Tài liệu Báo cáo khoa học: The role of N-glycosylation in the stability, trafficking and GABA-uptake of GABA-transporter 1 Terminal N-glycans facilitate efficient GABA-uptake activity of the GABA transporter pptx

The role of N-glycosylation in the stability, trafficking and
GABA-uptake of GABA-transporter 1
Terminal N-glycans facilitate efficient GABA-uptake activity
of the GABA transporter
Guoqiang Cai
1,2
, Petrus S. Salonikidis
3
, Jian Fei
1
, Wolfgang Schwarz
3
, Ralf Schu
¨
lein
4
,
Werner Reutter
2
and Hua Fan
2
1 Institute of Biochemistry and Cell Biology, SIBS, CAS, Shanghai, China
2 Institut fu
¨
r Molekularbiologie und Biochemie, CBF, Charite
´
Universita
¨
tsmedizin Berlin, Berlin-Dahlem, Germany
3 Max-Planck Institut fu
¨

r Biophysik, Frankfurt, Germany
4 Forschungsinstitut fu
¨
r Molekulare Pharmakologie, Berlin-Buch, Germany
The cellular membrane transporter for the inhibitory
neurotransmitter c-aminobutyric acid (GABA) belongs
to a family of secondary active systems that are driven
by electrochemica1 gradients of Na
+
and Cl
–
[1]. The
main physiological function of the transporter is
believed to be the control of the concentration and
dwell time of GABA in the synaptic cleft. Because the
transport of one molecule of GABA is coupled to the
Keywords
GABA transporter; N-glycosylation; N-glycan
trimming; membrane trafficking; patch-
clamp
Correspondence
H. Fan, Institut fu
¨
r Molekularbiologie und
Biochemie, Campus Bejamin Franklin,
Charite
´
Universita
¨
tsmedicin Berlin,

Arnimallee 22, D-14195 Berlin-Dahlem,
Germany
Fax: +49 30 84451541
Tel: +49 30 84451544
E-mail:
(Received 17 July 2004, revised 24 January
2005, accepted 2 February 2005)
doi:10.1111/j.1742-4658.2005.04595.x
Neurotransmitter transporters play a major role in achieving low concen-
trations of their respective transmitter in the synaptic cleft. The GABA
transporter GAT1 belongs to the family of Na
+
- and Cl
–
-coupled trans-
port proteins which possess 12 putative transmembrane domains and three
N-glycosylation sites in the extracellular loop between transmembrane
domain 3 and 4. To study the significance of N-glycosylation, green fluor-
escence protein (GFP)-tagged wild type GAT1 (NNN) and N-glycosylation
defective mutants (DDQ, DGN, DDN and DDG) were expressed in CHO
cells. Compared with the wild type, all N-glycosylation mutants showed
strongly reduced protein stability and trafficking to the plasma membrane,
which however were not affected by 1-deoxymannojirimycin (dMM). This
indicates that N-glycosylation, but not terminal trimming of the N-glycans
is involved in the attainment of a correctly folded and stable conformation
of GAT1. All N-glycosylation mutants were expressed on the plasma mem-
brane, but they displayed markedly reduced GABA-uptake activity. Also,
inhibition of oligosaccharide processing by dMM led to reduction of this
activity. Further experiments showed that both N-glycosylation mutations
and dMM reduced the V

max
value, while not increasing the K
m
value for
GABA uptake. Electrical measurements revealed that the reduced transport
activity can be partially attributed to a reduced apparent affinity for extra-
cellular Na
+
and slowed kinetics of the transport cycle. This indicates that
N-glycans, in particular their terminal trimming, are important for the
GABA-uptake activity of GAT1. They play a regulatory role in the GABA
translocation by affecting the affinity and the reaction steps associated with
the sodium ion binding.
Abbreviations
CHO, Chinese hamster ovary; dMM, 1-deoxymannojirimycin; ER, endoplasmic reticulum; FACS, fluorescence activated cell sorting; GABA,
c-aminobutyric acid; GAT1, GABA transporter type I; GFP, green fluorescence protein.
FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS 1625
cotransport of two Na
+
ions and one Cl
–
ion [2–4],
the translocation across the membrane is associated
with a current that can be measured by voltage clamp-
ing. In the absence of GABA the transport cycle is
not completed, but transient charge movements can be
detected that reflect partial reactions associated with
extracellular Na
+
binding and hence provide kinetic

information about the transport cycle [5–7].
Four subtypes of GABA transporters (GAT1–4)
have been found so far [8,9]. GABA transporter type
1 (GAT1) is a single polypeptide of about 67 kDa
with 12 putative transmembrane domains. Both
N- and C-termini are located in the cytoplasm. The
large extracellular loop between transmembrane
domains 3 and 4 contains three conserved N-glyco-
sylation sites (Asn176, Asn181 and Asn184). It has
been demonstrated that all three N-glycosylation sites
are used in vivo and that no additional sites are pre-
sent [10].
N-glycosylation is a major post-translational modi-
fication in eukaryotic cells. Recent results suggest
that this post-translational modification may influence
many of the physicochemical and biological proper-
ties of the proteins, such as protein folding, stability,
targeting, dynamics and ligand binding, as well as
cell-matrix and cell–cell interactions [11–16]. It has
been suggested that N-glycosylation is involved in
the regulation of the transport activity and surface
expression of neurotransmitter transporters [10,17].
Functional expression of the GABA transporter is
abolished by tunicamycin, a potent inhibitor of
N-glycosylation [18]. Experiments with HeLa trans-
fectants showed that removal of one or two glycosy-
lation sites by site-directed mutagenesis had little
effect on the expression of GABA-uptake activity.
However, removal of all three N-glycosylation sites
resulted in a reduction of GABA-uptake activity [10].

Although such experiments indicate that N-glycosyla-
tion mutations lead to a reduction of GABA-uptake
activity, we do not know how the N-linked oligosac-
charide side chains influence the function of this trans-
porter. Liu et al. demonstrated in Xenopus oocytes
that mutations of two of the three N-glycosylation
sites led to a reduction in turnover rates and complex
changes in the interaction of external Na
+
with the
transport protein as measured by voltage clamping [7].
However, the question remained as to whether the
reduction in function of the mutants was due to a
change in the biochemical properties of this transpor-
ter or to a reduction in the number of GABA trans-
porters per cell. In order to clarify the functional
significance of N-glycosylation and N-linked oligosac-
charides in GAT1, green fluorescence protein (GFP)-
tagged wild type GAT1 (NNN) and four glycosylation
mutants (DDQ, DGN, DDN, and DND) were stably
expressed in CHO cells lacking endogenous GAT1.
The influence of N-glycosylation mutations and inhibi-
tion of N-glycosylation processing on biochemical
properties and function were investigated.
In this work, we demonstrated that defective N-gly-
cosylation resulted in reduction of the stability and a
decrease in the cell surface expression of this protein.
The GAT1 mutants containing two N-glycosylation
mutations showed a delayed intracellular translocation,
but they targeted to the plasma membrane and showed

reduced GAT1-specific GABA-uptake activity. If all
three N-glycosylation sites were eliminated, a decreased
percentage of DDQ mutants was found on the cell
surface. However, the GABA-uptake activity could
hardly be detected in this mutant. Inhibition of
N-glycosylation processing by 1-deoxymannojirimycin
(dMM) affected neither the cell surface expression nor
stability of this protein, but it resulted in marked reduc-
tion of GABA-uptake activity. This suggests that
N-glycans, in particular terminal structures of N-gly-
cans, are involved in the GABA-uptake process of
GAT1. Finally, we found that deficiency of N-glycosy-
lation did not affect the affinity of GAT1 for GABA.
The observed reduction of GAT1-specific GABA-
uptake due to deficiency of N-glycans was attributed to
a reduced apparent affinity for extracellular Na
+
ions,
resulting in a reduction of the kinetics of the transport
cycle.
Results
Expression of GAT1/GFP fusion proteins
in CHO cells
cDNAs of GFP tagged wild type (NNN) and mutants
DND, DDN, DGN and DDQ were transfected into
CHO cells, which do not express endogenous GAT1
and GFP. Stable transfectants were selected by fluores-
cence activated cell sorting (FACS). Flow cytometry
analysis showed the expression of NNN and the
mutants on the surface of transfected CHO cells

(Fig. 1A). Fluorescence and immunofluorescence micro-
scopy showed that both GFP-fluorescence and anti-
GAT1 antibodies can be used to detect the expression
of the GAT1 ⁄ GFP fusion protein (NNN) on surface
and interior of CHO cells (Fig. 1B,C).
The expression of NNN was determined by West-
ern blotting with either anti-GAT1 pAb (Fig. 2A) or
anti-GFP mAb (Fig. 2B) following immunoprecipita-
tion with anti-GFP pAb. This GAT1 ⁄ GFP fusion
protein showed several bands in SDS ⁄ PAGE, two
Role of N-glycosylation and N-glycan trimming of GAT1 G. Cai et al.
1626 FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS
monomeric forms running as a main band of about
108 kDa and a small band of about 96 kDa. The
108 kDa polypeptide was resistant to Endo H diges-
tion, while the 96 kDa polypeptide was converted
into a polypeptide of 90 kDa after digestion with
Endo H (Fig. 2B). Digestion of both monomeric
forms with PNGase F resulted in a single 90 kDa
N-glycan-free peptide (Fig. 2C, lane 2). This indicates
that the 108 kDa peptide contains mature N-glycans
of the complex type, while the 96 kDa peptide con-
tains only N-glycans of the mannosidic type. The
210 kDa band may represent a dimeric form or a
protein aggregate. In addition, inhibition of N-glyco-
sylation processing of NNN by 1-deoxymannojirimy-
cin (dMM) leads to the reduction of NNN molecule
mass to 96 kDa (Fig. 2C, lanes 3 and 6). After diges-
tion with either PNGase F or Endo H, this 96 kDa
polypeptide was converted to a 90 kDa N-glycan-free

polypeptide (Fig. 2C, lanes 4 and 5), indicating that
the 96 kDa polypeptide contains only N-glycans of
oligomannosidic type.
Fig. 1. Flow cytometry, fluorescence microscopy and immunofluo-
rescence microscopy of GFP-tagged GAT1 in transfected CHO
cells. (A) Flow cytometry of GFP-tagged GAT1 wild type and
mutants. The polyclonal anti-GAT1 IgG was used for immunostain-
ing. Visualization was performed with R-phycoerythrin-conjugated
goat anti-(rabbit IgG) Ig. NNN, GFP-tagged wild type GAT1. DND,
DDN, DGN and DDQ, GFP-tagged N-glycosylation mutants. (B)
Fluorescence microscopy of NNN. The fluorescence of GFP in
GFP ⁄ GAT-fusion protein (NNN) was detected. (C) Immunofluores-
cence microscopy of NNN. Anti-GAT1 polyclonal antibodies were
used for immunostaining after cell fixation and permeabilization.
Visualization was performed with R-phycoerythrin-conjugated goat
anti-(rabbit IgG) Ig.
Fig. 2. Protein expression and N-glycosylation processing of GFP-
tagged GAT1 in CHO cells. NNN stable transfected CHO cells were
incubated with and without dMM (1 m
M) for 72 h. The solubilized
protein of transfected cells (1 · 10
7
) was subjected to immunopre-
cipitation with anti-GFP Igs. Aliquots of each immunoprecipitate
were treated either with Endo H or PNGase F. The resulting mix-
ture and the other aliquots of the immunoprecipitate were analyzed
by SDS ⁄ PAGE (7.5%) and immunoblotting with anti-GAT1 pAb (A)
or anti-GFP mAb (B, C).
G. Cai et al. Role of N-glycosylation and N-glycan trimming of GAT1
FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS 1627

Expression of N-glycosylation mutants
on the plasma membrane of CHO cells
The expression of NNN and N-glycosylation mutants
DGN, DDN, DND and DDQ on the plasma membrane
of CHO cells was investigated. As shown in Fig. 3A,
N-glycosylation mutants exhibited a reduced molecular
mass in comparison to that of the wild type NNN.
Nevertheless, all N-glycosylation mutants, as well as the
wild type NNN, were expressed on the plasma mem-
brane. Although all three N-glycosylation sites are
absent in DDQ, this mutant was also detected on the
plasma membrane of CHO cells, suggesting that N-gly-
cosylation or N-linked oligosaccharides are important,
but not essential for the translocation of GAT1 to the
cell surface. All intracellular proteins of wild type as
well as mutants (with the exception of DDQ) gave two
bands, while plasma membrane proteins gave only one
large band (Fig. 3A). On the basis of the Endo H diges-
tion (Fig. 2B,C), the large band is assigned to proteins
with N-glycans of mature complex type and the small
band to the proteins with N-glycans of mannose-rich
type. All bands of the mutants had reduced molecular
mass, compared with that of wild type NNN. The
reduced molecular masses of mutants are compatible
with the absence of N-glycans at the two eliminated
N-glycosylation sites, suggesting that the mutants in the
cell interior contain N-glycans of both mannosidic and
complex types, while those in the plasma membrane
contain only N-glycans of the mature complex type.
The relative levels of surface vs. intracellular GAT1

and mutants in a steady expression state were quanti-
fied. The distributions between cell surface and cell
interior of the mutants DGN, DND and DDN were
not significantly different from that of wild type NNN.
About 46 ± 4.7% is found on the cell surface. How-
ever, the percentage of the cell surface expression in
mutant DDQ which lacks all three N-glycosylation
sites was only 30 ± 4.4% in the steady expressed state
(Fig. 3B).
N-Glycosylation mutations result in reduction
of GABA-uptake activity
For quantitative measurement of the specific activity
of GABA-uptake, an aliquot of the stable CHO trans-
fectants was used for the GABA-uptake assay, and
another aliquot was used to determine the amount of
the membrane-expressed wild type or mutant proteins.
The GABA-uptake activities were normalized to the
same amount of cell surface proteins of wild type and
mutants. Compared with that of the wild type, the
GABA-uptake activities of the N-glycosylation
mutants were reduced significantly. Figure 3C shows
that the GABA-uptake activities of mutants with
double N-glycosylation mutations, DND, DGN and
Fig. 3. Determination of expression of GFP-tagged GAT1 mutants
on the surface of transfected CHO cells and measurement of
GABA-uptake by GFP-tagged GAT1 wild type and mutants in trans-
fected CHO cells. (A) Cell surface and intracellular expression of
GFP-tagged GAT1 wild type (NNN) and mutants (DGN, DND, DDN,
and DDQ) were analyzed by biotin labelling and Western blotting.
Anti-GAT1 serum or anti-GFP mAb MAB2510 were used for immu-

nostaining. I, intracellular expression; M, plasma membrane expres-
sion. (B) The protein bands obtained in western blotting were
analyzed by phosphoimager scanning. Each value represents the
mean ± SEM of three separate experiments. The total protein of
the cell surface and intracellular bands of each wild type or mutant
were set at 100%. (C) Measurement of GABA-uptake by GFP-
tagged GAT1 wild type and mutants in transfected CHO cells. The
measured GABA-uptake activity was normalized to the amount of
GAT1 or mutant protein expressed on the plasma membrane. The
activity of GABA-uptake by NNN was set at 100%. All other values
were expressed relative to this value. The values represent the
mean ± SEM of four separate experiments.
Role of N-glycosylation and N-glycan trimming of GAT1 G. Cai et al.
1628 FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS
DDN, were reduced to 64% (±5.6%), 42% (±12.4%)
and 32% (±8.2%) of that of NNN, respectively.
GAT1-mediated transport could hardly be detected in
the mutant DDQ, although this mutant was expressed
on the plasma membrane. Mutant DDQ does not exhi-
bit any N-glycosylation site. Because all values were
normalized to the transporter proteins in the plasma
membrane, the reduced specific activities of the
mutants are not due to a reduced number of GABA
transporters per cell, but to a reduced transport rate.
This suggests that N-linked oligosaccharide side-chains
are important for the GABA transport activity.
1-Deoxymannojirimycin inhibits the GABA-uptake
of GAT1
In order to gain further insight into the role of the
terminal structures of the N-glycans of GAT1, N-gly-

cosylation processing of NNN was inhibited by 1-de-
oxymannojirimycin (dMM). Inhibition by dMM leads
to the formation of NNN molecules containing N-gly-
cans of oligomannosidic type. Figure 4A shows that
after treatment with dMM (1 mm) for 72 h, the
amount of plasma membrane NNN containing man-
nosidic N-glycans was in the same range as that of
NNN containing mature complex N-glycans without
treatment with dMM. However, the activity of GABA-
uptake was reduced to 37% after treatment with dMM
(Fig. 4B). This indicates that the terminal trimming of
N-oligosaccharides is not involved in the regulation of
plasma membrane trafficking of GAT1, but in the
regulation of GABA uptake.
As well as wild type, mutant DND, DGN and
DDN exhibited only one small band on SDS ⁄ PAGE
after treatment with dMM (data not shown), indica-
ting that, like wild type, they contain only mannosidic
N-glycans. The level of cell surface expression was sim-
ilar with and without dMM treatment for both wild
type and mutants (Figs 4A and 5A). However, their
GABA-uptake activity was reduced to half after treat-
ment with dMM (1 mm) for 48 h (Fig. 5B). Although
mutant DND, DGN and DDN contain only one
N-glycosylation site, deficiency of terminal trimming of
their N-oligosaccharides strongly affected their GABA-
uptake activity. These indicate that the terminal
structure of the oligosaccharides facilitate efficient
GABA-uptake activity of the GABA transporter.
Defective N-glycosylation results in reduction

of the stability of GAT1
In order to study the influence of the N-glycosylation
and N-linked oligosaccharides on protein stability, the
intracellular decay time of the GAT1, GAT1 treated
with dMM, and the mutants was determined by pulse-
chase experiments (Fig. 6). Figure 6B shows that wild
type NNN was very stable with an exponential half-life
of about 22 h, whereas the half-life of mutant DDN
containing two N-glycosylation mutations was reduced
to 12 h. The half-life of DDQ containing all three
N-glycosylation mutations was reduced even more,
compared with that of DDN, showing a value of only
5.5 h. In contrast, the stability of NNN containing
only mannosidic N-glycans after treatment with dMM
Fig. 4. Influence of dMM on plasma membrane trafficking and
GABA-uptake of GFP-tagged GAT1 wild type. NNN stable trans-
fected CHO cells were incubated with and without dMM (1 m
M)
for 72 h. (A) Aliquots of cells were used for membrane biotinyla-
tion. After solubilization, 300 lg total proteins of cell lysates were
precipitated with streptavidin beads. The eluates were analyzed by
Western blotting using anti-GFP mAb. (B) Another aliquot of cells
was used for measurement of GABA-uptake as described above.
The values represent the mean ± SEM of three separate experi-
ments.
G. Cai et al. Role of N-glycosylation and N-glycan trimming of GAT1
FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS 1629
is similar to that of NNN containing N-glycans of the
mature complex type. The results suggest that N-glyco-
sylation is important for the stability of this protein,

but the terminal structure of the N-glycans is not.
Defective N-glycosylation reduces the trafficking
of GAT1 to the plasma membrane
In order to study the influence of N-glycosylation on
plasma membrane trafficking of GAT1, the distribu-
tion of wild type and mutants on the cell surface and
in the cell interior was kinetically analyzed by pulse-
chase experiments. Figure 7 shows that after a 40 min
chase, 34% of total wild type (NNN) proteins, whereas
only 18 and 12% of total mutant DDN and DDQ
proteins, respectively, were expressed on the plasma
membrane. After a 120-min chase, the membrane
expression of the NNN was increased to 50%, whereas
that of mutant DDN and DDQ was increased only to
40% and 15%, respectively. This result suggests that
deficiency of N-glycosylation impairs the plasma mem-
brane trafficking of GAT1.
Defective N-glycosylation or dMM treatment did
not increase the K
m
GABA values of GAT1
The above results show that both N-glycosylation
mutations and terminal structures of N-linked oligo-
saccharide side chains have a measurable effect on the
GABA-uptake activity of GAT1. To determine whe-
ther the N-linked oligosaccharide side chains of GAT1
influence the affinity of GAT1 for GABA, concentra-
tion dependencies were analyzed on the basis of the
Michaelis–Menten equation
V ¼ V

max
GABA
½GABA
K
m
GABA þ½GABA
and the parameters for NNN with and without treat-
ment with dMM, and for N-glycosylation mutant
DDN were determined. As shown in Fig. 8, the V
max
Fig. 5. Analysis of the cell surface expres-
sion and GABA-uptake activity of GFP-tag-
ged GAT1 wild type and mutants after
treatment with dMM. CHO stable transfect-
ants were incubated with and without dMM
(1 m
M) for 48 h (A) Cell surface expression
by FACS analysis. Anti-GAT IgG was used
for the immunostaining. Visualization was
performed with R-phycoerythrin-conjugated
goat anti-(rabbit IgG) Ig. (B) GABA-uptake
activity. The GABA-uptake activities were
normalized to the amount of GAT1 or muta-
nt protein expressed on the plasma mem-
brane. The activity of GABA-uptake by NNN
was set at 100%. The values represent the
mean ± SEM of five separate experiments.
Role of N-glycosylation and N-glycan trimming of GAT1 G. Cai et al.
1630 FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS
GABA values of NNN treated with dMM, and of

mutant DDN were reduced significantly. The V
max
GABA value of NNN without dMM is 1.21
pmolÆlgÆprotein
)1
Æmin
)1
, whereas the value for mutant
DDN was only 0.29 pmolÆlgÆprotein
)1
Æmin
)1
. After
treatment of NNN with dMM, the V
max
GABA value
of NNN containing mannose-rich N-glycans was
strongly reduced to 0.55 pmolÆlgÆprotein
)1
Æmin
)1
.
Although mutations at N-glycosylation sites, as well as
N-glycans of the oligomannosidic type reduced the
V
max
value of rate of GABA uptake markedly, the K
m
GABA values were not affected. The data in Fig. 8
were fitted with a common K

m
GABA value of 4.1 lm.
These results suggest that the defect of N-linked oligo-
saccharides did not reduce the binding affinity of
GAT1 to GABA. As treatment with dMM did not
affect GAT1 protein translocation to the plasma mem-
brane, the decreased GABA-uptake activity of NNN
Fig. 6. Biological stability of GFP-tagged GAT1 wild type and
mutants in transfected CHO cells. (A) CHO stable transfectants
(2 · 10
6
cells per dish) were preincubated with and without dMM
(1 m
M) for 72 h, then pulse-labelled with 3.7 · 10
6
Bq per dish
[
35
S]methionine for 1 h and immediately chased for the stated
times. Immunoprecipitates of cell lysates obtained at the indicated
chase-times were analyzed by SDS ⁄ PAGE. (B) The results of the
pulse-chase experiments were analyzed by phosphoimager scan-
ning. The radioactivities obtained by immunoprecipitation of the
pulse-labelled cells without chase were set at 100%. All other val-
ues were expressed relative to this value. Each time point repre-
sents the mean ± SEM of three separate experiments. Solid lines
represent the exponential fit with half-lives of 22 and 23 h for
wild type GAT1 in the absence and presence of dMM, respectively,
and of 12 and 5.5 h for the DDN and DDQ mutants, respectively.
Fig. 7. Plasma membrane trafficking of GFP-tagged GAT1 wild type

and mutants in transfected CHO cells. (A) CHO stable transfectants
were pulse-labelled with 3.7 · 10
6
Bq per dish [
35
S]methionine for
1 h and chased for 0 min, 40 min, 80 min, 120 min and 180 min.
Membrane biotinylation was performed after chase. After cell solu-
bilization, total GFP-tagged GAT1 wild type and mutant proteins
were immunoprecipitated with anti-GFP pAb and eluted with
100 lL sample buffer containing 0.5% SDS. The eluates were dilu-
ted with NaCl ⁄ P
i
buffer to 400 lL. The biotin-labelled membrane
proteins were isolated from the diluted eluates with streptavidin
beads. After removal of all membrane proteins, the intracellular pro-
teins were immunoprecipitated with anti-GFP antibodies. Both M
(membrane) and I (intracellular) precipitates were eluted and ana-
lyzed by SDS ⁄ PAGE. (B) The results of the pulse-chase experi-
ments were analyzed by phosphoimager scanning. The total
radioactivity of membrane and intracellular fractions obtained by
immunoprecipitation at each chase time were set at 100%. Each
value represents the mean ± SEM of membrane fractions derived
from three separate experiments.
G. Cai et al. Role of N-glycosylation and N-glycan trimming of GAT1
FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS 1631
after treatment with dMM may be caused by the
reduction in substrate translocation by GAT1 (turn-
over rate).
Defective N-glycosylation results in reduced

GAT1-mediated currents and reduced rate of
external Na
+
interaction
To obtain additional information on the mechanism of
the reduced rate of GABA-uptake due to the muta-
tions, we performed electrical measurements under
voltage clamp for wild type and mutant DDN and
DGN. The number of transporters was calculated
from the transient charge movement in the absence of
extracellular Na
+
[5–7]. Figure 9A shows the depend-
ence of the GAT1-mediated current, expressed as
charges translocated per functional transporter on the
cell surface per second, on the extracellular Na
+
con-
centration. The results were similar to those for the
GABA uptake, in that the current produced by a sin-
gle transporter was reduced by the mutation to 46 and
57% for DGN and DDN, respectively, which is close
to the reduced GABA uptake seen in the flux measure-
ments. A signal from DDQ could hardly be detected.
Though treatment with dMM makes the CHO cells
very unstable for the patch-clamp method, we were,
nevertheless, able to obtain evidence for a reduced
GAT1-mediated current (data not shown). The
dependence on Na
+

concentration reveals that muta-
tion of the two N-glycosylation sites reduced the
apparent affinity from 24 m
)1
to about 8 m
)1
. The
transient currents in the absence of GABA were ana-
lyzed for jumps in potential to the holding potential of
)30 mV. The kinetics of the reaction step associated
with the extracellular Na
+
binding was slowed down
by both mutations (Fig. 9B). All the rate constants
slightly increased with increasing Na
+
concentration,
Fig. 8. Kinetic analysis of GABA-uptake by GFP-tagged GAT1 wild
type (NNN) with and without dMM and N-glycosylation mutant
(DDN). Kinetic analysis of GABA-uptake by GFP-tagged GAT1 wild
type (NNN) with and without dMM and N-glycosylation mutant
(DDN). GABA-uptake assays of wild type NNN pre-incubated with
and without dMM (1 mM) and mutant DDN were performed with
different GABA concentrations. All values presented were calcula-
ted after subtraction of the mock values. The data were fitted by a
Michealis–Menten equation with a common K
m
value of 4.1 lM
and V
max

values of 1.21, 0.55 and 0.29 pmolÆlg protein
)1
Æmin
)1
for
wild type without and with dMM and for DDN, respectively. The
values represent the mean ± SEM of three separate experiments.
Fig. 9. Electrophysiological characterization of GAT1-mediated
steady-state and transient currents. CHO transient transfectants
were subjected to whole-cell patch clamp. (A) Steady-state, GABA-
induced currents were determined during voltage pulses from a
holding potential of )30 mV to )100 mV at different extracellular
Na
+
concentrations. The solid lines represent fits of the Hill equa-
tion with Hill coefficients of n ¼ 1.3 for the wild type GAT1 and
n ¼ 1 for the mutants (as used previously by Liu et al. 1998) and
K
m
Na values of 40 and about 130 mM for wild type and the
mutants, respectively. (B) The rate constants of the GAT1-mediated
current decline in response to a voltage jump from +100 mV to
)30 mV were determined at different extracellular Na
+
concentra-
tions. All data are averages of 3 to 9 determinations ± SEM.
Role of N-glycosylation and N-glycan trimming of GAT1 G. Cai et al.
1632 FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS
the value for the wild type NNN being about twice
that for the mutants.

Discussion
There is increasing evidence that cotranslational N-gly-
cosylation crucially influences the three-dimensional
structure, the biological half-life and intracellular traf-
ficking of proteins. It is also essential for many recog-
nition processes [13,14,16]. Previous studies showed
that the mutation of N-glycosylation sites resulted in a
reduction of GABA-uptake activity by GAT1 [7,10].
However, the possibility that this reduction in function
results from a decrease in the number of GABA trans-
porters per cell was not excluded. In order to clarify
whether N-glycans are directly involved in the GABA-
uptake process and whether the modulation of N-gly-
cosylation influences the biochemical properties of this
protein, quantitative and kinetic analysis of GABA
transport expression and activity was performed using
stable CHO transfectants. Both our own and commer-
cially available anti-GAT1 antibodies were unsuitable
for the quantitative analysis of GAT1, as they bind
very weakly to this protein. Therefore, wild type
GAT1 and N-glycosylation defective mutants were
tagged with GFP, which has been reported not to
influence the intracellular distribution of GAT1; more-
over, the tag does not modulate the relevant functions
of GAT1 [20]. For the quantitative analysis of GABA
transport activity, the cell surface expression of GAT1
wild type and N-glycosylation mutants was determined
by cell surface biotinylation and the resulting values
were used for normalization. This is a well established
method for the quantitative analysis of cell surface

proteins, in which the biotinylation reagent does not
react with intracellular proteins [21].
We found that all N-glycosylation mutants were
expressed on the cell surface, even when all three
N-glycosylation sites have been removed, e.g. in
mutant DDQ (Fig. 3A), indicating that N-glycosyla-
tion is not essential for the plasma membrane traffick-
ing of GAT1. However, the mutant DDQ was
expressed at lower levels on the cell surface, indicating
that deficiency of N-glycosylation impairs plasma
membrane translocation of GAT1. It was reported that
deficiency of N-glycosylation influenced the intracellu-
lar trafficking of some glycoproteins [22–25]. In order
to examine whether the N-glycosylation of GAT1 has
influence on its plasma membrane trafficking, a kinetic
analysis by pulse-chase experiments was performed.
These experiments revealed that the plasma membrane
trafficking of N-glycosylation mutant proteins was
reduced (Fig. 7), although the distributions between
cell surface and cell interior of the mutants DGN,
DND and DDN in the steady expression state were
not significantly different from that of wild type NNN
(Fig. 3B). It has been reported that, in the endoplasmic
reticulum (ER) and in the early secretory pathway, the
N-glycans play a pivotal role in protein folding, olig-
omerization, quality control, sorting, and transport.
Thus defective N-glycosylation may lead to a misfold-
ing of this protein, resulting in its partial retention in
the ER and its rapid digestion thereafter [16,19].
Accordingly, our results indicate that the intracellular

trafficking of N-glycosylation mutants was delayed,
and ⁄ or partly inhibited due to retention of some
mutant protein in the ER, followed by digestion.
The transfectants of GAT1 wild type (NNN) and
mutant DGN, DND and DDN exhibited in SDS ⁄
PAGE (Fig. 3A) two intracellular bands, whereas only
one large band was found in the plasma membrane
fraction. However, treatment of the transfectants with
dMM, which inhibits N-glycosylation processing,
resulted in only small band in the SDS ⁄ PAGE for
both the wild type (NNN) (Figs 2C and 4A) and
mutant DNG, DND and DDN (data not shown). The
mutant DDQ, which does not possess any N-glycosyla-
tion site, expressed only one N-glycan-free band of
90 kDa in both the intracellular and the plasma mem-
brane compartments (Fig. 3A). The 108 kDa large
band of NNN was Endo H resistant, whereas digestion
with PNGase F converted it to a 90 kDa N-glycan free
polypeptide (Fig. 2C), indicating a mature N-glycan of
complex type. However, the 96 kDa small band of
NNN was converted to a 90 kDa polypeptide after
either Endo H- or PNGase F-digestion (Fig. 2B,C),
indicating an N-glycan of the mannosidic type. The
N-glycosylation mutants DGN, DDN and DND
exhibited a reduced molecular mass in accordance with
the absence of N-glycans at the two eliminated N-gyl-
cosylation sites in those proteins. This suggests that
the mutants DGN, DND and DDN, as well as wild
type NNN, were N-glycosylated in CHO cells and
their N-glycans were processed before they arrived at

the cell surface.
To determine the influence of N-glycosylation on the
quality control of GAT1, the half-life of GAT1 wild
type and mutants was also investigated by kinetic ana-
lysis of pulse-chase experiments. We found that the
half-life of mutants containing either double (DDN) or
triple (DDQ) N-glycosylation mutations was remark-
ably reduced (Fig. 6). Keynan et al. reported that the
functional expression of the GABA transporter in
HeLa cells was abolished by tunicamycin, a potent
inhibitor of N-glycosylation [18]. We found that inhibi-
tion of N-glycosylation processing by dMM did not
G. Cai et al. Role of N-glycosylation and N-glycan trimming of GAT1
FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS 1633
affect either the protein stability (Fig. 6) or intracellu-
lar trafficking (Figs 4A and 5A). This suggests that
cotranslational N-glycosylation, but not the terminal
trimming of N-glycans is involved in the regulation of
the stability and trafficking of GAT1. It has been
reported that a variety of molecular chaperones and
folding enzymes assist the folding of newly synthesized
proteins in the ER. If N-glycosylation is inhibited,
some glycoproteins fail to fold or assemble efficiently,
resulting in a prolonged retention in the ER and an
increased proteolytic breakdown [26–29]. Our results
indicate that the impaired plasma membrane traffick-
ing and reduced stability of the N-glycosylation
mutants of GAT1 must be a result of misfolding of
these proteins.
The deficiency of N-glycosylation results in a mark-

edly reduced GABA-uptake activity (Fig. 3C) as well
as a GAT1-mediated current in CHO cells (Fig. 9A).
This is in accordance with our previous work using the
expression system of the Xenopus oocyte [7]. In order
to exclude the possibility that the reduction in function
in the mutants could be due to a reduction in the num-
ber of GABA transporters per cell, values for the
transport activity were normalized for the surface pro-
tein of these mutants. Double N-glycosylation mutants
showed a marked reduction of GABA-uptake activity
of 60–40% of that of the wild type. GAT-mediated
GABA transport activity could hardly be detected in
the mutant lacking all three N-glycosylation sites
(DDQ), despite the fact that this protein was expressed
on the surface of CHO cells (Fig. 3A). The N-glyco-
sylation processing inhibitor 1-deoxymannojirimycin
(dMM) also strongly inhibited GABA-uptake (Figs 4B
and 5B), although the amount of cell surface expres-
sion and the intracellular trafficking of GAT1 were
not affected by dMM (Figs 4A and 5A). This indicates
that the observed reduction of GABA-uptake activity
is a result of a deficiency of N-glycans. The possibility
that the reduced GAT1 activity could be due to a gen-
eral effect of the inhibitor on other glycoproteins
required for GAT1 activity is very unlikely. It has been
demonstrated that GAT transport function can be
reconstructed in liposomes and that no other pro-
teins are needed for GABA-uptake activity [30]. Our
results suggest that N-glycans, in particular their
terminal structure, are involved in the GABA-uptake

process of GAT1. However, the GABA-uptake tolerates
the modification of neuraminic acid to N-propanoyl
neuraminic acid, as incubation with N-propanoylman-
nosamine (P-NAP), a synthetic precursor of N-propa-
noyl neuraminic acid [31–33], did not significantly
change the GABA-uptake activity of GAT1 (data not
shown).
How do the oligosaccharides of GAT1 influence
GABA-uptake? In order to clarify the functional mech-
anism of oligosaccharide side chains in GABA-uptake,
a kinetic analysis was performed. Deficient N-glycosy-
lation decreased the V
max
values of GABA-uptake by
GAT1, while the K
m
GABA values were not affected.
Similar results were also obtained after treatment with
dMM (Fig. 8). Our results indicate that the turnover
rate of the transporter is affected, but not the substrate
binding process. This provides strong evidence that
N-glycans, in particular their terminal structures, are
involved in regulating the GABA translocation of
GAT1, but not in binding of GAT1 to GABA.
Transport of GABA by GAT1 across the cell mem-
brane is driven by an electrochemical gradient of Na
+
and Cl
–
[1,34] with a stoichiometry that results in an

electrogenic substrate transport. Voltage-clamp experi-
ments suggest that deficient N-glycosylation reduces
the affinity of GAT1 for Na
+
[7]. The present work
revealed that the reduced transport activity can at least
partially be attributed to a reduced apparent affinity of
GAT1 for extracellular Na
+
and slowed kinetics of
the transport cycle (Fig. 9). This was observed in both
wild type and mutants after inhibition with dMM. As
the GABA transport process is driven by the gradient
of Na
+
, it is reasonable to deduce that the affinity of
GAT1 for Na
+
determines the turnover rate of GABA
transport. As the data presented in Fig. 9 are for a sin-
gle, functional transporter expressed on the cell sur-
face, the reduced GABA-uptake cannot be due to
reduced cell surface expression of transporters. In this
event the oligosaccharides of GAT1 play a role in the
regulation of GABA-uptake by affecting the affinity
for sodium ions.
In conclusion, cotranslational N-glycosylation is
important for the correct folding of GAT1 to a func-
tional conformation. Defective N-glycosylation leads
to decreased protein stability and disturbed intracellu-

lar trafficking. N-Linked oligosaccharides, in particular
their terminal structures, are involved in the regulation
of GABA-transport of GAT1 by influence on its affin-
ity for sodium ions.
Experimental procedures
Construction of N-glycosylation mutants of GAT1
and of GAT1/GFP fusion proteins
The mutants were based on the neuronal wild type GABA
transporter type 1 of mouse (mGAT1). The cDNAs of
N-glycosylation mutants DGN (N176D, N181G), DDN
(N176D, N181D) and DND (N176D, N184D)
were constructed earlier [7]. In each mutant, two of three
Role of N-glycosylation and N-glycan trimming of GAT1 G. Cai et al.
1634 FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS
N-glycosylation sites Asn (N) were mutated to Asp (D) or
Gly (G). The mutant DDQ (N176D, N181D and N184Q)
was constructed by site-directed mutagenesis in DDN with
synthetic oligonucleotides bearing the designated mutation:
Forward, 5¢-ACCACCCAAATGACCAGC-3¢; reverse,
5¢-GCTGGTCATTTGGGTGGT-3¢. The reaction was per-
formed using the QuikChange
TM
site-directed mutagenesis
kit from Stratagene (Heidelberg, Germany). All mutants
were cloned into the expression vector pCDNA3 (Invitrogen,
Karlsruhe, Germany) and confirmed by sequence analysis.
To construct the GAT1 ⁄ GFP fusion proteins, HindIII—
StuI fragments containing the cDNAs of GAT1 wild type
or mutants were cut from the pAlter-1 vector, then cloned
into HindIII and SmaI sites of pEGFP-N1 vector

(Clontech, Heidelberg, Germany) containing the cDNA
encoding the red-shifted GFP-variant. In this construct,
the cDNAs of GAT1 wild type or mutants were ligated
with cDNA of GFP with an identical reading frame, which
was confirmed by sequence analysis.
Preparation of polyclonal anti-GAT
and anti-GFP sera
Four different oligopeptides: LPWKQCDNPWNTDR
(159–172), MHQMTDGLDKPGQIRC(197–211), DEYPR-
LLRNRRELFC(409–423) and SEDIVRPENGPEQPQAC
(584–599), corresponding to sequences in the extracellular
and intracellular loops of GAT1, were synthesized and used
for immunization of rabbits. Specificity of the antiserum
was verified by immunoblotting with single or mixed
peptides. A polyclonal anti-GFP serum was prepared as
described previously [35].
Transfection of CHO cells and selection
of stable transfectants
In order to produce stable transfectans, each plasmid DNA
(2–4 lg) was transfected into 4 · 10
5
CHO cells using the
Eppendorf Multiporator and an appropriate Eppendorf
protocol (Wesseling-Berzdorf, Germany). Transfectants
were cultured in six-well plates in alpha-modified Eagle’s
medium (MEM alpha) containing 440 mgÆL
)1
l-glutamine
and 10% (v ⁄ v) fetal bovine serum for 2 days, then selected
with 400 mgÆL

)1
geneticin G418 for 1–3 weeks. The expres-
sion of GAT1 ⁄ GFP fusion proteins was detected by flow
cytometry, fluorescence microscopy and immunofluores-
cence microscopy. The stable transfectants expressing
GAT1 ⁄ GFP fusion proteins were selected by flow cytometry
with the FACS Vantage cell-sorter (Becton Dickinson,
Erembodegem, Belgium).
Higher transfection efficiencies were necessary for the
electrophysiological investigations. These were achieved
with transiently transfected CHO cells. The transient trans-
fection was performed using Superfect (Qiagen, Hilden,
Germany) or FuGENE6 (Roche, Mannheim, Germany) as
reagents according to the protocols of Qiagen or Roche,
respectively. The cells were used for electrophysiological
experiments 1–2 days after transfection.
Flow cytometry and immunofluorescence
microscopy
For analysis of surface expression of GFP tagged GAT1
and mutants, the anti-GAT1 peptide-specific polyclonal
IgG was used for immunostaining transfectants at room
temperature for 30 min. After washing with NaCl ⁄ P
i
cells
were incubated with R-phycoerythrin-conjugated goat anti-
(rabbit IgG) Ig at room temperature for 20 min and then
analyzed by flow cytometry.
For immunofluorescence microscopy cells were fixed with
3% (v ⁄ v) formaldehyde in NaCl ⁄ P
i

at room temperature for
10 min. After permeabilization with 0.1% (v ⁄ v) Triton
X-100 in NaCl ⁄ P
i
at room temperature for 5 min, cells were
then extensively washed with NaCl ⁄ P
i
and blocked with 5%
bovine serum albumin and 0.1 m glycine in NaCl ⁄ P
i
for
30 min and washed with NaCl ⁄ P
i
again. Polyclonal antibod-
ies against GAT1 were used for immunostaining at room
temperature for 2 h. After further washing with NaCl ⁄ P
i
,
the cells were incubated with R-phycoerythrin conjugated
goat anti-(rabbit IgG) Ig (diluted 1 : 200) at room tempera-
ture for 1 h. The cells were extensively washed again with
NaCl ⁄ P
i
and then mounted with glycerol ⁄ NaCl ⁄ P
i
(10 : 1,
by volume) for fluorescence microscopy.
Immunoprecipitation and western blotting
analysis
Harvested cells were solubilized; followed by centrifugation

at 40 000 g for 30 min. Aliquots of the supernatant were
subjected to Western blotting as described previously [36].
For immunoprecipitation experiments, the supernatant was
incubated with protein A-Sepharose-bound anti-GFP poly-
clonal antibodies for 12 h at 4 °C. After intensive washing,
immunoprecipitates were eluted by boiling for 4 min in
SDS sample buffer. SDS ⁄ PAGE was performed according
to Laemmli [37]. After electrophoresis, the separated pro-
teins were transferred onto a nitrocellulose membrane. The
anti-GAT1 polyclonal antiserum or anti-GFP mAb
MAB2510 (Chemicon, Temecula, CA, USA) was used for
immunostaining. Visualization was performed with peroxi-
dase-conjugated goat anti-(rabbit IgG) or rabbit anti-
(mouse IgG) Ig (Sigma, St. Louis, MD, USA) and the
chemiluminescent reagent luminol.
Pulse-chase experiments
Pulse-chase experiments were performed as described earlier
[26]. Cells (2 · 10
6
cells per dish) were incubated at 37 °C
G. Cai et al. Role of N-glycosylation and N-glycan trimming of GAT1
FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS 1635
for 4 h in Dulbecco’s modified essential medium lacking
cysteine and methionine. The cells were pulse-labelled with
[
35
S]methionine (ICN, Irvine, CA, USA) for the indicated
times, using 3.7 · 10
6
Bq per dish. After different chase

times, cells were solubilized. The expressed GAT1 ⁄ GFP
fusion proteins were immunoprecipitated with polyclonal
anti-GFP antiserum and analyzed by SDS ⁄ PAGE (7.5%).
Quantification of radio-labeled protein was carried out on a
PhosphorImager
TM
(Molecular Dynamics, Sunnyvale, CA,
USA) using iplabgel software. The total protein of the cell
surface and intracellular bands of each wild type or mutant
were set at 100%.
Endoglycosidase H treatment
Immunoprecipitates were eluted by boiling for 4 min in
buffer containing 0.4% SDS, 1% 2-mercaptoethanol and
40 mm EDTA. Endoglycosidase H (Endo H, Boehringer
Mannheim) treatment was performed with Endo H
(0.02 U ⁄ 80 lL) at 37 °C for 16 h in 50 mm sodium acetate
containing 0.5 lL protease inhibitor cocktail (Sigma) at
pH 5.5.
PNGase F treatment
Immunoprecipitates were eluted by boiling for 4 min in
buffer containing 0.5% (v ⁄ v) SDS, 50 mm 2-mercaptoetha-
nol. PNGase F (Roche) treatment was performed with
PNGase F (15 UÆ40 lL
)1
)at37°C 16 h in 500 mm
NaCl ⁄ P
i
containing 0.5% (w ⁄ v) Mega 10 and 0.5 lL pro-
tease inhibitor cocktail (Sigma) at pH 7.5.
Labelling of cell surface proteins with sulfo-NHS-

biotin and isolation of membrane proteins
Cell surface proteins were labelled with NHS-LC-biotin
according to the protocol of Chen et al. [38]. Cells were
washed three times with ice-cold NaCl ⁄ P
i
(pH 8.0) and incu-
bated with a freshly prepared solution of NHS-LC-biotin
for 20 min at 4 ° C [1.5 mgÆmL
)1
EZ-Link
TM
sulfo-NHS-
LC-biotin (Pierce, Rockford, IL, USA) in 10 mm Hepes
buffer pH 9.0, 2 mm CaCl
2
, 150 mm NaCl]. Cells were
washed with 100 mm glycine then incubated in 100 mm gly-
cine for 20 min. The supernatant of the solubilized cells was
incubated with 50 lL streptavidin beads (Pierce) at 4 °C for
10 h. After extensive washing, all the membrane proteins
attached to beads were eluted by boiling for 4 min in SDS
sample buffer, then subjected to western blotting analysis.
Analysis of plasma membrane and intracellular
expressions of GAT1/GFP fusion proteins
After biotinylation, cell solubilization and centrifugation,
all the membrane proteins were isolated from the superna-
tants using streptavidin beads (see above). The intracellular
GAT1 ⁄ GFP fusion proteins in remaining fractions were
isolated by immunoprecipitation with anti-GFP Igs. Both
the plasma membrane and intracellular proteins were eluted

by boiling for 4 min in SDS sample buffer, and then ana-
lyzed by SDS ⁄ PAGE and western blotting. Either anti-
GFP mAb MAB2510 or anti-GAT1 pAb was used for the
immunostaining. The protein bands obtained in western
blotting were analyzed by phosphoimager scanning. The
total protein of the cell surface and intracellular bands of
each wild type or mutant were set at 100%.
Measurement of [
3
H]GABA uptake
To determine the transport activity, uptake of [
3
H]GABA
(Amersham-Pharmacia Biotech, Freiburg, Germany) was
measured in the presence of 128 mm external Na
+
and
10 lm total GABA. Cells incubated in 96-well tissue culture
plates were washed three times with wash buffer (128 mm
NaCl, 5.2 mm KCl, 2.1 mm CaCl
2
, 2.9 mm MgSO
4
,5mm
dextrose and 10 mm Hepes) and then incubated with
200 lL wash buffer containing 3.7 · 10
4
Bq [
3
H]GABA,

10 lm cold GABA, 3.7 · 10
4
Bq [
14
C]sucrose (Amersham-
Pharmacia Biotech) and 100 lm cold sucrose for 15 min at
room temperature. The sucrose was used to detect leaky
cell batches, which were then excluded from the analysis.
The uptake was stopped by washing cells three times with
cold wash buffer, followed by solubilization of the cells
with 100 lL of 0.5% (w ⁄ v) SDS solution for 1 h at 4 °C.
Aliquots were used for measurement of the remaining
[
3
H]GABA and [
14
C]sucrose. The protein concentration in
the supernatant was determined using the bicinchoninic
acid protein assay reagent (Pierce, USA). Unspecific uptake
was determined in mock transfected cells. The GABA-
uptake activity was measured as pmÆlg protein
)1
Æmin
)1
.
For quantitative measurement of the activity of GABA-
uptake, an aliquot of the stable CHO transfectants was
used for GABA-uptake assay, and another aliquot was
used to determine the quantity of plasma membrane pro-
teins of GAT1 and mutants (see above). The relative

amounts of plasma membrane proteins of GAT1 or
mutants were analyzed by imager scanning of western blots.
The GABA-uptake activity was normalized with the same
amount of plasma membrane proteins of wild type and
mutants. The activity of GABA-uptake by NNN was set
at 100%. All other values were expressed relative to this
value.
Patch-clamp experiments
Voltage-clamp experiments were performed on CHO tran-
sient transfectants in the whole-cell patch-clamp configur-
ation. Steady-state and transient currents were measured in
response to rectangular voltage jumps from a holding
Role of N-glycosylation and N-glycan trimming of GAT1 G. Cai et al.
1636 FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS
potential of )30 mV to potentials of )100 or +40 mV,
using the EPC9 patch-clamp system and pulse software
(HEKA, Lambrecht, Germany). From transient charge
movements in the absence of GABA, the amount and volt-
age dependence of external Na
+
interaction with the trans-
porter can be determined [7]. From the time course of the
exponential current decline, the rate constant for a step
associated with extracellular Na
+
binding can be deter-
mined. In addition, the total amount of charge Q
max
moved
by the transporters and the effective valency of the charge

z* moved by a single transporter can be calculated. Hence,
the number of transporters N is given by N ¼ Q
max ⁄
z* and
the turnover rate k of the charges transported across the
membrane is given by k ¼ I ⁄ Q
max
, where I is the steady-
state current generated by the single functioning GAT1
molecule on the cell surface in the presence of GABA.
The external bath solution contained 150 mm NaCl,
2mm MgCl
2
,2mm CaCl
2
and 5 mm Mops (pH ¼ 7.2).
For solutions with reduced Na
+
, the NaCl was replaced by
equimolar concentrations of TMACl. The pipette solution
in contact with the cell interior contained 150 mm
K-d-gluconate, 2 mm MgCl
2
,5mm EGTA and 5 mm
Mops (pH ¼ 7.2).
Acknowledgements
This work was supported in part by the Special Funds
for Major State Basic Research of China (Grant
G1999053907), a grant from the Chinese Academy of
Sciences, a grant from the Deutsche Forschungsgeme-

inschaft Bonn, the Sonnenfeld-Stiftung and the Fonds
der Chemischen Industrie, Frankfurt⁄ Main. The
cooperation between China and Germany was suppor-
ted on the basis of an agreement between the Max-
Planck Gesellschaft and the Chinese Academy of
Sciences. We are grateful to P Donner and A Becker
(Schering, AG) for the synthesis of peptides, to A
Niedergesa
¨
ss for production of anti-GFP sera, to Q
Gu and M Richter for technical assistance.
References
1 Schloss P, Mayser W & Betz H (1992) Neurotransmitter
transporters: a novel family of integral plasma mem-
brane proteins. FEBS Lett 307, 76–80.
2 Radian R & Kanner BI (1983) Stoichiometry of
sodium- and chloride-coupled c-aminobutyric acid
transport by synaptic plasma membrane vesicles isolated
from rat brain. Biochemistry 22, 1236–1241.
3 Keynan S & Kanner BI (1988) c-Aminobutyric acid
transport in reconstituted preparations from rat brain:
coupled sodium and chloride fluxes. Biochemistry 27,
12–17.
4 Loo DD, Eskandari S, Boorer KJ, Sarkar HK & Wright
EM (2000) Role of Cl
–
in electrogenic Na
+
-coupled
cotransporters GAT1 and SGLT1. J Biol Chem 275 ,

37414–37422.
5 Mager S, Naeve J, Quick M, Labarca C, Davidson N &
Lester HA (1993) Steady states, charge movements, and
rates for a cloned GABA transporter expressed in Xeno-
pus oocytes. Neuron 10, 177–188.
6 Mager S, Kleinberger-Doron N, Keshet GI, Davidson
N, Kanner BI & Lester HA (1996) Ion binding and per-
meation at the GABA transporter GAT1. J Neurosci
16, 5405–5414.
7 Liu Y, Eckstein-Ludwig U, Fei J & Schwarz W (1998)
Effect of mutation of glycosylation sites on the Na
+
dependence of steady-state and transient currents gener-
ated by the neuronal GABA transporter. Biochim Bio-
phys Acta 1415, 246–254.
8 Schloss P, Puschel AW & Betz H (1994) Neurotransmit-
ter transporters: new members of known families. Curr
Opin Cell Biol 6, 595–599.
9 Honda S, Yamamoto M & Saito N (1995) Immunocyto-
chemical localization of three subtypes of GABA trans-
porter in rat retina. Brain Res Mol Brain Res 33, 319–
325.
10 Bennett ER & Kanner BI (1997) The membrane topol-
ogy of GAT-1, a (Na
+
+Cl
–
)-coupled gamma-ami-
nobutyric acid transporter from rat brain. J Biol Chem
272, 1203–1210.

11 Varki A (1993) Biological roles of oligosaccharides: all
of the theories are correct. Glycobiology 3, 97–130.
12 Lis H & Sharon N (1993) Protein glycosylation. Struc-
tural and functional aspects. Eur J Biochem 218, 1–27.
13 Helenius A (1994) How N-linked oligosaccharides affect
glycoprotein folding in the endoplasmic reticulum. Mol
Biol Cell 5, 253–265.
14 Fiedler K & Simons K (1995) The role of N-glycans in
the secretory pathway. Cell 81, 309–312.
15 Molinari M & Helenius A (1999) Glycoproteins form
mixed disulphides with oxidoreductases during folding
in living cells. Nature 402, 90–93.
16 Helenius A & Aebi M (2001) Intracellular functions of
N-linked glycans. Science 291, 2364–2369.
17 Melikian HE, Ramamoorthy S, Tate CG & Blakely RD
(1996) Inability to N-glycosylate the human norepi-
nephrine transporter reduces protein stability, surface
trafficking, and transport activity but not ligand recog-
nition. Mol Pharmacol 50, 266–276.
18 Keynan S, Suh YJ, Kanner BI & Rudnick G (1992)
Expression of a cloned gamma-aminobutyric acid
transporter in mammalian cells. Biochemistry 31,
1974–1979.
19 Verbert A & Cacan R (1999) [‘Glyco-deglyco’ processes
during the biosynthesis of glycoproteins]. J Soc Biol
193, 101–110.
G. Cai et al. Role of N-glycosylation and N-glycan trimming of GAT1
FEBS Journal 272 (2005) 1625–1638 ª 2005 FEBS 1637
20 Chiu CS, Jensen K, Sokolova I, Wang D, Li M, Desh-
pande P, Davidson N, Mody I, Quick MW, Quake SR

& Lester HA (2002) Number, density, and surface ⁄ cyto-
plasmic distribution of GABA transporters at presynap-
tic structures of knock-in mice carrying GABA
transporter subtype 1-green fluorescent protein fusions.
J Neurosci 22, 10251–10266.
21 Law RM, Stafford A & Quick MW (2000) Functional
regulation of gamma-aminobutyric acid transporters by
direct tyrosine phosphorylation. J Biol Chem 275,
23986–23991.
22 Gut A, Kappeler F, Hyka N, Balda MS, Hauri HP &
Matter K (1998) Carbohydrate-mediated Golgi to cell
surface transport and apical targeting of membrane pro-
teins. EMBO J 17, 1919–1929.
23 Benting JH, Rietveld AG & Simons K (1999) N-Gly-
cans mediate the apical sorting of a GPI-anchored, raft-
associated protein in Madin-Darby canine kidney cells.
J Cell Biol 146, 313–320.
24 Scheiffele P, Peranen J & Simons K (1995) N-Glycans
as apical sorting signals in epithelial cells. Nature 378,
96–98.
25 Martina JA, Daniotti JL & Maccioni HJ (1998) Influ-
ence of N-glycosylation and N-glycan trimming on the
activity and intracellular traffic of GD3 synthase. J Biol
Chem 273, 3725–3731.
26 Fan H, Meng W, Kilian C, Grams S & Reutter W (1997)
Domain-specific N-glycosylation of the membrane
glycoprotein dipeptidylpeptidase IV (CD26) influences its
subcellular trafficking, biological stability, enzyme
activity and protein folding. Eur J Biochem 246, 243–251.
27 Trombetta ES & Helenius A (1998) Lectins as chaper-

ones in glycoprotein folding. Curr Opin Struct Biol 8,
587–592.
28 Molinari M & Helenius A (2000) Chaperone selection
during glycoprotein translocation into the endoplasmic
reticulum. Science 288, 331–333.
29 Fan H, Dobers J & Reutter W (2001) DPPIV ⁄ CD26:
structural and biological characteristics of asparagine
and cysteine mutants. In Cell-Surface Aminopeptidases:
Basic and Clinical Aspects (Mizutani S, ed), pp. 303–
316. Elsevier, Amsterdam, the Netherlands.
30 Kleinberger-Doron N & Kanner BI (1994) Identification
of tryptophan residues critical for the function and tar-
geting of the gamma-aminobutyric acid transporter
(subtype A). J Biol Chem 269, 3063–3067.
31 Kayser H, Geilen CC, Paul C, Zeitler R & Reutter W
(1992) Incorporation of N-acyl-2-amino-2-deoxy-hexoses
into glycosphingolipids of the pheochromocytoma cell
line PC 12. FEBS Lett 301, 137–140.
32 Keppler OT, Horstkorte R, Pawlita M, Schmidt C &
Reutter W (2001) Biochemical engineering of the N-acyl
side chain of sialic acid: biological implications. Glyco-
biology 11, 11R–18R.
33 Oetke C, Brossmer R, Mantey LR, Hinderlich S, Isecke
R, Reutter W, Keppler OT & Pawlita M (2002) Versa-
tile biosynthetic engineering of sialic acid in living cells
using synthetic sialic acid analogues. J Biol Chem 277,
6688–6695.
34 Nelson N (1998) The family of Na
+
⁄ Cl

–
neurotransmit-
ter transporters. J Neurochem 71, 1785–1803.
35 Schulein R, Zuhlke K, Oksche A, Hermosilla R, Furk-
ert J & Rosenthal W (2000) The role of conserved extra-
cellular cysteine residues in vasopressin V2 receptor
function and properties of two naturally occurring
mutant receptors with additional extracellular cysteine
residues. FEBS Lett 466, 101–106.
36 Dobers J, Grams S, Reutter W & Fan H (2000) Roles
of cysteines in rat dipeptidyl peptidase IV ⁄ CD26 in pro-
cessing and proteolytic activity. Eur J Biochem 267,
5093–5100.
37 Laemmli UK (1970) Cleavage of structural proteins
during the assembly of the head of bacteriophage T4.
Nature 227, 680–685.
38 Chen JG, Liu-Chen S & Rudnick G (1997) External
cysteine residues in the serotonin transporter. Biochemis-
try 36, 1479–1486.
Role of N-glycosylation and N-glycan trimming of GAT1 G. Cai et al.
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