Q
displayed a 40%
reduction in mean surface expression when compared with
the wild-type channel and quantified by flow cytometry.
Channel current for wild-type and glycosylation mutant
K
2P
9.1 channels were not significantly different when ana-
lyzed by patch clamp analyses. These data support the con-
clusion that although K
2P
9.1 is clearly glycosylated at Asn-
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FIGURE 5. Subcellular localization of GFP-tagged rK
2P
3.1 and rK
2P
3.1
N53Q
.A, COS-7 cells expressing GFP-rK
2P
3.1 (upper panels,WT,green) or GFP-
rK
2P
3.1
N53Q
(lower panels, N53Q, green) were fixed and stained with antibodies against the 58-kDa Golgi complex protein (Golgi, red). Merge, superimposed
channel and Golgi complex images. The images are single confocal sections. The white bar represents 10
m. Transects in the Merge images (yellow) reveal the
extent of co-localization of channel and the Golgi complex signals, because both signal intensities across the length of each transect are plotted in the graphs
to the right of the confocal images. B,asA, but cells were stained with an antibody against ERGIC-53.
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53, the lack of glycosylation at this site is not critical to the
channel’s surface expression and delivery and does not sig-
nificantly affect channel function but has a negative impact
on the overall level of channel maintained on the cell surface
either by altering the efficiency of channel delivery to or
removal from the cell surface.
Such a varied response of two closely related ion channels to
glycosylation is not unprecedented. Indeed, members of hyper-
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polarization-activated cyclic nucleotide-gated (HCN) channels
show similar findings. HCN1 and HCN2 are both glycosylated
in embryonic mouse heart (29), but whereas the surface expres-
sion of HCN2 is highly dependent on glycosylation, HCN1 is
markedly less sensitive. The authors propose that the channels
diverged after gene duplication from an ancestor HCN channel
that did not require glycosylation for efficient cell surface
localization.
The recently solved crystal structures of two K
2P
channels:
K
2P
1.1 and K
2P
4.1 (30, 31) were obtained using recombinant
proteins from which putative glycosylation sites had been
removed. The respective predicted glycosylation sites occur
within a disordered region after the extracellular cap/helical
cap domain and appear to be positioned away from the ion
selectivity filter in each channel. We can postulate a similar
arrangement for Asn-53 in K
2P
3.1 and K
2P
9.1, which likely
places the N-glycan away from the channel pore and selectivity
filter: in support of this, the biophysical properties of
K
2P
9.1
N53Q
did not appear to be altered when examined by
whole cell patch clamp recordings.
Among their other roles, glycans have been proposed to act
as targeting determinants for a number of channels, includ-
ing aquaporins (AQP-2), voltage-gated potassium channels
(Kv1.2), HCN channels, and acid-sensing ion channels (ASIC-
1a) (29, 32–34). It is difficult, however, to discriminate between
sorting per se and the protein quality control processes that
precede targeting. The
-subunit of gastric HK-ATPase is an
example of a protein where N-linked glycans have been shown
to play a distinct role in targeting to the apical plasma mem-
brane (35). The almost complete absence of K
2P
3.1
N53Q
on the
plasma membrane led us to investigate whether intracellular
localization of the glycosylation mutant channel was disrupted.
When the intracellular localization of wild-type K
2P
3.1 and
mutant K
2P
3.1
N53Q
were compared by immunofluorescence,
both channels were detected in both the ERGIC and the GC.
Indeed, although a degree of overlap between the GC marker
and the channel was detected, substantially greater overlap
between ERGIC marker and both channels was observed. Sig-
nificantly, ERGIC contributes to the concentration, folding,
and quality control of newly synthesized proteins. These find-
ings are significant because they suggest that although the non-
glycosylated channel is not detected on the cell surface, it does
escape the ER and traffics to the ERGIC and GC. Therefore it
appears that glycosylation does not in itself alter the processing
pathway of K
2P
3.1 channels.
If glycosylation mutant channels are not retained within the
ER and pass through the secretory pathway, these channels
have the possibility of two fates. Mutant channels may reach the
cell surface but show lower stability and hence a higher rate of
turnover, or they may be targeted directly for degradation.
Because N-linked glycosylation of K
2P
3.1 channels does not
appear to alter intracellular transport of the channel, we sought
to determine whether glycosylation had an impact on channel
turnover. By flow cytometry, a reduction in the total (intracel-
lular and cell surface) amount of K
2P
3.1
N53Q
per cell compared
with the wild type was observed. Similarly, tunicamycin-treated
cells expressing K
2P
3.1 showed a similar reduction, with total
channel per cell comparable with ablation of the glycosylation
site. These observations are significant when considered
together with our recycling data. When channel retrieval from
the cell surface and entry into the endocytic pathway is studied,
although K
2P
3.1
N53Q
is not detected on the cell surface,
K
2P
3.1
N53Q
which was tagged by surface labeling with biotin
(and therefore must have reached the cell surface) is detected
within the endocytic pathway. Together these data suggest that
although K
2P
3.1 cannot be detected at the cell surface, a pro-
portion of the channel population does reach the plasma mem-
brane but is retrieved and likely targeted for degradation via the
endocytic pathway. These studies provide strong evidence that
channel glycosylation plays an important role in the stability of
K
2P
3.1 expression on the cell surface.
How Might Glycosylation Contribute to the Stability of K
2P
3.1
Once Delivered to the Plasma Membrane?—The two glycans on
wild-type CFTR have been shown to promote proper apical
recycling; in their absence, CFTR is more rapidly internalized
and is targeted to the basolateral membrane, where it appears to
have a shorter half-life than glycosylated CFTR (36). The same
glycans also play a role in the stability of the channel once it has
exited the ER (36). A subset of the N-glycans attached to the
HK-ATPase
is responsible for delivery of the protein to the
apical membrane and retaining it there (35). It has been sug-
gested that extracellular lectins may play a role in binding HK-
ATPase and stabilizing it at the cell surface (37). Glycosylation
increases thermodynamic stability by reducing the amount of
surface area accessible to solvent, which in turn influences
structural dynamics and protein function (26); this effect has
been clearly demonstrated for the thermostability of human
aquaporin 10 protein (39). Although a number of potential
mechanisms exist, it will be important to find out how glycosyl-
ation contributes to the stability of K
2P
3.1 and to determine the
process that renders K
2P
9.1 less sensitive to this regulation.
TASK channels are expressed in both neuronal and cardiac
cell populations and significantly have been identified in glu-
cose-sensing neurons of the hypothalamus, as well as periph-
eral specialized chemo- and nutrient-sensing cells (40, 41). Sen-
sitivity of K
2P
3.1 cell surface expression and turnover to its
glycosylation state represents a potential link between meta-
bolic status and cellular activity. Down-regulation of TASK
channels is known to influence cellular depolarization. Hence,
decreased cell surface expression of K
2P
3.1 channels in
FIGURE 6. Is reduced function of glycosylation mutant rK
2P
3.1
N53Q
caused by lower channel stability? A, flow cytometric analysis of COS-7 cells expressing
GFP-rK
2P
3.1-HA without (solid black line) and in the presence of 1
g/ml tunicamycin (dashed black line) or GFP-rK
2P
3.1
N53Q
HA without (solid gray line) or with
tunicamycin (dashed gray line). The population of GFP-positive cells is denoted by Transfected cells, derived by comparing with cells transfected with empty
vector alone (data not shown). B,upper panels, confocal microscopic images of COS-7 cells transfected with GFP-rK
2P
3.1 (WT,green), surface biotinylated (Biotin,
red), and then allowed to endocytose for 20 min before fixing and staining with anti-EEA1 (EEA1, blue). Merge, superimposed images of channel, biotin and
EEA1. Examples of triple-stained vesicles are denoted with white arrows, and the numbers correspond to transects (transects themselves not shown on the
Merge image), which appear in the lower panels 1– 4, as graphs of fluorescence intensity against distance. The white scale bar represents 10
m. C, as described
for B, except cells express GFP-rK
2P
3.1
N53Q
(N53Q, green).
Glycosylation of TASK Channels
3262 JOURNAL OF BIOLOGICAL CHEMISTRY VOLUME 288 •NUMBER 5 •FEBRUARY 1, 2013
by guest on February 14, 2017http://www.jbc.org/Downloaded from
response to decreased glucose would lead to increased neuronal
excitation. Furthermore, in the diabetic patient, with sustained
higher blood glucose levels, one would predict that this envi-
ronment would promote K
2P
3.1 channel surface expression
with resultant dampening of cellular activity.
The importance of the findings presented in this study lies in the
numerous roles TASK channels may play in cellular regulation.
Their varied sensitivity and stability to glycosylation, and by asso-
ciation glucose concentration, opens a host of potential regulatory
pathways in which these important channels may be involved.
Acknowledgments—We are grateful to Dr. David Johnston of the Uni-
versity of Southampton Biomedical Imaging Unit for assistance with
confocal microscopy, Kelly Wilkinson for expert technical support,
and Prof. Stephen High at the University of Manchester for helpful
comments.
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Glycosylation of TASK Channels
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Merge
Merge
N53Q
WT ERGIC
ERGIC
WT
ERGIC
03.6 µm
N53Q
ERGIC
03.6 µm
Supplementary Figure 1
A
B
WT
ERGIC
N53Q
ERGIC
Fluorescence intensityFluorescence intensityFluorescence intensityFluorescence intensity
C
0
0
7.0 µm
7.0 µm
WT
Golgi
WT
Golgi
N53Q
Golgi
N53Q
Golgi
Subcellular localisation of rK 9.1 and
2P
rK 9.1
2P N53Q
A: Fluorescence images of COS-7 cells
transfected with GFP-rK 9.1 (WT, green,
2P
upper panels) or GFP-rK 9.1 (N53Q,
2P N53Q
green, lower panels), counter-stained
with an antibody against the 53 kDa
ERGIC protein (ERGIC, red). Merge:
superimposed images of channel and
ERGIC, showing areas of signal overlap.
White scale bar: 10 µm.
B: As in panel A, Merge, but single
confocal sections. Yellow arrows point to
transects shown in the corresponding
graphs of fluorescence intensity to the
right of each image. White scale bars: 10
µm.
C: As in panel B, but the cells were
counter-stained with an antibody against
the 58 kDa Golgi protein, instead of
ERGIC.
Ita O'Kelly
Alexandra Mant, Sarah Williams, Laura Roncoroni, Eleanor Lowry, Daniel Johnson and
9.1
2P
3.1 and K
2P
Potassium Channels K
-Glycosylation-dependent Control of Functional Expression of BackgroundN
doi: 10.1074/jbc.M112.405167 originally published online December 18, 2012
2013, 288:3251-3264.J. Biol. Chem.
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This article cites 40 references, 11 of which can be accessed free at
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... Furthermore, in several voltage-gated ion channels, negatively charged sialic acid residues located at the glycan structures are thought to contribute to the external negative surface potential, thereby impacting channel gating through electrostatic mechanisms [35][36][37]. While N-linked carbohydrate modifications have been described to alter the functional characteristics of the K 2P channels hK 2P 3.1 (hTASK-1, TWIK-related acid-sensing K + channel 1), hK 2P 9.1 (hTASK-3, TASK-3, TWIK-related acid-sensing K + channel 3) and hK 2P 18.1 (TRESK, TWIK-related spinal cord K + channel) [38,39], it remains uncertain whether other members of the K 2P channel family are subjected to N-glycosylation in a similar fashion. ...
... With the exception of hK2P15.1 (hTASK-5), a channel with unclear functional relevance, all members of the K2P channel family carry at least one putative N-glycosylation motif in their M1-P1 interdomain [38]. While N-glycosylation of the channels hK2P1.1 (hTWIK-1), K2P3.1 (TASK-1), K2P9.1 (TASK-3) and K2P18.1 (TRESK) has been experimentally validated [38,39,47,48], relatively little is known about the glycosylation of the remaining K2P channels. ...
... With the exception of hK2P15.1 (hTASK-5), a channel with unclear functional relevance, all members of the K2P channel family carry at least one putative N-glycosylation motif in their M1-P1 interdomain [38]. While N-glycosylation of the channels hK2P1.1 (hTWIK-1), K2P3.1 (TASK-1), K2P9.1 (TASK-3) and K2P18.1 (TRESK) has been experimentally validated [38,39,47,48], relatively little is known about the glycosylation of the remaining K2P channels. Functional validation of putative Nglycosylation motifs, however, is indispensable as Egenberger et al. [39] showed that TRESK harbors two putative N-glycosylation motifs, but only one is of functional relevance. ...
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... Disruption of TASK1 and TASK3 glycosylation lowers the number of cell surface channels, which leads to a reduced current flow. 28 Similarly, inhibiting N-glycosylation of Kv1.4 channel decreases protein stability, induces intracellular retention, and decreases cell surface protein levels. 29 Thus, enhanced N-glycosylation observed in APA may promote steroidogenesis via increasing cell surface expression and activity of relevant receptors and ion channels. ...
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... PCR mutagenesis was carried out using Pfu Ultra DNA polymerase (Agilent Technologies, Stockport, UK) and the following thermo cycle: 2 min at 95°C, (12× cycles of 1 min at 95°C, 1 min at 55°C, 7.5 min at 68°C), 10 min at 68°C. 17 DNA was digested with Dpn1 for 1 hour at 37°C to cleave the parental DNA template. The mutated vector was sequenced to confirm the AMMECR1 mutation (Source BioScience). ...
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Background Deletions in the Xq22.3–Xq23 region, inclusive of COL4A5, have been associated with a contiguous gene deletion syndrome characterised by Alport syndrome with intellectual disability (Mental retardation), Midface hypoplasia and Elliptocytosis (AMME). The extrarenal biological and clinical significance of neighbouring genes to the Alport locus has been largely speculative. We sought to discover a genetic cause for two half-brothers presenting with nephrocalcinosis, early speech and language delay and midface hypoplasia with submucous cleft palate and bifid uvula. Methods Whole exome sequencing was undertaken on maternal half-siblings. In-house genomic analysis included extraction of all shared variants on the X chromosome in keeping with X-linked inheritance. Patient-specific mutants were transfected into three cell lines and microscopically visualised to assess the nuclear expression pattern of the mutant protein. Results In the affected half-brothers, we identified a hemizygous novel non-synonymous variant of unknown significance in AMMECR1 (c.G530A; p.G177D), a gene residing in the AMME disease locus. Transfected cell lines with the p.G177D mutation showed aberrant nuclear localisation patterns when compared with the wild type. Blood films revealed the presence of elliptocytes in the older brother. Conclusions Our study shows that a single missense mutation in AMMECR1 causes a phenotype of midface hypoplasia, mild intellectual disability and the presence of elliptocytes, previously reported as part of a contiguous gene deletion syndrome. Functional analysis confirms mutant-specific protein dysfunction. We conclude that AMMECR1 is a critical gene in the pathogenesis of AMME, causing midface hypoplasia and elliptocytosis and contributing to early speech and language delay, infantile hypotonia and hearing loss, and may play a role in dysmorphism, nephrocalcinosis and submucous cleft palate.
... Consistent with our previous study showing a glucose-dependent potentiation of the T-type calcium conductance [37], our data revealed that elevated external glucose concentration caused a significant increase of the expression of the channel protein at the cell surface, without affecting the total expression of the channel. This observation is consistent with previous results obtained with two-pore domain potassium (K2P) channels showing that elevated glucose levels potentiate surface expression of the channel [17]. In contrast, surface expression of the glycosylation-deficient N192Q/N1466Q mutant channel was not influenced by external glucose levels, suggesting that glucosedependent potentiation of hCa v 3.2 channels requires the presence of the glycan tree on the channel protein. ...
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T-type calcium channels are key contributors to neuronal physiology where they shape electrical activity of nerve cells and contribute to the release of neurotransmitters. Enhanced T-type channel expression has been causally linked to a number of pathological conditions including peripheral painful diabetic neuropathy. Recently, it was demonstrated that asparagine-linked glycosylation not only plays an essen- tial role in regulating cell surface expression of Cav3.2 chan- nels, but may also support glucose-dependent potentiation of T-type currents. However, the underlying mechanisms by which N-glycosylation and glucose levels modulate the ex- pression of T-type channels remain elusive. In the present study, we show that site-specific N-glycosylation of Cav3.2 is essential to stabilize expression of the channel at the plasma membrane. In contrast, elevated external glucose concentra- tion appears to potentiate intracellular forward trafficking of the channel to the cell surface, resulting in an increased steady-state expression of the channel protein at the plasma membrane. Collectively, our study indicates that glucose and N-glycosylation act in concert to control the expression of Cav3.2 channels, and that alteration of these mechanisms may contribute to the altered expression of T-type channels in pathological conditions.
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