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Division of Endocrinology, Department of Medicine (L.W., A.C.P.), and the Department of Pharmacology (J.D.F.), Vanderbilt University; and the Department of Veterans Affairs Medical Center (A.C.P.), Nashville, Tennessee 37232
Address all correspondence and requests for reprints to: Dr. Alvin C. Powers, Division of Endocrinology, 715 MRB II, Vanderbilt University, Nashville, Tennessee 37232. E-mail: Al.Powers{at}mcmail.vanderbilt.edu
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Introduction |
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30
mM compared with 1–10 mM for the other GLUT
isoforms), elevations in the blood glucose level during the
postprandial period linearly increase the velocity of glucose transport
and the intracellular glucose concentration. Both transport of glucose
into the liver during the postprandial state and transport of glucose
out of the hepatocyte after gluconeogenesis benefit from GLUT2’s high
Km, because increases in the velocity transport would
parallel increases in either the postprandial glucose concentration or
the intracellular hepatocyte glucose concentration. Furthermore, there
is a correlation between the glucose affinity of GLUT isoforms and the
glucose affinity of the hexokinases with which the GLUT isoforms are
expressed. For example, GLUT2 is usually expressed in tissues that
express glucokinase, which has the highest affinity of the hexokinases
(16, 17). Second, introduction of GLUT2, as opposed to GLUT1, into
insulin-secreting cells results in a more normal insulin secretory
profile (18, 19). Third, animal studies of transplanted pancreatic
islets suggest that GLUT2 expression is required for normal
glucose-stimulated insulin secretion (20, 21). Fourth, although all
members, except GLUT5, can efficiently transport glucose, only GLUT2
and GLUT5, are capable of transporting fructose (14, 22, 23). This
property allows for the uptake of fructose by liver and for transport
of fructose across the basolateral membrane of epithelial cells of the
small intestine. GLUT2’s ability to transport fructose may have
physiological relevance, because fructose utilization by the liver
influences glucose metabolism, and fructose metabolites modulate the
activity of glucokinase and its regulatory protein (24, 25). The molecular determinants that endow GLUT2 with these specialized functions are incompletely understood. One hypothesis is that regions of the GLUT2 protein may be important in the specialized functions of GLUT2 in glucose utilization and insulin secretion. To address this question, we constructed a series of chimeric proteins comprised of GLUT2 and GLUT3 components and analyzed the function of the chimeric GLUT proteins in both Xenopus oocytes and mammalian cells. GLUT3, the primary glucose transporter of neurons, has a high affinity for glucose (14, 26). We used these chimeric proteins to identify the region(s) that is responsible for GLUT2’s distinctive high Km and its ability to transport fructose.
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Materials and Methods |
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-32P]deoxy-CTP (3000 Ci/mmol) were obtained from New
England Nuclear (Boston, MA). 14C-Labeled
D-fructose (290 mCi/mmol) was purchased from Sigma (St.
Louis, MO). The sources for reagents were: phloretin and
3-O-methyl-D-glucose, Aldrich (Milwaukee, WI);
D-fructose, Fisher Scientific Co. (Fairlawn, NJ); and
collagenase (CLS-2), Worthington Biochemical Corp. (Freehold, NJ).
In vitro RNA transcription kits were purchased from
Stratagene (La Jolla, CA). RNeasy minikits for isolation of total RNA
were obtained from Qiagen (Chatsworth, CA). Oligolabeling kits for
synthesizing double stranded DNA probes were purchased from Pharmacia
Biotech (Piscataway, NJ). Affinity-purified rabbit antihuman GLUT3 was
obtained from Charles River Pharmservices (Southbridge MA).
Peroxidase-conjugated affinity-purified goat antirabbit IgG was
purchased from Jackson ImmunoResearch Laboratories (West Grove, PA).
ECL Western blotting detection kits were purchased from Amersham
(Arlington Heights, IL). Plasmids pCA14 and pJM17 as well as 293 cells
were purchased from Microbix Biosystems (Toronto, Canada).
Oligonucleotides were synthesized in the DNA Core Laboratory of the
Vanderbilt Diabetes Research and Training Center.
Construction of GLUT2/GLUT3 chimeric complementary DNAs (cDNAs)
The cDNAs encoding the human glucose transporters GLUT2 and
GLUT3 were gifts from Dr. Graeme Bell (University of Chicago, Chicago,
IL) and were used as previously reported to construct chimeric GLUT
cDNAs (27). The chimeric glucose transporter cDNAs, comprised of
fragments from human GLUT2 and GLUT3, were constructed by gene overlap
extension PCR or the introduction of unique restriction enzyme sites
into GLUT2 cDNA by PCR and exchange of the cDNA fragments between GLUT2
and GLUT3. The design of these chimeric cDNAs was based on the
predicted two-dimensional structure of GLUT proteins proposed by
Mueckler et al. (8, 9). All chimeric GLUTs were created so
that the predicted transmembrane segments were not disrupted but were
replaced by the corresponding regions of the other glucose transporter
protein. For overlap extension PCR, the GLUT2 and GLUT3 cDNA fragments
were amplified separately in a first round of PCR from the coding
sequence of human GLUT2 and human GLUT3 cDNAs. The generated GLUT2 and
GLUT3 cDNA fragments were subsequently annealed in a second round of
PCR and extended in a third round of PCR as described previously (27).
DNA polymerase pfu (Stratagene, La Jolla, CA) was chosen to increase
the fidelity of DNA synthesis during the PCR. The chimeric GLUT cDNAs
were subcloned into a modified oocyte expression vector (pGOV) and
checked by sequencing as previously described (27).
The chimeric GLUTs created in the present study are listed in Table 1
.
The nomenclature for these chimeric proteins is based on the primary
structure of the glucose transporter proteins. Each name starts with
the NH2-terminal portion of the chimeric GLUT protein, with
the number in parentheses indicating the amino acid boundary
of this portion. GLUT2 is referred to as G2, and GLUT3 is referred to
as G3. As GLUT2 is 524 amino acids in length and GLUT3 is 479 amino
acids in length (due to a larger extracellular loop between
transmembrane segments 1 and 2 of GLUT2), the locations of predicted
transmembrane segments do not occur at identical amino acid positions.
The predicted two-dimensional structures of these chimeric GLUT
proteins are shown in Fig. 2.
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Expression of GLUT isoforms in Xenopus oocytes
Oocytes for microinjection were isolated from female
Xenopus laevis (Nasco, Fort Atkinson, WI) as previously
described (27). Capped RNA (10–20 ng) in a volume of 10–20 nl was
microinjected into each oocyte within 24 h of isolation, using a
Picospritzer II (General Valve Corp., Fairfield, NJ) and a
micromanipulator (Singer Instruments Co., Somerset, UK). Microinjected
oocytes were incubated at 18 C for 48 h before assay, during which
time the medium was changed every 12 h, and unhealthy oocytes were
removed.
3OMG and fructose transport assays
Two days after RNA injection, healthy oocytes were selected for
3OMG transport assay. To determine maximal GLUT expression, oocytes
injected with increasing amounts of GLUTs RNA were incubated in 1
mM 3OMG and 1.2 µCi [3H]3OMG at room
temperature for 15 min. The transport of methylglucose was stopped by
washing oocytes four times with ice-cold buffer containing 100 µm
phloretin. The oocytes were transferred to scintillation vials and
lysed; radioactivity was quantitated by liquid scintillation counting.
To determine the equilibrium exchange Km for methylglucose,
the oocytes were incubated in a range of 3OMG concentrations (1, 5, 10,
15, 20, 30, and 40 mM) at 18 C for 15–18 h to reach
equilibrium exchange conditions. Ten oocytes for each glucose
concentration were used as a group, and each assay was performed in
triplicate. Before each assay, a preliminary uptake was performed by
incubating oocytes (under equilibrium exchange conditions with 1
mM 3OMG) with 1.2 µCi [3H]3OMG at room
temperature for 60 min. This incubation time is sufficient for the
oocytes to reach maximal uptake of [3H]3OMG. The uptake
of [3H]3OMG reached with the 60-min incubation and with
incubations of 1–10 min was used to select a transport assay time
(1, 2, 3, 4, 5), so that the uptake of [3H]3OMG was less than 20%
of that reached during the 60-min incubation. For the final assay,
transport was initiated by adding 1.2 µCi [3H]3OMG into
each group of oocytes equilibrated within the different 3OMG
concentrations. The transport of methylglucose was stopped by washing
oocytes four times with ice-cold buffer containing 100 µm phloretin.
The oocytes were transferred to scintillation vials and lysed;
radioactivity was quantitated by liquid scintillation counting. Each
assay was repeated at least four times for each chimeric glucose
transporter.
For measurement of fructose transport, 2 days after RNA injection healthy oocytes were selected and incubated in 1 mM of fructose at 18 C for 15–18 h to reach equilibrium exchange conditions. Ten healthy oocytes were used as a group, and each assay was performed in triplicate. To initiate the transport assay, 0.13 µCi [14C]fructose was added to each group of oocytes. The oocytes were then incubated at room temperature for 60 min. The uptake of fructose was stopped by washing the oocytes four times with ice-cold buffer containing 100 µm phloretin. The oocytes were transferred to scintillation vials and lysed. The radioactivity was quantitated by liquid scintillation counting. A transport assay for 1 mM 3OMG was performed in parallel for each glucose transporter.
Kinetic analysis of 3OMG and fructose transport
The kinetic analysis of sugar uptake was performed by
calculating the clearance of the oocyte medium at various sugar
concentrations. Linear regression of a Hanes plot [(substrate
concentration/velocity) vs. substrate concentration] was
used to calculate the Km for methylglucose transport. A
time course of methylglucose transport at 1 mM 3OMG was
performed for each experiment to ensure that initial rates of transport
were obtained. The Km values from individual experiments
were averaged and presented as the mean and SEM.
Statistical comparisons of the Km for chimeric GLUTs used
Student’s unpaired t test.
Construction of recombinant adenoviruses carrying wild-type or
chimeric GLUT cDNAs
Recombinant adenoviruses carrying wild-type GLUT2, wild-type
GLUT3, or chimeric GLUT2-(1–303)/GLUT3-(270–497) (chimera 9 in Table 1
) cDNAs were generated by homologous
recombination as previously described (28). The cDNA encoding each GLUT
was subcloned into plasmid pCA14. The resulting plasmid was
cotransfected with pJM17 into 293 cells by calcium phosphate
transfection. Two to 3 weeks later, individual clones were chosen and
confirmed by Southern blot analysis. The recombinant adenovirus was
amplified in 293 cells and purified by cesium chloride centrifugation.
The multiplicity of infection (MOI) of the viral stock was determined
by a plaque assay with 293 cells and expressed as infectious units per
microliter. The viral stocks were stored at -70 C until use. The terms
for the recombinant adenoviruses used in this study are AdGLUT2
(containing wild-type GLUT2 cDNA), AdGLUT3 (containing wild-type GLUT3
cDNA), and AdGLUT2/GLUT3 [containing chimeric
GLUT2-(1–303)/GLUT3-(270–497) cDNA].
In vitro transduction of RIN 1046-38 cells
RIN 1046-38 cells were cultured in DMEM supplemented with 10%
FBS, 100 U/ml penicillin, and 100 µg/ml streptomycin. The cells were
infected with a recombinant adenovirus by adding the desired amount of
viral stock directly to the culture medium. The MOI was selected by
examining cell survival and transgene expression as a function of
increasing MOI. AdGLUT2 was used at a MOI of 80. AdGLUT3 was used at a
MOI from 10–50. AdGLUT2/GLUT3 was used at a MOI from 5–50. The
viruses were removed 24 h later by washing the cells twice with
PBS. The cells were harvested for Northern blot analysis after washing
or were cultured for another 24 h in culture medium for immunoblot
analysis.
Northern blot analysis of GLUT RNA
Total RNA was prepared from RIN 1046-38 cells using the RNeasy
minikit. Northern blot analysis was performed by separating 15 µg
total RNA on 1.5% formaldehyde-agarose gels. The RNA was transferred
onto a nylon membrane by capillary blotting. 32P-Labeled
double stranded DNA probes were synthesized from human
GLUT2-(1–303)/GLUT3-(270–497) cDNA using an oligolabeling kit. This
probe hybridizes to both GLUT2 and GLUT3 mRNA. The hybridizations were
performed at 42 C overnight with formamide hybridization solution. The
membranes were then washed with 2 x SSC-0.1% SDS twice at room
temperature and with 0.1 x SSC-0.1% SDS twice at 65 C, then
subjected to autoradiography.
Western blot analysis of GLUT protein
RIN 1046-38 cells were lysed in ice-cold lysis buffer [0.5%
Nonidet P-40, 5 mM MgCl2, 50 mM
Tris (pH 7.4), and 0.1 mM phenymethylsulfonylfluoride].
The total cellular proteins were separated on 10% SDS-PAGE gels and
electrotransferred to an Immobilon-P membrane (Millipore, Bedford, MA).
The affinity-purified rabbit antihuman GLUT3 was used as primary
antibody at a 1:500 dilution. This antibody reacts only with the
carboxy-terminus of human GLUT3. The affinity-purified
peroxidase-conjugated goat antirabbit IgG was used as a secondary
antibody at a dilution of 1:10,000. Antibody binding was detected by
chemiluminescence using the ECL Western blotting detection kit
according to the manufacturer’s protocol.
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Results |
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). A time
course of 3OMG accumulation, performed under the condition of maximal
GLUT expression, was linear for approximately 15 min and reached a
maximum at 60 min for GLUT2- and GLUT3-expressing oocytes (data not
shown). The oocytes expressing chimeric GLUTs demonstrated properties
similar to those of GLUT2- or GLUT3-expressing oocytes for
methylglucose transport (data not shown). Under the maximal GLUT
expression and equilibrium exchange conditions, the average velocity of
1 mM 3OMG transport was 24.83 ± 0.41
pmol/min·oocyte (mean ± SEM) for wild-type GLUT2
and 25.21 ± 0.14 pmol/min·oocyte (mean ± SEM)
for wild-type GLUT3 (Table 1). All chimeric proteins containing the
NH2-terminal portion of GLUT3 (chimeras 2–7 in Table 1)
had uptake rates for 3OMG similar to those of wild-type GLUT2 and
wild-type GLUT3 (P > 0.05), with the exception of
chimera 1, which had a rate significantly lower than those of the
wild-type GLUTs (P < 0.01). The two chimeric proteins
with the NH2-terminal portion of GLUT2 (chimeras 8 and 9 in
Table 1) did not transport more 3OMG than sham-injected oocytes (Table 1).
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). GLUT3/GLUT2 chimeric proteins
containing a region from transmembrane segment 9 to the initial 20
amino acid residues of the COOH-terminus of GLUT2 (chimeras 1, 2, and 4
in Table 1 and Fig. 2) had Km values similar to that of
wild-type GLUT2 and different from that of wild-type GLUT3
(P > 0.05 compared with the Km of
wild-type GLUT2 and P < 0.01 compared with the
Km of wild-type GLUT3). Based on these results, we
constructed two additional chimeric proteins to determine the specific
region(s) responsible for GLUT2’s high Km. Replacing the
region from transmembrane segment 9 to the loop connecting
transmembrane segments 10 and 11 of GLUT2 with the corresponding region
of GLUT3 (chimera 5) resulted in a Km intermediate between
those of wild-type GLUT2 and wild-type GLUT3 (P < 0.05
compared with the Km of wild-type GLUT3 and
P = 0.055 compared with the Km of wild-type
GLUT2). When only transmembrane segments 9 and 10 and the connecting
loop between these two segments of GLUT2 were present in the chimeric
GLUT3/GLUT2 (chimera 6), the Km was similar to that of
wild-type GLUT3 and different from that of wild-type GLUT2
(P > 0.05 compared with the Km of
wild-type GLUT3 and P < 0.01 compared with the
Km of wild-type GLUT2; Fig. 2). These results indicate that
the amino acid residues responsible for GLUT2’s distinctive high
Km reside mainly in the region from transmembrane segments
9–12. Chimera 3 contains a region from GLUT2 that is similar to
chimera 2 and also has nine amino acid residues from GLUT2’s
intracellular loop between transmembrane segments 6 and 7. This chimera
had a Km of 48 ± 4.62 mM (mean ±
SEM), which is significantly higher than the Km
of wild-type GLUT2 (Fig. 2).
Fructose transport of wild-type and chimeric GLUTs
GLUT2 can also transport fructose across the cell membrane,
whereas GLUT3 does not have this ability (Fig. 3). Under the condition of maximal GLUT
expression, the equilibrium exchange uptake rate for 1 mM
fructose was 2.46 ± 0.21 pmol/min·oocyte (mean ±
SEM) for wild-type GLUT2. The wild-type GLUT3 had an uptake
rate of 0.58 ± 0.02 pmol/min·oocyte (mean ±
SEM), which was not significantly different from that in
sham-injected oocytes (data not shown). To localize the region(s) of
GLUT2 responsible for this property, fructose transport by our series
of GLUT3/GLUT2 chimeric proteins was examined. The GLUT3/GLUT2 chimera
containing a region from transmembrane segment 7 to part of the
COOH-terminus of GLUT2 (chimera 3) retained the ability to transport
fructose (P > 0.05 compared with wild-type GLUT2 and
P < 0.01 compared with wild-type GLUT3). Further
narrowing of this region in the GLUT3/GLUT2 chimeric proteins (chimeras
4–6) resulted in a complete loss of the fructose transport ability
(Fig. 3). In parallel experiments, all chimeric proteins had similar
uptake rates for 1 mM 3OMG (Fig. 3). This suggests, but
does not prove, that GLUT expression is similar among the chimeric
proteins (the differences in Km may somewhat influence the
transport rate).
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).
To investigate the molecular basis for the inability to transport
glucose, we also expressed chimeric GLUT2-(1–303)/GLUT3-(270–497),
wild-type GLUT2, and wild-type GLUT3 in RIN 1046-38 cells using
adenovirus-mediated gene transfer. Transduction of RIN 1046-38 cells
with AdGLUT2, AdGLUT3, and AdGLUT2/GLUT3 resulted in expression of
mRNAs with the expected sizes, as demonstrated by Northern blot
analysis (Fig. 4A). However, no
GLUT2/GLUT3 protein was detected by immunoblotting (Fig. 4B).
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Discussion |
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Glucose affinity of GLUT2
Based on prior studies, the distinctive glucose affinity of GLUT2
is a function of the carboxy half of the protein (27, 29). Using a
series of chimeric proteins that exchanged transmembrane segments 7–12
of GLUT2 for corresponding regions of GLUT3, we found that
transmembrane segments 9, 10, 11, and 12 are required for this
property. GLUT2’s distinctive glucose affinity requires the presence
of all four transmembrane segments, as a chimeric protein that excluded
transmembrane segments 9 and 10 (chimera 5) had a Km
intermediate between those of wild-type GLUT2 and wild-type GLUT3,
whereas a chimeric protein that contained only transmembrane segments 9
and 10 (chimera 6) had a Km similar to that of GLUT3. These
results suggest that amino acids in transmembrane segments 9–12, which
are GLUT2 specific, interact with the glucose molecule to facilitate
its transport. Models of glucose transporter topography predict that
these transmembrane segments are part of the pore through which glucose
traverses the membrane (9). One envisions that amino acids specific for
GLUT2 in these transmembrane segments determine its glucose affinity,
but that the overall membrane pore and the ability to transport glucose
are common to all glucose transporter isoforms. These results also
point out a limitation of using chimeric proteins to study protein
function, in that regions that are not contiguous by primary amino acid
sequence may cooperate functionally, and this must be taken into
consideration when designing chimeric proteins. In addition, further
narrowing of this regions using additional chimeric proteins is
unlikely. As continuous amino acids are not required, future efforts
will require assessment of glucose affinity after mutation of
combinations of amino acids widely separated in the primary amino acid
sequence of GLUT2.
The present results contrast with those of Arbuckle et al., who concluded that transmembrane segment 7 alone was responsible for GLUT2’s Km (30). The reason for this disagreement is not clear, as both laboratories examined the function of chimeric GLUT2/GLUT3 proteins in Xenopus oocytes. One possible explanation is that the current study used 3OMG, which measures only glucose transport, whereas Arbuckle et al. used 2-deoxyglucose, which requires both glucose transport and phosphorylation (31). In addition, the current study used a series of chimeric proteins that span transmembrane segments 7–10, whereas Arbuckle et al. used only a single chimeric protein to reach their conclusion. The Km of this chimeric protein in the work of Arbuckle et al. was lower than the Km of GLUT2, but higher than the Km of GLUT3. This suggests that additional regions of GLUT2 participate in fully determining GLUT2’s glucose affinity. As none of the chimeric proteins examined by Arbuckle et al. involved exchanges of transmembrane segments 8–10, the contribution of this area to GLUT2’s Km was not detected. The reason why chimera 3 (containing transmembrane segments 7–12, part of the carboxy-terminal tail, and nine amino acids of the intracellular loop of GLUT2) had a Km significantly higher than that of wild-type GLUT2 is not clear. One possible explanation is that this altered the glucose-binding site and therefore affected the glucose affinity (32, 33). Prior work by our laboratory and by Noel and Newgard has demonstrated a minor contribution of GLUT2’s intracellular carboxy-tail to GLUT2’s glucose affinity (27, 29, 34). Thus, the current findings, integrated with our prior work and the findings of Arbuckle et al. and Noel and Newgard, support the concept that several regions in the carboxy half of a glucose transporter protein cooperate to determine its glucose affinity.
Fructose transport
Fructose transport is a property of only GLUT2 and GLUT5 (14, 22),
and the current study indicates that transmembrane segments 7 and 8 of
GLUT2 are essential for this property (chimera 3 in Fig. 3). These
results indicate that the requirements for binding and transporting
fructose are slightly different from the protein regions responsible
for GLUT2’s glucose affinity. Our findings clarify conflicting results
from other laboratories regarding fructose transport. Based on a series
of GLUT5/GLUT1 chimeric proteins expressed in selected clones of
Chinese hamster ovary cells, Inukai et al. concluded that
the entire GLUT5 protein was required for fructose transport (35).
However, none of the chimeric proteins used in that study exhibited
increased glucose transport, so a conclusion about their ability to
transport fructose is not possible. Noel and Newgard found that a
GLUT2/GLUT1 chimera consisting of the amino-terminus, the first
transmembrane segment, and the extracellular loop of GLUT1, with the
remainder of the protein being GLUT2, was able to transport fructose
but at a reduced velocity compared with wild-type GLUT2 (34). These
researchers concluded that the amino-terminal region of GLUT2 was
important for fructose transport. The current study and the work of
Arbuckle et al. (30) indicate that the amino-terminus of
GLUT2 is not required for fructose transport and that transmembrane
segments 7 and 8 are the essential regions. Both studies examined a
series of GLUT2/GLUT3 chimeric proteins, whereas Noel and Newgard
studied only one GLUT2/GLUT1 chimeric protein. It is not known whether
using GLUT1 rather than GLUT3 is responsible for the different results.
Arbuckle et al. further suggested that the affinity for
fructose was modulated by transmembrane segments 8–12. Interestingly,
this is the same region identified by the current study to determine
GLUT2’s glucose affinity. The current study cannot assess whether
transmembrane segments 9–12 modulate the affinity for fructose. Thus,
the ability of GLUT2 to transport fructose requires amino acids in
transmembrane segments 7 and 8, but transmembrane segments 9–12 may
modulate the affinity for fructose.
Role of amino-terminal region(s) of GLUT2
Chimeric proteins that contained the amino-terminus, the first
transmembrane segment, and the extracellular loop of GLUT2 were not
expressed either in the oocyte system or in mammalian cells using
adenovirus-mediated gene transfer. Even when the entire amino half of
the chimeric protein was GLUT2, there was no glucose transport. This
finding with GLUT2/GLUT3 chimeric proteins confirms our earlier
observation with similar GLUT2/GLUT4 chimeric proteins (27), the
findings of Noel and Newgard with GLUT2/GLUT1 chimeric proteins (34),
and the findings of Arbuckle et al. with GLUT2/GLUT3
chimeric proteins (30). Both our laboratory and that of Noel and
Newgard were able to in vitro translate chimeric proteins
with the amino-terminal regions of GLUT2 into a protein of the expected
size (34). Neither the current study nor that by Noel and Newgard could
detect the chimeric GLUT protein from cells that express the mRNA for
the chimeric protein. Taken together, these findings from three
different laboratories with chimeric proteins of GLUT1, GLUT3, and
GLUT4 indicate that the amino-terminal region(s) of GLUT2 imposes
strict structural requirements on the carboxy-terminus of the glucose
transporter protein. As the amino-terminus of GLUT1, GLUT3, and GLUT4
can participate with the carboxy-terminus of GLUT2, this property is
peculiar to GLUT2. As GLUT2’s extracellular loop is distinctive (64
amino acids compared with 32 amino acids for GLUT1, GLUT3, and GLUT4,
with no sequence homology) (7), it is likely that this region is
responsible for the lack of expression of these chimeric proteins.
These findings suggest that a failure of the amino-terminal regions of
GLUT2 to interact with other GLUT isoforms interferes with protein
stability within the cell. Cope et al. proposed that the
amino-terminus of GLUT1 provides a scaffold for the folding of the
carboxy-terminus of the protein (33), and these findings with GLUT2
support this hypothesis. The amino-terminal region of GLUT2 does not
influence glucose or fructose transport substantially, so the
functional purpose of GLUT2’s distinctive extracellular loop remains
obscure.
Model of functional domains of GLUT2
Based on the results of the current study and the work of
other laboratories, a model of functional domains of glucose
transporter proteins is emerging. Transmembrane segments 7–8 appear to
form a functional domain that is responsible for binding the substrate
(glucose/fructose), whereas another functional domain in transmembrane
segments 9–12 may modulate the affinity of the transporter for the
substrate. The intracellular carboxy-tail of the transporter also
appears to independently module affinity for the substrate and is
involved in the targeting of GLUT4 to its distinctive intracellular
location (36, 37). These functional domains interact to produce the
final substrate specificity and substrate affinity that are distinctive
for each GLUT isoform. As yet, no functional role has been identified
for the extracellular loop between transmembrane segments 1 and 2 or
the intracellular loop between transmembrane segments 6 and 7, but
specific interactions of the amino-terminus of GLUT2 are required for
protein stability.
Whether regions of any GLUT isoforms interact with other membrane or cytoplasmic proteins is not known, but such interactions could provide yet another mechanism for modifying transporter function. It is also unresolved whether GLUT isoforms are interchangeable. For example, could any GLUT isoform substitute for GLUT2 in the liver or pancreatic ß-cell, or are certain features of GLUT2 required? Have the distinctive transporter kinetics and the tissue-specific expression patterns evolved in response to certain requirements of cellular physiology? Studies in vivo to ablate a GLUT isoform in a certain tissue, then introduce a different GLUT isoform and assess the physiological consequences will be required to answer such questions.
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Footnotes |
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Received March 30, 1998.
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