The transport of glucose across the plasma membrane is mediated by members of the glucose transporter family. In this study, we investigated glucose uptake through the yeast hexose transporter 1 (Hxt1) by measuring incorporation of 2-NBDG, a non-metabolizable, fluorescent glucose analog, into the yeast Saccharomyces cerevisiae. We find that 2-NBDG is not incorporated into the hxt null strain lacking all glucose transporter genes and that this defect is rescued by expression of wild type Hxt1, but not of Hxt1 with mutations at the putative glucose-binding residues, inferred from the alignment of yeast and human glucose transporter sequences. Similarly, the growth defect of the hxt null strain on glucose is fully complemented by expression of wild type Hxt1, but not of the mutant Hxt1 proteins. Thus, 2-NBDG, like glucose, is likely to be transported into the yeast cells through the glucose transport system. Hxt1 is internalized and targeted to the vacuole for degradation in response to glucose starvation. Among the mutant Hxt1 proteins, Hxt1N370A and HXT1W473A are resistant to such degradation. Hxt1N370A, in particular, is able to neither uptake 2-NBDG nor restore the growth defect of the hxt null strain on glucose. These results demonstrate 2-NBDG as a fluorescent probe for glucose uptake in the yeast cells and identify N370 as a critical residue for the stability and function of Hxt1.
Citation: Roy A, Dement AD, Cho KH, Kim J-H (2015) Assessing Glucose Uptake through the Yeast Hexose Transporter 1 (Hxt1). PLoS ONE 10(3): e0121985. https://doi.org/10.1371/journal.pone.0121985
Academic Editor: Hernâni Gerós, University of Minho, PORTUGAL
Received: December 1, 2014; Accepted: February 6, 2015; Published: March 27, 2015
Copyright: © 2015 Roy et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Data Availability: All relevant data are within the paper.
Funding: The work was supported by National Institutes of Health Grant GM087470. The funder had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing interests: The authors have declared that no competing interests exist.
Metastasized tumor cells metabolize large amounts of glucose through glycolysis and produce copious amounts of lactic acid even in the presence of oxygen [1,2]. This phenomenon, termed the Warburg effect, is a hallmark of cancer . The well-established elevated glucose consumption of malignant tissue forms the basis of the clinical imaging of cancer, [18F] FDG-PET (positron emission tomography) . The budding yeast Saccharomyces cerevisiae, like cancer cells, prefers to ferment rather than oxidize glucose [5,6]. Since energy generation by fermentation of glucose is inefficient, the yeast cells consume the available glucose vigorously by enhancing glucose uptake through glucose transporters [7,8].
Glucose uptake is measured using non-metabolizable glucose analogs such as 3-O-methylglucose (3-OMG) and 2-deoxyglucose (2-DG) [9–11]. After taken up by cells, 3-OMG cannot be phosphorylated by hexokinase; 2-DG is phosphorylated to 2-DG-6-phosphate (2-DG-6-P) but cannot be metabolized further. Hence, radioisotope-labeled 3-OMG and 2-DG have been widely used to measure glucose uptake. Recently, however, some inherent disadvantages associated with using radioactive glucose analogs have led to the development of improved methods for measuring glucose uptake using nonradioactive substances. 2-[N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)amino]-2-deoxy-D-glucose (2-NBDG) is a fluorescent derivative of 2-DG, which is converted to its phosphate form and accumulates in cells . Due to its non-metabolizable and fluorescent properties, 2-NBDG has been proven useful for evaluating glucose uptake in mammalian cells [13–15].
S. cerevisiae possesses at least six members of the glucose transporter family (Hxt1, 2, 3, 4, 6 and 7) with different affinities for glucose in order to cope with environmental changes in glucose availability [16,17]. Expression of several HXT genes (HXT1-4) is repressed by the Rgt1 repressor, which recruits the general corepressor complex Ssn6–Tup1 and the HXT corepressor Mth1 to the HXT promoters in the absence of glucose [18–26]. The yeast cells employ three major glucose signaling pathways—Rgt2/Snf3, AMPK, and cAMP-PKA—that operate in a highly regulated and cooperative manner to bring about glucose-induction of HXT gene expression by inactivating the Rgt1 repressor [7,27,28]. The yeast glucose transporters are regulated at both transcriptional and posttranslational levels: HXT genes are induced by the aforementioned mechanisms; Hxt proteins undergo endocytosis and vacuolar degradation when they are not needed [29, 30].
In this study, using 2-NBDG, we investigated glucose uptake through the yeast hexose transporter 1 (Hxt1). Our study was focused on whether 2-NBDG can be used as a proxy for glucose uptake in S. cerevisiae and whether 2-NBDG is transported through the putative glucose-binding residues, inferred from human glucose transporters (Gluts). Our results show that Hxt1 transports 2-NBDG in a mechanism similar to Gluts and, furthermore, that some of the putative glucose-binding residues of Hxt1 are involved in endocytosis. Also discussed is the possible roles of these residues in the stability and function of Hxt1.
2-NBDG as a fluorescent probe for glucose uptake in S. cerevisiae
Glucose uptake assays in yeast heavily rely on the use of radioactive glucose or its derivatives, which suffer from numerous problems inherent in the use of radioactive substances. This hampers the development of facile methods for measuring glucose transport activity. 2-NBDG is often used as a fluorescent probe for glucose uptake in mammalian cells [13–15]. However, limited numbers of studies, only two early works, have shown that 2-NBDG can be incorporated into yeast cells [31,32]. To directly determine whether 2-NBDG is transported into the yeast S. cerevisiae through the glucose transport system, the yeast cells lacking all HXT (glucose transporter) genes  were transformed with an empty plasmid or with a plasmid encoding Hxt1-HA. The resulting transformants were first grown in SC-glycerol/ethanol medium till mid log phase and shifted to the same medium containing 60 μM of 2-NBDG. The intensity of the fluorescence signal was used to measure for the concentration of 2-NBDG transported into the cell. The fluorescence signal was not noticeably observed in the hxt null strain, but was markedly increased in the strain expressing the Hxt1 glucose transporter (Fig. 1A). Glucose competition assay was conducted to study substrate specificity by varying glucose concentrations with a fixed concentration of 2-NBDG. The results show that the fluorescence intensity is inversely correlated with increasing concentrations of glucose in the medium and that treatment of cells with 0.5 mM glucose resulted in ~ 50% decrease in the uptake of 2-NBDG (Fig. 1B and 1C). These results suggest that 2-NBDG is transported into S. cerevisiae through glucose transporters and that glucose uptake activity in yeast can be directly evaluated by measuring the incorporation of 2-NBDG into the cells.
(A) Yeast cells (hxtΔ) expressing either empty vector or Hxt1-HA were first grown in SC-5% glycerol + 2% ethanol medium to mid log phase and shifted to the same medium containing 60 μM of 2-NBDG and incubated for 30 min. Fluorescence microscopy images (left panel) and quantification of relative 2-NBDG fluorescence (right panel, **P < 0.001) were shown. (B) Yeast cells (hxtΔ) expressing Hxt1-HA were grown as described in A. For glucose competition assay, glucose was added to different concentrations (mM) as shown and 2-NBDG uptake was analyzed by fluorescence microscopy. (C) Quantification of relative 2-NBDG uptake in glucose competition assay was shown. FL: Fluorescence, DIC: Differential Interference Contrast.
The yeast Hxt1 glucose transporter transports glucose in a similar mechanism as the human Glut1
Extensive mutational analyses of human glucose transporters and crystal structures of some sugar transporters have identified residues important for glucose transport [34–41]. Since yeast and human glucose transporters are highly conserved, we examined whether yeast glucose transporters transport glucose in a similar mechanism as human glucose transporters (Fig. 2A and 2B). To this end, we first mutated the putative glucose-binding residues of Hxt1 corresponding to the residues of human Glut1—Q209, Q335, Q336, S363, N370, and W473 —to alanine individually and tested the resulting mutant Hxt1 proteins for their ability to transport glucose. While the expression patterns of most of mutant Hxt1 proteins are similar to those of wild type Hxt1, the protein levels of Hxt1N370A and Hxt1W473A are constitutively high as compared with those of wild type Hxt1 (Fig. 2C). The hxt null strain is unable to grow on glucose as a sole carbon source and this defect is fully complemented by expression of wild type Hxt1 . To test the mutant Hxt1 proteins for their ability to transport glucose, the hxt null mutant strain was transformed with plasmids encoding the mutant Hxt1-HA transporters and scored for growth on glucose medium. We observe that the growth of the hxt null strain is not restored by expression of most of mutant Hxt1 proteins or only partially restored by expression of Hxt1S363A-HA or Hxt1W473A-HA (Fig. 2D). Thus, these results demonstrate that the putative glucose-binding residues—in particular Q209, Q335, Q336 and N370—may be critical for glucose uptake.
(A) Cartoon for the arrangement of the 12 transmembrane helices and the proposed model of the exofacial glucose-binding sites of Hxt1 protein as viewed from the outside of the cell. For the sake of simplicity, all transmembrane helices were drawn as perfect helices perpendicular to the plane of the membrane. Amino acid residues were grouped according to their chemical properties of side chains (red: neutral polar, purple: acidic or basic polar, green: neutral nonpolar and blue: hydrophobic). Putative glucose-binding residues are shown. (B) Sequence alignment of segments of glucose transporters of yeast (Hxt1-7 and Gal2) and human (Glut1-5) showing proposed amino acids (highlighted in box) predicted to interact with glucose. (C) Western blot analysis of Hxt1-HA levels at the plasma membrane. Yeast cells (WT) expressing indicated Hxt1-HA proteins were grown in SC-2% glucose (Glu) medium to mid log phase and shifted to SC medium containing 2% galactose (Gal) for 6 hr. Membrane fractions were analyzed using anti-HA antibody. Actin was served as loading control. (D) Yeast cells (hxtΔ) expressing either empty vector or indicated Hxt1-HA proteins were spotted on 2% glucose plate supplemented with Antimycin-A (1μg/ml). The first spot of each row represents a count of 5 x 107 cell/ml, which is diluted 1:10 for each spot thereafter. The plate was incubated for 3 days and photographed.
Hxt1 with mutations at the putative glucose-binding residues cannot transport 2-NBDG
Given that 2-NBDG is transported into the yeast cells through glucose transporters (Fig. 1), we examined whether the putative glucose-binding residues are involved in 2NBDG uptake. To this end, the hxt null strain was transformed with plasmids encoding the mutant Hxt1 proteins, and the resulting transformants were treated with 2-NBDG, as described above. 2-NBDG fluorescence is hardly detectable in cells expressing Hxt1Q209A, Hxt1Q336A and Hxt1N370A; by contrast, the fluorescence signals in cells expressing Hxt1Q335A, Hxt1S363A and Hxt1W473A are reduced by 50%-80%, compared with those in cells expressing wild type Hxt1 (Fig. 3A). Thus, the residues Q209, Q336 and N370 seem to be critical for 2-NBDG uptake, consistent with the above finding that these three residues may play a major role in glucose transport.
(A) Yeast cells (hxtΔ) expressing either empty vector or indicated Hxt1-HA proteins were grown as described in Fig. 1A. 2-NBDG uptake by the yeast cells was analyzed by fluorescence microscopy. FL: Fluorescence, DIC: Differential Interference Contrast. (B) Quantification of relative 2-NBDG uptake by the yeast cells was shown (*P < 0.05).
Mutation at the putative glucose-binding residues of Hxt1 does not affect its localization to the plasma membrane
Next, we determined whether mutation at the putative glucose-binding residues of Hxt1 affect its targeting to the plasma membrane by analyzing subcellular localization of GFP-fused mutant Hxt1 proteins. Expression of wild type Hxt1-GFP fully complemented the growth defect of the hxt null strain on glucose, suggesting that fusion of the GFP moiety to Hxt1 does not interfere with its function. However, the growth defect was not rescued by expression of the mutant Hxt1-GFP transporters or only partially rescued by expression of Hxt1S363A-GFP or Hxt1W473A-GFP (Fig. 4A). It should be noted that basal levels of glucose uptake in the hxt null strain expressing Hxt1-HA fusions were a little higher, compared with those in the strain expressing corresponding GFP fusions. This is presumably due to different expression levels of the two constructs; Hxt1-HA expressed from a 2μ-based, high copy number plasmid (Fig. 2) and Hxt-GFP expressed from a CEN-based, low copy plasmid (Fig. 4). Western blot analysis indicates that expression patterns of Hxt-GFP proteins are similar to those of Hxt-HA proteins: the protein levels of Hxt1Q209A-GFP, Hxt1Q335A-GFP, Hxt1Q336A-GFP and Hxt1S363A-GFP are high in glucose-grown cells but low in galactose (glucose-free)-grown cells; by contrast, those of Hxt1N370A-GFP and Hxt1W473A-GFP are constitutively high in both glucose and galactose-grown cells (Fig. 4B). In addition, fluorescence microscopy reveals that wild type Hxt1-GFP is localized to the plasma membrane in response to glucose, whereas Hxt1N370A-GFP and Hxt1W473A-GFP are found at the plasma membrane constitutively, regardless of the presence and absence of glucose (Fig. 4C). Of note, both HA and GFP fusions of Hxt1N370A are expressed at high levels in both glucose-containing and glucose-free medium but unable to rescue the growth defect of the hxt null strain. Thus, the expression patterns of mutant Hxt1 proteins are not correlated with their ability to uptake glucose.
(A) Yeast cells (hxtΔ) expressing either empty vector or indicated Hxt1-GFP proteins were spotted on 2% glucose plate supplemented with Antimycin-A (1μg/ml) as described in Fig. 2D. The plate was incubated for 3 days and photographed. (B) Western blot analysis of Hxt1-GFP levels at the plasma membrane. Yeast cells (WT) expressing indicated Hxt1-GFP proteins were grown as described in Fig. 2C. Membrane fractions were analyzed by Western blotting with anti-GFP antibody. Actin was served as loading control. (C) Yeast cells (WT) expressing indicated Hxt1-GFP proteins were grown as described in Fig. 2C and were analyzed by fluorescence microscopy (top). Relative GFP fluorescence in the plasma membrane was quantified (bottom). Relative GFP fluorescence intensities were plotted with the fluorescence of WT cells (2% glucose condition) set to 100%. The data represented were averages of at least 50 cell counts with error bars representing standard deviations (S.D).
Some of the putative glucose binding residues of Hxt1 are required for its endocytosis
We have recently shown that Hxt1 is endocytosed and degraded in the vacuole in response to glucose starvation . Thus, it is conceivable that the reduced protein levels of Hxt1 in glucose-starved cells (e.g., galactose-grown cells) may be due to endocytosis (Figs. 2 and 4). To validate this idea, we examined endocytosis of the mutant Hxt1 proteins (Hxt1S363A and Hxt1N370A) in a strain lacking End3, involved in the internalization step of endocytosis. We found that, in the end3Δ strain, the protein levels of wild type Hxt1 and mutant Hxt1S363A transporters are not reduced in glucose-starved cells (Fig. 5A) and that these proteins constitutively localize to the plasma membrane (Fig. 5B and 5C). Thus, like wild type Hxt1, Hxt1S363A (and presumably also Hxt1Q209A, Hxt1Q335A, and Hxt1Q336A) may be internalized and targeted to the vacuole for degradation in glucose-starved cells. By contrast, Hxt1N370A protein levels are constitutively high in both wild type and end3Δ strains, suggesting that Hxt1N370A (and presumably also Hxt1W473A) may not undergo endocytosis. Thus, these results identify N370 as a critical residue for the stability and function of Hxt1.
Yeast cells (WT and end3Δ) expressing indicated Hxt1-GFP proteins were grown as described in Fig. 2D. (A) Western blot analysis of Hxt1-GFP levels at the plasma membrane and (B) fluorescence microscopy of Hxt1-GFP proteins were shown. Actin was served as loading control. (C) Quantification of relative GFP fluorescence in the plasma membrane was performed as described in Fig. 4E. *Results of the Western blot and fluorescence microscopy analysis of wild type Hxt1-GFP protein in yeast cells (WT) depicted in Fig. 4C were shown for comparison.
In this study, using 2-NDBG uptake assay in combination with yeast growth restoration assay, we assessed glucose uptake through the yeast hexose transporter Hxt1 and found that it may transport glucose in a mechanism similar to but subtly different from that of human Glut1 (Fig. 3B). Previous evidence suggests that Q161, Q282 and W412 of Glut1 may be involved in glucose-binding. Substitution of an asparagine at Q161significantly reduces the affinity for the substrate-binding site of a nontransported glucose analog, suggesting that this residue is critical for transport activity and exofacial ligand binding . Q282 of Glut1 appears to have an important role in exofacial substrate binding but not to be critical in glucose transport, implicating the involvement of this residue in glucose-induced conformational change . Our results show that Hxt1 with mutations at Q209 and Q335, corresponding to Q161 and Q282 in Glut1, respectively, is unable to restore the growth defect of the hxt null strain on glucose and to transport 2-NBDG into the yeast cells (Fig. 3A). Interestingly, W412 of Glut1 appears to be critical for substrate binding by interacting with the C-6 position of the pyranose ring [34,46]; however, its corresponding residue in Hxt1 (W473) is not critically required for glucose uptake (Figs. 2D and 4A). A striking feature of glucose transporters is their distinctive substrate specificity: the human Glut1 transports glucose, galactose but not fructose , whereas the yeast Hxt1 transports glucose and fructose, but not galactose [48, 49]. This is presumably due to differences in the sequences within and outside the glucose-binding pockets of the yeast and human glucose transporters. For example, Q282, Q283, N288, N317 and N415 of Glut1 are involved in interaction with glucose through hydrogen bonds; the first four residues are conserved in yeast transporters, whereas the aromatic asparagine at 415 in Glu1 is replaced with the non-aromatic glycine (at position 476) in yeast Hxts . Therefore, it is conceivable that this difference, in part, may account for the different roles of W412 (Glut1) and W473 (Hxt1) in transporting glucose.
Yeast glucose transporters are removed from the plasma membrane and targeted to the vacuole for degradation when they are not needed. The high affinity glucose transporters Hxt2 and Hxt6/7 are endocytosed and degraded in high glucose-grown cells, whereas the low affinity glucose transporters Hxt1 and Hxt3, in glucose-starved cells [30,50–52]. We have also recently shown that the high affinity glucose sensor Snf3 is inactivated in a similar mechanism as Hxt2 and Hxt6/7 and that the low affinity glucose sensor Rgt2, as Hxt1 and Hxt3 . These observations suggest that turnover of glucose transporters and sensors might be associated with their inability to bind glucose. However, the results in this study show that the mutant Hxt1 transporters unable to restore the growth defect of the hxt null strain on glucose—Hxt1Q209A, Hxt1Q335A, Hxt1Q336A and Hxt1N370A—are endocytosed and degraded in response to glucose starvation except Hxt1N370A (Fig. 4). Furthermore, both Hxt1S363A and Hxt1W473A have a reduced ability to complement the growth defect of the Hxt null strain, but only Hxt1W473A is resistant to endocytosis (Fig. 5). Thus, it remains to be determined whether Hxt1 stability is related to its ability to bind glucose.
N370 and W473—N370 in particular—may have a pivotal role in both glucose transport and Hxt1 protein turnover. Hxt1 is ubiquitinated by the Rsp5 ubiquitin ligase prior to endocytosis , and often phosphorylation is a signal for ubiquitination [54–56]. We surmise that N370A and W473A mutations may induce conformational changes in Hxt1 and that the resulting mutant Hxt1 proteins are neither phosphorylated nor ubiquitinated. Consistently, a mutation of N370 of Hxt7 or N376 of Gal2, corresponding to N370 of Hxt1, to F (phenylalanine) abolishes hexose transport completely . It is also noted that residues involved in the endocytosis of the Hxt1 transporter are also needed for the transport of 2-NBDG (Figs. 2 and 3), raising a possibility that 2-NBDG binds to Hxt1 and then enters the cells by endocytosis. Therefore, 2-NBDG, like glucose, is likely transported into the cells through the glucose transporter system, but the possibility of 2-NBDG uptake by endocytosis cannot be ruled out.
Materials and Methods
Yeast strains and Growth Conditions
The Saccharomyces cerevisiae strains used in this study are BY4742 (WT, Mata his3Δ1 leu2Δ0 ura3Δ0 met15Δ), EBY.S7 (MATα hxt1-17Δgal2Δagt1Δstl1Δleu2-3,112 ura3-52 trp1-289 his3-Δ1 MAL2–8c SUC2 hxtΔfgy1-1 ) and KFY127 (Matα his3Δ1 leu2Δ0 lys2Δ0 ura3Δ0 end3::KanMX ). Yeast cells were grown in YP (2% bacto-peptone, 1% yeast extract) and SC (synthetic yeast nitrogen base medium containing 0.17% yeast nitrogen base and 0.5% ammonium sulfate) media supplemented with the appropriate amino acids and carbon sources.
The plasmids used in this study are listed in Table 1. Plasmids containing Hxt1-GFP, Hxt1 (Q209A)-GFP, Hxt1 (Q335A)-GFP, Hxt1 (Q336A)-GFP, Hxt1 (S363A)-GFP, Hxt1 (N370A)-GFP and Hxt1 (W473A)-GFP were constructed by ‘gap repair’ of BamHI-EcoRI linearized pUG35 vector. Hxt1-HA plasmid was mutagenized by QuikChange Site-Directed Mutegenesis kit (Stratagene) following manufacturer’s protocol to generate Hxt1 (Q209A)-HA, Hxt1 (Q335A)-HA, Hxt1 (Q336A)-HA, Hxt1 (S363A)-HA, Hxt1 (N370A)-HA and Hxt1 (W473A)-HA.
Yeast Membrane Preparation, Western Blotting and Protein Half-life Measurement
Membrane enriched fractions were essentially prepared as described previously , with some minor modifications. Briefly, after washing with phosphate buffer, pH 7.4 containing 10 mM sodium azide, the cell pellet was resuspended in ice cold membrane isolation buffer (100 mM Tris-Cl, pH 8, 150 mM NaCl, 5 mM EDTA) containing 10 mM sodium azide, protease and phosphatase inhibitors and vortexed with acid-washed glass beads. After diluting the samples with the same buffer, unbroken cells and debris were removed by centrifugation and membrane enriched fraction was collected by centrifuging the samples at 12,000 rpm for 40 min at 4° C. The pellets were resuspended in the aforementioned buffer containing 5M urea and incubated for 30 min on ice and further centrifuged at 12,000 rpm for 40 min at 4° C. The proteins were precipitated with 10% TCA, neutralized with 20 μl of 1M Tris base and finally dissolved in 80 μl of SDS buffer (50 mM Tris-HCl, pH, 6.8, 10% glycerol, 2% SDS, 5% β-mercaptoethanol). For Western blotting, proteins were resolved in 10% SDS-PAGE, transferred to PVDF membrane (Millipore) and the membranes were incubated with appropriate antibodies (anti-HA, anti-GFP or anti-Actin antibody, Santa Cruz) in TBST buffer (10 mM Tris-HCl, pH, 7.5, 150 mM NaCl, 0.1% Tween 20) and proteins were detected by the enhanced chemiluminescence (ECL) system (Pierce).
Microscopy and Image Analysis
Yeast cells expressing Hxt1-GFP were stained with FM4-64 (lipophilic styryl dye for selectively staining vacuolar membrane, 1μg/ml) to visualize vacuole and analyzed with Olympus FluoView confocal microscope under 63X oil immersion objective lens using GFP, Texas Red filters. Images from confocal microscope were captured by FluoView software (Olympus) and un-manipulated raw images were used to quantify fluorescence intensities by ImageJ v1.4r software (NIH). For each cell in a given image, regions of interest on plasma or vacuolar membrane and in an area outside the cell (background) were traced and mean fluorescence intensities (both GFP and FM4-64) were measured. After background subtraction, the GFP signals in the plasma membranes were normalized to the FM4-64 signal of vacuolar membrane. At least 200 cells were analyzed and the data represented were the averages with error bars representing standard deviation (S.D).
2-NBDG Uptake Assay and Quantification
Cells were first grown in glucose free (5% glycerol + 2% ethanol) medium to mid log phase (O.D600nm = 1.2–1.5). The cells were harvested, resuspended in the aforementioned medium and incubated with 60μM of 2-NBDG at 30° C for 30 min. The uptake reaction was stopped by washing the cells three times with 1X Phosphate-buffered saline, pH 7.4. Live cells were visualized with Olympus FluoView confocal microscope under 63X oil immersion objective lens using GFP filter. The quantification of 2-NBDG fluorescence was calculated using ImageJ v1.4r software (NIH). Briefly, the mean fluorescence intensity (MFI) of individual cells within the field of view was calculated and normalized by subtracting the background fluorescence signal from a region without any cells. At least 200 cells were analyzed from three independent experiments. Relative 2-NBDG fluorescence (the fluorescence of 2-NBDG from hxtΔ was set to 1.0) was plotted with data showing averages ± S.D.
Conceived and designed the experiments: JHK. Performed the experiments: AR AD. Analyzed the data: AR AD KHC JHK. Contributed reagents/materials/analysis tools: AR JHK. Wrote the paper: JHK AR.
- 1. Hsu PP, Sabatini DM (2008) Cancer cell metabolism: Warburg and beyond. Cell 134: 703–707. pmid:18775299
- 2. Vander Heiden MG, Cantley LC, Thompson CB (2009) Understanding the Warburg effect: the metabolic requirements of cell proliferation. Science 324: 1029–1033. pmid:19460998
- 3. Warburg O (1956) On the origin of cancer cells. Science 123: 309–314. pmid:13298683
- 4. Mandelkern M, Raines J (2002) Positron emission tomography in cancer research and treatment. Technol Cancer Res T 1: 423–439. pmid:12625770
- 5. Crabtree HG (1929) Observations on the carbohydrate metabolism of tumours. Biochem J 23: 536–545. pmid:16744238
- 6. Lagunas R (1979) Energetic irrelevance of aerobiosis for S. cerevisiae growing on sugars. Mol Cell Biochem 27: 139–146. pmid:390364
- 7. Rolland F, Winderickx J, Thevelein JM (2002) Glucose-sensing and-signalling mechanisms in yeast. FEMS Yeast Res 2: 183–201. pmid:12702307
- 8. Ozcan S, Johnston M (1999) Function and regulation of yeast hexose transporters. Microbiol Mol Biol Rev 63: 554–569. pmid:10477308
- 9. Vinters HV, Beck DW, Bready JV, Maxwell K, Berliner JA, et al. (1985) Uptake of glucose analogues into cultured cerebral microvessel endothelium. J Neuropath Exp Neur 44: 445–458. pmid:3897465
- 10. Khoo S, Cobb MH (1997) Activation of mitogen-activating protein kinase by glucose is not required for insulin secretion. Pros Natl Acad Sci USA 94: 5599–5604. pmid:9159118
- 11. Rumsey SC, Daruwala R, Al-Hasani H, Zarnowski MJ, Simpson IA, et al. (2000) Dehydroascorbic acid transport by GLUT4 in Xenopus oocytes and isolated rat adipocytes. J Biol Chem 275: 28246–28253. pmid:10862609
- 12. Yoshioka K, Takahashi H, Homma T, Saito M, Oh KB, et al. (1996) A novel fluorescent derivative of glucose applicable to the assessment of glucose uptake activity of Escherichia coli. Biochim Biophys Acta 1289: 5–9. pmid:8605231
- 13. Lloyd PG, Hardin CD, Sturek M (1999) Examining glucose transport in single vascular smooth muscle cells with a fluorescent glucose analog. Physiol Res 48: 401–410. pmid:10783904
- 14. Yamada K, Nakata M, Horimoto N, Saito M, Matsuoka H, et al. (2000) Measurement of glucose uptake and intracellular calcium concentration in single, living pancreatic beta-cells. J Biol Chem 275: 22278–22283. pmid:10748091
- 15. O'Neil RG, Wu L, Mullani N (2005) Uptake of a fluorescent deoxyglucose analog (2-NBDG) in tumor cells. Molecular imaging and biology: MIB: the official publication of the Academy of Molecular Imaging 7: 388–392.
- 16. Boles E, Hollenberg CP (1997) The molecular genetics of hexose transport in yeasts. FEMS Microbiol Rev 21: 85–111. pmid:9299703
- 17. Reifenberger E, Boles E, Ciriacy M (1997) Kinetic characterization of individual hexose transporters of Saccharomyces cerevisiae and their relation to the triggering mechanisms of glucose repression. Eur J Biochem 245: 324–333. pmid:9151960
- 18. Kim JH, Polish J, Johnston M (2003) Specificity and regulation of DNA binding by the yeast glucose transporter gene repressor Rgt1. Mol Cell Biol 23: 5208–5216. pmid:12861007
- 19. Kim JH, Brachet V, Moriya H, Johnston M (2006) Integration of transcriptional and posttranslational regulation in a glucose signal transduction pathway in Saccharomyces cerevisiae. Eukaryot Cell 5: 167–173. pmid:16400179
- 20. Lakshmanan J, Mosley AL, Ozcan S (2003) Repression of transcription by Rgt1 in the absence of glucose requires Std1 and Mth1. Curr Genet 44: 19–25. pmid:14508605
- 21. Palomino A, Herrero P, Moreno F (2005) Rgt1, a glucose sensing transcription factor, is required for transcriptional repression of the HXK2 gene in Saccharomyces cerevisiae. Biochem J 388: 697–703. pmid:15705057
- 22. Roy A, Shin YJ, Cho KH, Kim JH (2013) Mth1 regulates the interaction between the Rgt1 repressor and the Ssn6-Tup1 corepressor complex by modulating PKA-dependent phosphorylation of Rgt1. Mol Biol Cell 24: 1493–1503. pmid:23468525
- 23. Flick KM, Spielewoy N, Kalashnikova TI, Guaderrama M, Zhu Q, et al. (2003) Grr1-dependent inactivation of Mth1 mediates glucose-induced dissociation of Rgt1 from HXT gene promoters. Mol Biol Cell 14: 3230–3241. pmid:12925759
- 24. Schmidt MC, McCartney RR, Zhang X, Tillman TS, Solimeo H, et al. (1999) Std1 and Mth1 proteins interact with the glucose sensors to control glucose-regulated gene expression in Saccharomyces cerevisiae. Mol Cell Biol 19: 4561–4571. pmid:10373505
- 25. Moriya H, Johnston M (2004) Glucose sensing and signaling in Saccharomyces cerevisiae through the Rgt2 glucose sensor and casein kinase I. Proc Natl Acad Sci U S A 101: 1572–1577. pmid:14755054
- 26. Roy A, Jouandot D 2nd, Cho KH, Kim JH (2014) Understanding the mechanism of glucose-induced relief of Rgt1-mediated repression in yeast. FEBS Open Bio 4: 105–111. pmid:24490134
- 27. Johnston M, Kim JH (2005) Glucose as a hormone: receptor-mediated glucose sensing in the yeast Saccharomyces cerevisiae. Biochem Soc Trans 33: 247–252. pmid:15667318
- 28. Kim JH, Roy A, Jouandot D 2nd, Cho KH (2013) The glucose signaling network in yeast. Biochim Biophys Acta 1830: 5204–5210. pmid:23911748
- 29. Horak J (2013) Regulations of sugar transporters: insights from yeast. Current genetics 59: 1–31. pmid:23455612
- 30. Roy A, Kim YB, Cho KH, Kim JH (2014) Glucose starvation-induced turnover of the yeast glucose transporter Hxt1. Biochim Biophys Acta 1840: 2878–2885. pmid:24821015
- 31. Yoshioka K, Oh KB, Saito M, Nemoto Y, Matsuoka H (1996) Evaluation of 2-[N-(7-nitrobenz-2-oxa-1,3-diazol-4-yl)amino]-2-deoxy-D-glucose, a new fluorescent derivative of glucose, for viability assessment of yeast Candida albicans. Appl Microbiol Biot 46: 400–404. pmid:8987729
- 32. Achilles J, Muller S, Bley T, Babel W (2004) Affinity of single S. cerevisiae cells to 2-NBDglucose under changing substrate concentrations. Cytometry 61: 88–98. pmid:15351993
- 33. Wieczorke R, Dlugai S, Krampe S, Boles E (2003) Characterisation of mammalian GLUT glucose transporters in a heterologous yeast expression system. Cell Physiol Biochem 13: 123–134. pmid:12876383
- 34. Hruz PW, Mueckler MM (2000) Cysteine-scanning mutagenesis of transmembrane segment 11 of the GLUT1 facilitative glucose transporter. Biochemistry 39: 9367–9372. pmid:10924131
- 35. Hruz PW, Mueckler MM (2001) Structural analysis of the GLUT1 facilitative glucose transporter (review). Mol Membr Biol 18: 183–193. pmid:11681785
- 36. Salas-Burgos A, Iserovich P, Zuniga F, Vera JC, Fischbarg J (2004) Predicting the three-dimensional structure of the human facilitative glucose transporter glut1 by a novel evolutionary homology strategy: insights on the molecular mechanism of substrate migration, and binding sites for glucose and inhibitory molecules. Biophys J 87: 2990–2999. pmid:15326030
- 37. Mueckler M, Makepeace C (2005) Cysteine-scanning mutagenesis and substituted cysteine accessibility analysis of transmembrane segment 4 of the Glut1 glucose transporter. J Biol Chem 280: 39562–39568. pmid:16172126
- 38. Heinze M, Monden I, Keller K (2004) Cysteine-scanning mutagenesis of transmembrane segment 1 of glucose transporter GLUT1: extracellular accessibility of helix positions. Biochemistry 43: 931–936. pmid:14744136
- 39. Sun L, Zeng X, Yan C, Sun X, Gong X, et al. (2012) Crystal structure of a bacterial homologue of glucose transporters GLUT1-4. Nature 490: 361–366. pmid:23075985
- 40. Iancu CV, Zamoon J, Woo SB, Aleshin A, Choe JY (2013) Crystal structure of a glucose/H+ symporter and its mechanism of action. Pros Natl Acad Sci USA 110: 17862–17867. pmid:24127585
- 41. Deng D, Xu C, Sun P, Wu J, Yan C, et al. (2014) Crystal structure of the human glucose transporter GLUT1. Nature 510: 121–125. pmid:24847886
- 42. Mueckler M, Makepeace C (2009) Model of the exofacial substrate-binding site and helical folding of the human Glut1 glucose transporter based on scanning mutagenesis. Biochemistry 48: 5934–5942. pmid:19449892
- 43. Lin CH, MacGurn JA, Chu T, Stefan CJ, Emr SD (2008) Arrestin-related ubiquitin-ligase adaptors regulate endocytosis and protein turnover at the cell surface. Cell 135: 714–725. pmid:18976803
- 44. Mueckler M, Weng W, Kruse M (1994) Glutamine 161 of Glut1 glucose transporter is critical for transport activity and exofacial ligand binding. J Biol Chem 269: 20533–20538. pmid:8051152
- 45. Hashiramoto M, Kadowaki T, Clark AE, Muraoka A, Momomura K, et al. (1992) Site-directed mutagenesis of GLUT1 in helix 7 residue 282 results in perturbation of exofacial ligand binding. J Biol Chem 267: 17502–17507. pmid:1517202
- 46. Garcia JC, Strube M, Leingang K, Keller K, Mueckler MM (1992) Amino acid substitutions at tryptophan 388 and tryptophan 412 of the HepG2 (Glut1) glucose transporter inhibit transport activity and targeting to the plasma membrane in Xenopus oocytes. J Biol Chem 267: 7770–7776. pmid:1560011
- 47. Uldry M, Thorens B (2004) The SLC2 family of facilitated hexose and polyol transporters. Pflug Arch Eur J Phy 447: 480–489.
- 48. Reifenberger E, Freidel K, Ciriacy M (1995) Identification of novel HXT genes in Saccharomyces cerevisiae reveals the impact of individual hexose transporters on glycolytic flux. Mol Microbiol 16: 157–167. pmid:7651133
- 49. Reifenberger E, Boles E, Ciriacy M (1997) Kinetic characterization of individual hexose transporters of Saccharomyces cerevisiae and their relation to the triggering mechanisms of glucose repression. Eur J Biochem 245: 324–333. pmid:9151960
- 50. Kruckeberg AL, Ye L, Berden JA, van Dam K (1999) Functional expression, quantification and cellular localization of the Hxt2 hexose transporter of Saccharomyces cerevisiae tagged with the green fluorescent protein. Biochem J 339 (Pt 2): 299–307. pmid:10191260
- 51. Krampe S, Stamm O, Hollenberg CP, Boles E (1998) Catabolite inactivation of the high-affinity hexose transporters Hxt6 and Hxt7 of Saccharomyces cerevisiae occurs in the vacuole after internalization by endocytosis. FEBS Lett 441: 343–347. pmid:9891967
- 52. Snowdon C, van der Merwe G (2012) Regulation of Hxt3 and Hxt7 turnover converges on the Vid30 complex and requires inactivation of the Ras/cAMP/PKA pathway in Saccharomyces cerevisiae. PloS one 7: e50458. pmid:23227176
- 53. Roy A, Kim JH (2014) Endocytosis and vacuolar degradation of the yeast cell surface glucose sensors Rgt2 and Snf3. J Biol Chem 289: 7247–7256. pmid:24451370
- 54. Hicke L (1999) Gettin' down with ubiquitin: turning off cell-surface receptors, transporters and channels. Trends Cell Biol 9: 107–112. pmid:10201076
- 55. Hicke L, Dunn R (2003) Regulation of membrane protein transport by ubiquitin and ubiquitin-binding proteins. Annu Rev Cell Dev Biol 19: 141–172. pmid:14570567
- 56. Galan JM, Moreau V, Andre B, Volland C, Haguenauer-Tsapis R (1996) Ubiquitination mediated by the Npi1p/Rsp5p ubiquitin-protein ligase is required for endocytosis of the yeast uracil permease. J Biol Chem 271: 10946–10952. pmid:8631913
- 57. Farwick A, Bruder S, Schadeweg V, Oreb M, Boles E (2014) Engineering of yeast hexose transporters to transport D-xylose without inhibition by D-glucose. Pros Natl Acad Sci USA 111: 5159–5164. pmid:24706835