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Cytoplasmic ATP inhibits human erythrocyte glucose transport protein (GLUT1)–mediated glucose transport in human red blood cells by reducing net glucose transport but not exchange glucose transport (Cloherty, E.K., D.L. Diamond, K.S. Heard, and A. Carruthers. 1996. Biochemistry. 35:13231–13239). We investigated the mechanism of ATP regulation of GLUT1 by identifying GLUT1 domains that undergo significant conformational change upon GLUT1–ATP interaction. ATP (but not GTP) protects GLUT1 against tryptic digestion. Immunoblot analysis indicates that ATP protection extends across multiple GLUT1 domains. Peptide-directed antibody binding to full-length GLUT1 is reduced by ATP at two specific locations: exofacial loop 7–8 and the cytoplasmic C terminus. C-terminal antibody binding to wild-type GLUT1 expressed in HEK cells is inhibited by ATP but binding of the same antibody to a GLUT1–GLUT4 chimera in which loop 6–7 of GLUT1 is substituted with loop 6–7 of GLUT4 is unaffected. ATP reduces GLUT1 lysine covalent modification by sulfo-NHS-LC-biotin by 40%. AMP is without effect on lysine accessibility but antagonizes ATP inhibition of lysine modification. Tandem electrospray ionization mass spectrometry analysis indicates that ATP reduces covalent modification of lysine residues 245, 255, 256, and 477, whereas labeling at lysine residues 225, 229, and 230 is unchanged. Exogenous, intracellular GLUT1 C-terminal peptide mimics ATP modulation of transport whereas C-terminal peptide-directed IgGs inhibit ATP modulation of glucose transport. These findings suggest that transport regulation involves ATP-dependent conformational changes in (or interactions between) the GLUT1 C terminus and the C-terminal half of GLUT1 cytoplasmic loop 6–7.
Blood–tissue barriers, which comprise endothelial cells connected by tight junctions (Takata et al., 1997; Mann et al., 2003), protect the brain, peripheral nerve, myocardium, retina, olfactory epithelium, and the inner ear from the external environment. Metabolism in these tissues is fueled by glucose that is transported across the endothelial barrier via a transcellular mechanism mediated by the type I facilitative glucose transport protein, GLUT1 (Takata et al., 1997; Mann et al., 2003; Leybaert, 2005). GLUT1 is a prototypic member of the facilitative glucose transporter family (Joost et al., 2002) and of the wider major facilitator superfamily (MFS) of structurally and functionally related transport proteins (Saier et al., 1999; Saier, 2000).
GLUT1 is expressed at very high levels in endothelial cells and erythrocytes (Takata et al., 1997) where it displays substrate affinities, kinetic properties, and inhibitor pharmacodynamics that distinguish it from other GLUT facilitative sugar transporter members (Takakura et al., 1991). Glucose metabolism in erythrocytes and endothelial cells is not rate limited by transport because the glucose transport capacities of these cells greatly exceeds their glycolytic capacities (Jacquez, 1984; Gerritsen et al., 1988). In spite of this, GLUT1-mediated sugar transport displays acute and adaptive regulation in endothelial cells (Takata et al., 1997; Loaiza et al., 2003; Mann et al., 2003) and acute regulation in erythrocytes (Jung et al., 1971; Taverna and Langdon, 1973; Jacquez, 1983; Weiser et al., 1983; Levine et al., 2005) where cellular ATP depletion enhances GLUT1-mediated sugar import capacity (Carruthers and Zottola, 1996; Heard et al., 2000; Levine et al., 2002; Levine et al., 2005; Leitch and Carruthers, 2007). Acute responses occur within seconds to minutes and involve stimulation of existing cell surface glucose transporters (Diamond and Carruthers, 1993; Shetty et al., 1993; Cloherty et al., 1996). Adaptive responses occur over several hours in response to hypoxia and hypoglycemia and involve changes in glucose transporter expression (Mann et al., 2003).
GLUT1 is a nucleotide binding protein that, when complexed with ATP, displays reduced glucose import capacity but increased affinity for sugar (Carruthers and Helgerson, 1989; Levine et al., 1998; Levine et al., 2002). ATP modulation of GLUT1-mediated transport is competitively inhibited by AMP and ADP, but does not require ATP hydrolysis (Heard et al., 2000). Peptide mapping studies of azidoATP-labeled GLUT1 demonstrate that ATP interacts with GLUT1 residues 301–364 (Levine et al., 1998). This sequence spans transmembrane helices 8 and 9 (TM8 and TM9) and cytoplasmic loop 8–9. Residues 332–343 of this region display 50% sequence identity with a component of the adenylate kinase ATP binding pocket (Levine et al., 1998) and mutagenesis of key residues within this subdomain abolishes ATP modulation of transport (Levine et al., 2002). These observations suggest that nucleotide binding pocket minimally consists of L8–9 and a portion of TM9. Competitive antagonism of ATP regulation of transport by AMP eliminates the possibility that transport regulation is a simple consequence of nucleotide binding to GLUT1. Rather, regulation must involve nucleotide- induced GLUT1 conformational changes but the nature of these changes is unknown. Neither is it known whether the details of GLUT1 regulation are isoform specific or reflect a mechanism fundamental to all structurally related GLUT family members that extends to transport catalyzed by other structurally and functionally related MFS proteins. GLUT1 regulation appears to involve rapid changes in GLUT1 intrinsic activity while regulation of the insulin-sensitive transporter GLUT4 involves rapid redistributions of GLUT4 proteins between intracellular and cell surface membranes in addition to GLUT4 activation (Joost et al., 1986; Simpson and Cushman, 1986).
The results of the present study suggest that ATP binding to GLUT1 causes the GLUT1 carboxyl terminus to interact with GLUT1 cytoplasmic loop 6–7 in a sequence-specific fashion to inhibit transport.
Fresh, de-identified human blood was purchased from Biological Specialties Corporation. Protein assays, Pro Blue coomassie stain, and Supersignal chemiluminescence kits were from Pierce Chemical Co. Nitrocellulose and Immobilon-P were purchased from Fisher Scientific. Purified rabbit IgGs raised against synthetic peptides corresponding to GLUT1 subdomains were obtained from Animal Pharm Services, Inc. These are N-Ab (GLUT1 residues 1–13); L2–3-Ab (GLUT1 residues 85–95); L6–7-Ab (GLUT1 residues 217–231); L7–8-Ab (GLUT1 residues 299–311); C-Ab (GLUT1 residues 480–492). All other reagents were purchased from Sigma-Aldrich.
Saline comprises 150 mM NaCl, 10 mM Tris-HCl, and 0.5 mM EDTA, pH 7.4. Lysis medium contained 10 mM Tris-HCl and 0.2 mM EDTA, pH 7.2. Stripping solution contained 2 mM EDTA, 15.2 mM NaOH, pH 12. Tris medium contained 50 mM Tris-HCl, pH 7.4. Kaline consisted of 150 mM KCl, 5 mM HEPES, 4 mM EGTA, and 5 mM MgCl2. Ammonium bicarbonate was 0.5% (63 mM), pH 9.0. PBS containing Tween (PBS-T) comprised 140 mM NaCl, 10 mM Na2HPO4, 3.4 mM KCl, 1.84 mM KH2PO4, 0.1% Tween, pH 7.3. Stop solution comprises ice-cold Kaline plus cytochalasin B (CB) (10 μM) and phloretin (100 μM).
Glucose transporter (plus endogenous lipids) was purified from human erythrocyte membranes in the absence of reductant as described previously (Hebert and Carruthers, 1992). The resulting GLUT1 proteoliposomes contain (by protein mass) 90% GLUT1, 8% RhD protein, 2% nucleoside transporter (ENT1) and have a lipid:total protein mass ratio of 1:1 (Zottola et al., 1995). Experiments were restricted to the use of GLUT1 preparations in which the stoichiometry of proteoliposomal cytochalasin B binding is 0.48 ± 0.07 mol CB per mol nonreduced GLUT1 and 0.9 ± 0.1 mol CB per mol reduced GLUT1.
Transport in red cell ghosts was measured as described previously (Leitch and Carruthers, 2007).
Purified GLUT1 (10 μg) was digested with a 20:1 (protein: enzyme) ratio of purified porcine trypsin (Princeton Separations) in 50 mM Tris-HCl (pH 7.5), 5 mM MgCl2, 4 mM ATP (pH 7.5). Digestions were performed at 4°C for 60 min, or the indicated time period. Reactions were immediately loaded onto 15% SDS-PAGE.
After SDS-PAGE, samples were transferred to nitrocellulose and blocked overnight in 25% nonfat dry milk/PBS-T. Primary antibody (N-Ab [1:200], L2–3-Ab [1:200], L6–7-Ab [1:500], L7–8-Ab [1:200], L8–9 [1:200], C-Ab [1:15,000], δ-Ab [1:1,000]) was incubated in 3% nonfat dry milk/PBS-T for 1 h at room temperature. Blots were washed three times in PBS-T and incubated with horseradish peroxidase (HRP)–conjugated goat anti-rabbit secondary antibody (1:5,000 dilution) at room temperature for 1 h. Blots were washed three times in PBS-T, developed using Pierce SuperSignal West Pico Chemiluminescent substrate and visualized by autoradiography.
Purified GLUT1 (200 ng) in PBS was adsorbed to each well of the ELISA plate for 2 h at 37°C. Plates were blocked with PBS + 3% BSA for 2 h at 37°C. Primary antibody (N-Ab [1:200], L2–3-Ab [1:200], L6–7-Ab [1:500], L7–8-Ab [1:200], L8–9 [1:100 to 1:500], C-Ab [1:15,000], δ-Ab [1:1,000]) was added to each well in PBS + 0.1% BSA ± ATP and binding was allowed to proceed for 2 h. The plate was washed five times with PBS, and then each well was incubated with HRP-conjugated goat anti-rabbit secondary antibody in PBS + 0.1% BSA (1:5,000 dilution) for 1 h at 37°C. The plate was washed five times with PBS and wells were developed using 100 μl of 1-Step ABTS solution (Pierce Chemical Co.). Primary IgG binding was quantitated as absorbance at 415 nm.
This chimera substitutes the middle loop (L6–7) with that of its rat GLUT4 counterpart and was constructed using a six-step PCR protocol. In PCR 1A, a HindIII primer complimentary to the 5′ end of human GLUT1 and a reverse primer complimentary to nucleotides 600–619 of GLUT1 and 667–685 of GLUT4 was used to generate a fragment containing TM 1–6 (nucleotides 1–620) of GLUT1. In a separate reaction, PCR1B, primer (TGCATCGTGCTGCCCTTCTGTCCTGAGAGCCCCCGA) containing sequence complimentary to nucleotides 600–619 of GLUT1 and 667–685 of GLUT4 (5′end of L6–7) and a reverse primer (containing a NotI restriction site) complimentary to the 3′ end of GLUT4 was used to generate a fragment containing the middle loop and TM 7–12 (nucleotides 667–1531) of GLUT4. In PCR2, the HindIII primer complimentary to the 5′ end of human GLUT1 and the reverse primer (containing a NotI restriction site) complimentary to the 3′ end of GLUT4 were used along with the products from PCR1A and B to generate an intermediate chimera containing sequence from nucleotides 1–620 of GLUT1 and 667–1531 of GLUT4. This intermediate chimera contained TM 1–6 of GLUT1 in frame with L6–7 and TM 7–12 of GLUT4. In PCR 3A, the HindIII primer complimentary to the 5′ end of human GLUT1 and a primer (CGCACCCAGGGGCAGCCTATCCTCATCGCTGTGGTC) complimentary to nucleotides 844–862 of GLUT4 and 815–829 of GLUT1 were used along with the product from PCR2 to generate a fragment containing TM 1–6 of GLUT1 (nucleotides 1–619) in frame with L6–7 (nucleotides 667–862) of GLUT4. In PCR 3B, a primer (CGCACCCAGGGGCAGCCTATCCTCATCGCTGTGGTC) complimentary to nucleotides 844–862 of GLUT4 and nucleotides 815–829 of GLUT1 were used with wild-type GLUT1 plasmid as a template to generate a fragment containing TM 7–12 (nucleotides 815–1480) of GLUT1. In the final reaction, PCR 4, HindIII primer complimentary to the 5′ end of human GLUT1 and the NotI primer complimentary to the 3′ end of GLUT1 were used with the products of PCR 3A and B as a template generating the final product, which contained TM 1–6 (nucleotides 1–619) of GLUT1, followed by L6–7 of GLUT4 (nucleotides 667–862), followed by TM 7–12 (nucleotides 815–1480) of GLUT1. This PCR product was digested with HindIII and NotI and ligated into the mammalian expression vector pcDNA 3.1+, cut with the same enzymes. Sequences were confirmed by sequencing and the chimera was transiently expressed in HEK 293 cells as described previously (Levine et al., 2005).
GLUT1 was covalently modified at accessible lysine residues (±ATP) using sulfosuccinimidyl-6-(biotinamido) hexanoate (sulfo-NHS- LC-biotin) as described previously (unpublished data). Detection of modified residues was achieved by limited proteolysis of labeled GLUT1 followed by RP-HPLC separation of fragments and ESI-MS/MS identification of peptides as described previously (unpublished data).
Human GLUT1 copurifies with red cell membrane lipids and is reconstituted into unsealed proteoliposomes upon detergent removal (Baldwin et al., 1979; Appleman and Lienhard, 1985; Sultzman and Carruthers, 1999). Although sealed proteoliposomes may be formed if exogenous lipid is added before detergent dialysis (Baldwin et al., 1982; Carruthers and Melchior, 1984; Zeidel et al., 1992), we employed purified human GLUT1 in unsealed proteoliposomes to ensure that added reagent has access to both surfaces of the lipid bilayer and thus full access to exo- and endofacial GLUT1 domains.
Exposure of GLUT1 proteoliposomes to trypsin (62.5:1 by mass) at 37°C results in the rapid loss of intact GLUT1 as judged by silver stain detection of peptides resolved by 10% SDS-PAGE. The time course of GLUT1 proteolysis is characterized by a rapid burst phase of proteolysis in which ~30% of GLUT1 is hydrolyzed with a τ of 4.5 s (Fig. 1 A). The remaining intact GLUT1 is proteolyzed with a τ of 2,500 s (see Fig. 1 B). Addition of ATP (0.1 to 4 mM always in the presence of 5 mM MgCl2) to the reaction reduces the size and rate of the proteolysis burst phase significantly (extent = 22%, τ = 56 s; Fig. 1 B). AMP, ADP, GTP, and CTP do not affect the rate or extent of trypsin-catalyzed GLUT1 proteolysis. ATP inhibits GLUT1 digestion at 60 s by 50% at 627 ± 268 μM ATP (Fig. 1 C).
To evaluate which GLUT1 domains were most susceptible to proteolysis, we examined GLUT1 fragmentation patterns by immunoblot analysis of GLUT1 digests using a series of GLUT1 peptide-directed IgGs. These included N-Ab (directed against GLUT1 amino acids 1–13), L6–7-Ab (directed against GLUT1 residues 217– 231), L7–8-Ab (GLUT1 residues 299–311), and C-Ab (residues 480–492). ATP (4 mM) reduces the efficiency of GLUT1 digestion (100:1 GLUT1:trypsin by mass, 20°C, 30 min reaction) as detected by immunoblot using IgGs specific to each of these GLUT1 subdomains (Fig. 2 A). For example, the C-Ab panel indicates the absence of higher molecular weight C-Ab–reactive fragments when GLUT1 proteolysis proceeds in the absence of ATP. When ATP is included, however, both high and low molecular weight C-Ab–reactive fragments are visible and more abundant, suggesting a general slowing of proteolysis of both intact and partially cleaved GLUT1. To identify protected peptides, we aligned each GLUT1 peptide detected by immunoblot analysis to GLUT1 sequence using two criteria: (1) the fragment must contain the reactive epitope, and (2) the mass (electrophoretic mobility) of the fragment must be consistent with previously detected GLUT1 tryptic cleavage sites (unpublished data). The trypsin-accessible sites observed here are generally consistent with the surface accessibility maps produced by systematic cysteine scanning mutagenesis of GLUT1 (Mueckler and Makepeace, 2006).
Fig. 2 B summarizes potential alignments of protected peptides. GLUT1 is most susceptible to tryptic digestion at cytoplasmic loop 6–7 and at the C terminus. ATP protection is distributed across GLUT1 primary structure. ATP inhibition of trypsin-catalyzed GLUT1 proteolysis increases in a saturable manner with ATP concentration and is half maximal at ~400 μM ATP (n = 1; Fig. 2 C).
The results of the proteolysis experiments suggest that ATP promotes significant GLUT1 conformational changes. To determine whether this also occurs in intact GLUT1, we analyzed ATP-dependent changes in GLUT1 epitope accessibility by measuring peptide-directed IgG binding to membrane-resident GLUT1. GLUT1 proteoliposomes were adsorbed to ELISA wells and then incubated ± ATP and peptide-directed IgGs. As reported previously (Carruthers and Helgerson, 1989), ATP inhibits C-Ab binding to GLUT1 proteoliposomes (Fig. 3 A; Table I). This is also observed with C-Ab binding to unsealed red cell membranes (Fig. 3 B) and in membranes isolated from HEK cells heterologously expressing wild-type human GLUT1 (Fig. 3 C). When HEK293 cells are transfected with a GLUT1–GLUT4 chimera in which the GLUT1 large middle loop (L6–7) is replaced by equivalent GLUT4 sequence, ATP inhibition of C-Ab binding to the GLUT1 C terminus of the chimera is lost (Fig. 3 D; Table I). Analysis of equilibrium C-Ab binding to these membranes indicates that ATP significantly reduces C-Ab binding to purified GLUT1, red cell–resident GLUT1 and wtGLUT1 (P < 0.001) but not to the GLUT1–GLUT4 loop 6 chimera (P > 0.1). This chimera is expressed efficiently (Fig. 3 D) and reaches the cell surface where it facilitates 2-deoxy-d-glucose transport. Untransfected HEK cells are characterized by Vmax and Km(app) for 2-deoxy-d-glucose uptake at 30°C of 1.2 ± 0.1 pmol/μg cell protein/min and 3.6 ± 1.4 mM, respectively. HEK cells transfected with wild-type GLUT1 (1.6 μg DNA per 106 cells) show significantly greater 2-deoxy-d-glucose uptake and are characterized by Vmax and Km(app) of 29.3 ± 9.4 pmol/μg cell protein/min and 3.6 ± 1.4 mM, respectively. Cells transfected with the loop 6–7 GLUT1–GLUT4 chimera (1.6 μg DNA per 106 cells) are characterized by Vmax and Km(app) for 2- deoxy-d-glucose uptake of 21.6 ± 2.6 μmol/106 cells/min and 1.7 ± 0.7 mM, respectively.
To understand whether this response is restricted to the GLUT1 C terminus or more widespread, we examined the available peptide-directed IgGs for ability to bind to intact GLUT1 and for sensitivity of binding to ATP (Table II). ATP does not affect binding of -Ab, loop 2–3-Ab or loop 6–7-Ab to membrane-resident GLUT1 but does reduce loop 7–8-Ab and C-Ab binding to GLUT1 proteoliposomes. N-Ab and loop 8–9-Ab binding to native GLUT1 structure are undetectable, indicating that these epitopes are inaccessible in membrane-resident GLUT1.
We assessed whether ATP-dependent conformational changes in loop 6–7 and the C terminus are reflected at the level of specific amino acid side chains by analysis of ATP modulation of sulfo-NHS-LC-biotin covalent modification of loop 6–7 and C terminus lysine residues. GLUT1 proteoliposomes were preincubated in the presence or absence of 4 mM ATP before addition of a 20-fold molar excess of sulfo-NHS-LC-biotin to initiate GLUT1 labeling. This ratio of probe to GLUT1 produces significant labeling of GLUT1 cytoplasmic loop lysine residues without affecting the ability of GLUT1 to bind cytochalasin B (unpublished data). ATP reduces the extent of GLUT1 modification by ~40%, without slowing the reaction rate (Fig. 4 A). ATP inhibition of labeling is half maximal at 2 mM ATP (Fig. 4 B). AMP (0 to 4 mM) alters neither the rate nor the extent of GLUT1 modification but increases the amount of ATP required to inhibit GLUT1 modification. When the concentration dependence of ATP inhibition of GLUT1 modification is measured in the presence of 2 mM AMP, the effects of ATP are half maximal at 3.8 mM. Assuming AMP competitively antagonizes ATP inhibition of modification, Ki(app) for AMP antagonism of ATP modulation of GLUT1 is 2.2 mM (Fig. 4 B).
GLUT1 lysine residues whose accessibility to sulfo-NHS-LC-biotin is specifically affected by ATP were identified by ESI-MS/MS analysis of labeled GLUT1. After GLUT1 biotinylation in the presence of 4 mM AMP (control) or ATP and subsequent tryptic digestion/ESI-MS-MS, we identified all peptides originating from a specific GLUT1 region and quantitated each peak area. The peak areas of biotinylated peptides in a denoted region were summed and expressed as the fraction of all peptides (labeled and unlabeled) in that region. Fig. 5 shows a representative analysis of lysine 245 modification.
To simplify the analysis, cytoplasmic GLUT1 lysine residues were grouped into four regions based on sequence location (see Fig. 6): region 1 contains K225, K229, and K230; region 2 contains K245; region 3 comprises K255 and K256; region 4 contains K477. Regions 1, 2, and 3 are located in cytoplasmic loop-6–7. Region 4 lies in the GLUT1-C terminus.
Region 1 shows no discernible change in the quantity of labeled peptide in the presence of ATP (Fig. 6). Regions 2, 3, and 4 show decreased lysine modification in the presence of ATP (Fig. 6). ATP reduces labeling of K245 of region 2 by 38% (see Fig. 5 A [AMP] and Fig. 5 B [ATP]). Similarly, region 3 displays a 24% reduction and labeling of K477 of region 4 shows a 26% decrease in the presence of ATP (Fig. 6). Overall, ATP reduces labeling of lysine residues found within the C-terminal half of L6–7 and the C terminus of GLUT1 by ~30%.
These findings reinforce the hypothesis that the GLUT1 C terminus and loop 6–7 undergo significant conformational change in the presence of cytoplasmic ATP. We asked, therefore, how complexation of the C terminus using intracellular C-Ab might impact ATP modulation of sugar transport. ATP increases the rate of sugar transport at subsaturating 3MG because at these sugar concentrations the rate of transport, v, is given by:
where [S] is the sugar concentration. ATP reduces Vmax and Km(app) for net 3MG uptake by red cell ghosts at 4°C by 8- and 18-fold, respectively (Helgerson et al., 1989), thereby increasing the ratio Vmax/ Km(app) and the rate of subsaturating transport by 2.3-fold (Table III).
Incorporation of GLUT1 C-Ab into resealed erythrocyte ghosts severely blunts ATP stimulation of 3MG uptake at 4°C (Table III). Preimmune serum and loop 6–7-Ab are without effect on ATP action.
Exogenous, intracellular GLUT1 C-terminal peptide (EELFHPLGADSQV), but not 1D4 peptide (ETQSVAPA, a rhodopsin peptide), mimics the ability of ATP to stimulate sugar transport and acts synergistically with ATP (Table III). In the absence of ATP, C-terminal peptide triples the rate of 3MG uptake and is half maximally active at 9 μM.
This study examines the structural basis of GLUT1 modulation by cytoplasmic ATP and AMP. Our findings suggest that the GLUT1 cytoplasmic C terminus and the cytoplasmic loop linking TMs 6 and 7 interact in an ATP- dependent fashion to modulate transport.
Human GLUT1 responds acutely to cellular ATP depletion with enhanced sugar transport capacity (Jung et al., 1971; Taverna and Langdon, 1973; Weiser et al., 1983; Cloherty et al., 1996; Levine et al., 2002). This effect is direct (Hebert and Carruthers, 1986; Carruthers and Helgerson, 1989), does not require ATP hydrolysis, and is antagonized by intracellular H+, AMP, and ADP (Carruthers and Helgerson, 1989; Helgerson et al., 1989). GLUT1 residues 301–364 (TM8 through TM 9) form at least one element of the nucleotide binding domain (Levine et al., 1998) and site-directed mutagenesis of loop 8–9 residues (E329A or R332A/R333A) produces an ATP-insensitive, dominant-negative transporter in HEK cells (Levine et al., 2002). The molecular mechanism of GLUT1 modulation by ATP is unknown but may involve GLUT1 C terminus conformational change (Carruthers and Helgerson, 1989).
Our results demonstrate that ATP–GLUT1 interactions promote global changes in GLUT1 conformation. ATP-dependent GLUT1 protection against trypsin digestion, ATP inhibition of IgG binding to GLUT1 subdomains, and ATP protection of specific cytoplasmic lysine residues against modification by NHS-biotin confirm that the ATP–GLUT1 complex is conformationally distinct from its ATP-free counterpart. ATP inhibits IgG binding to the C terminus and to exofacial loop 7–8 of membrane-resident transporter. ATP does not affect the extent of covalent modification of loop 6–7 lysine residues 225, 229, and 230 or binding of an antibody directed against residues 217–231, indicating that exposure of this flexible loop subdomain to cytoplasmic water is unchanged by ATP. This ATP-insensitive region contains a putative ATP binding motif (KSVLK; residues 225–229) that is also present in GLUT4 loop 6–7 (KSLK), lending further support to the hypothesis that this domain is not involved in ATP regulation of GLUT1. Labeling of loop 6–7 lysine residues 245, 255, and 256 is inhibited by ATP, and substitution of GLUT1 loop 6–7 with the equivalent GLUT4 domain greatly diminishes ATP inhibition of IgG binding to the chimera's GLUT1 C terminus. These observations suggest that GLUT1 requires sequence present in GLUT1 loop 6–7 in order to occlude the C terminus from exogenous IgG. Two of the three ATP-sensitive lysine residues of loop 6–7 are retained in GLUT4 loop 6–7 primary structure. This suggests that loop 6–7 ATP-protected lysine residues do not play a direct role in ATP regulation of transport (e.g., ATP binding or salt bridge formation with C-terminal residues). Rather, these residues may reflect altered conformational accessibility of loop 6–7.
AMP is without effect on GLUT1 loop 6–7 NHS-biotinylation but antagonizes ATP inhibition of labeling. AMP also antagonizes ATP inhibition of C-Ab binding to GLUT1 (Carruthers and Helgerson, 1989). This behavior confirms that the functional antagonism between ATP and AMP is replicated at the molecular level and that ATP modulation of GLUT1 conformation results in modulated transport function.
We propose that GLUT1–ATP association promotes conformational change within the cytoplasmic GLUT1 C terminus and loop 6–7, resulting in their interaction and alignment with sugar translocation pathway (Fig. 7). This alignment/interaction limits sugar release into cytoplasm and is supported by several observations. (a) Sequestration of the GLUT1 C terminus by exogenous intracellular C-terminal IgGs blunts ATP modulation of sugar transport. (b) Exogenous, intracellular C-terminal peptide mimics ATP action of sugar transport. (c) Substitution of GLUT1 loop 6–7 with GLUT4 loop 6–7 (40% sequence identity with GLUT1 over residues 249–259) prevents ATP inhibition of IgG binding to the C terminus.
The action of ATP is thus extrinsic to the membrane-spanning translocation pathway and intrinsic to cytoplasmic domains. This hypothesis is further supported by rapid quench flow studies of erythrocyte glucose transport, demonstrating that rapid translocation of sugar through the sugar translocation pathway is unaffected by ATP, but sugar release from the pathway into cytosol is regulated by ATP (Blodgett and Carruthers, 2005; Leitch and Carruthers, 2007). Loop 6–7 and the C terminus may be necessary but not sufficient for GLUT1 regulation because GLUT1 loop 8–9 mutagenesis causes the loss of GLUT1 ATP sensitivity but retention of GLUT1 ATP binding (Levine et al., 2002).
The parallels between channel inactivation and inhibition of GLUT1 via cytosolic “inactivation domains” are striking. Inactivation of multisubunit, voltage-dependent K+ channels and large-conductance, Ca2+-activated K+ channels involves insertion of a cytosolic N-terminal peptide segment into the ion permeation pathway, thereby obstructing ion flux (Kobertz et al., 2000).
Less is known about the intrinsic regulation of other members of the GLUT family of proteins or whether the broader family of MFS proteins coopt a similar regulatory mechanism. MFS proteins contain loop 6–7 domains ranging from 30 to 65 amino acids (Weinglass and Kaback, 2000). The LacY and GlpT structures indicate that loop 6–7 and the C terminus cytoplasmic domains do not impede access to the substrate export site (Abramson et al., 2003; Huang et al., 2003). However, loop 6–7 and the C termini of these structures are considerably shorter than equivalent domains in the mammalian sugar porters. Progressive truncation of GLUT1 loop 6–7 results in loss of transport function followed by loss of GLUT1 expression (Monden et al., 2001). The former result may reflect the loss of residues crucial to transport. The latter may reflect impaired cotranslational insertion/folding (Weinglass and Kaback, 2000).
The available evidence suggests that the GLUT1 and GLUT4 C termini are essential for transport but contain isoform-specific subsequence that suppresses intrinsic GLUT activity. Truncation of the GLUT1 C terminus by 12 amino acids is without effect on GLUT1 activity (Lin et al., 1992), whereas removal of 37 residues abolishes GLUT1 intrinsic activity (Oka et al., 1990). Substitution of the GLUT1 C-terminal tail with the equivalent GLUT4 sequence stimulates GLUT1 intrinsic activity twofold but abolishes accelerated exchange transport (Dauterive et al., 1996). The C-terminal 20 residues of GLUT4 suppress GLUT4 intrinsic activity and deletion or replacement of this region with equivalent GLUT1 sequence increases GLUT4 activity fourfold (Dauterive et al., 1996). C-terminal truncation of the organic anion transporter OAT1 reduces OAT1 intrinsic activity (Xu et al., 2006). These observations support the hypothesis that the C-terminal domains of the GLUT sugar porter and the organo anion porter MFS subfamilies are essential for transport function and that GLUT1 and GLUT4 C termini contain isoform-specific subsequence that suppresses intrinsic GLUT activity. Insulin unmasks the GLUT4 C terminus in L6 muscle cells (Ishiki et al., 2005), adipocytes (Smith et al., 1991), and muscle (Wang et al., 1996) and desuppresses GLUT4 intrinsic activity (Michelle Furtado et al., 2003). The present study demonstrates that ATP depletion unmasks the GLUT1 C terminus, resulting in desuppression of GLUT1 intrinsic activity.
Sugar transport in astrocytes, vascular smooth muscle cells, basal cardiomyocytes, and cells of the reticuloendothelial system is mediated by GLUT1 (Hruz and Mueckler, 2001; Mann et al., 2003; Simpson et al., 2007). This may result in ATP-sensitive glucose transport in these tissues whereby cellular sugar transport capacity is increased in response to ATP depletion. What advantage is obtained by GLUT1 regulation in erythrocytes and endothelial cells where transport exceeds metabolic demand? The benefit may not be realized by the transporting cell but rather by the cells to which glucose is subsequently transferred.
Glucose transfer from blood to brain proceeds across capillary endothelial cells that form the blood–brain barrier (Takata et al., 1990). Endothelial cells constitute only 0.1% of brain mass yet almost all glucose used by cerebral neurons and astrocytes enters the brain by GLUT1-mediated, transendothelial cell transport (Harik, 1992). Cerebral endothelial cell GLUT1 content is adaptively up-regulated during long-term hypoglycemia and hypoxia to meet increased astrocytic and neuronal demand for glycolytic ATP (Mann et al., 2003). Blood–brain barrier glucose transport is acutely stimulated during seizure-induced stimulation of neuronal and astrocytic glycolysis (Cornford et al., 2000). Regulation of barrier GLUT1 is, therefore, an important physiologic adaptation and deficits in GLUT1 expression or hypoglycemia inevitably impact human cognitive function and development as observed in GLUT1-deficiency syndrome (Pascual et al., 2004) and hypoglycemia (Guettier and Gorden, 2006).
The very high GLUT1 content of higher primate and odontocete erythrocytes may contribute to glucose transfer from blood to tissue (Craik et al., 1998) by allowing human red cells to exchange upwards of 82–98% of intracellular glucose with serum in the 2–4 s required for an erythrocyte to transit the capillary bed of peripheral tissues (Honig et al., 1977; Regittnig et al., 2003). Regulation of erythrocyte glucose transport therefore may permit controlled expansion/contraction of the blood glucose space available for exchange with the interstitium of glycolytically active tissues.
GLUT1–ATP interaction restructures GLUT1 cytoplasmic loop 6–7 and C-terminal domains, resulting in their interaction and selective inhibition of sugar release from the transmembrane sugar translocation pathway into cytosol. This enables GLUT1 to respond to cytoplasmic energy charge with low or high capacity glucose transport. Interactions between cytoplasmic domains may be important in the regulation of other mammalian MFS transport proteins.
This work was supported by National Institutes of Health grants DK 36081 and DK 44888 and by ADA 1-06-IN-04 (Gail Patrick Innovation Award supported by a generous gift from the estate of Gail Patrick).
Lawrence G. Palmer served as editor.
D.M. Blodgett and J.K. De Zutter contributed equally to this work.
Abbreviations used in this paper: CB, cytochalasin B; GLUT1, human erythrocyte glucose transport protein; ESI-MS, electrospray ionization mass spectrometry; HRP, horseradish peroxidase; MFS, major facilitator superfamily; PBS-T, PBS containing Tween; sulfo-NHS-LC-biotin, sulfosuccinimidyl-6-(biotinamido) hexanoate.