Glucose transporters and transport kinetics in retinoic acid-differentiated T47D human breast cancer cells

Dalia Rivenzon-Segal1, Edna Rushkin1, Sylvie Polak-Charcon2, and Hadassa Degani1

1 Department of Biological Regulation, Weizmann Institute of Science, Rehovot 76100; and 2 Department of Pathology, Sheba Medical Center, Tel-Hashomer 52621, Israel


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The rates of glucose transport and of glycolysis and the expression of the glucose transporters GLUT-1 through GLUT-4 were measured in T47D human breast cancer cells that underwent differentiation by retinoic acid. Glucose transport was found to be the rate-limiting step of glycolysis in control and differentiated cells. The transporters GLUT-1, GLUT-3, and GLUT-4 were present in the cell membrane and in the cytoplasm, and GLUT-2 was present solely in the cytoplasm. Differentiation led to a reduction in GLUT-1 and to an increase in cytoplasmic GLUT-2 and GLUT-3 with no change in GLUT-4. Differentiation also caused a reduction in the maximal velocity of glucose transport by ~40% without affecting the Michaelis-Menten constant of glucose transport. These changes did not alter the steady-state concentration of the phosphate metabolites regulating cell energetics but increased the content of phospholipid breakdown phosphodiesters. In conclusion, differentiation of human breast cancer cells appears to be associated with decreased glycolysis by a mechanism that involves a reduction in GLUT-1 and a slowdown of glucose transport.

glycolysis; magnetic resonance spectroscopy; 2-deoxy-D-glucose; GLUT-1


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INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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TUMOR CELLS EXHIBIT HIGH GLYCOLYTIC ACTIVITY and accumulate lactate to high levels compared with normal tissues (2, 43). These processes were attributed to changes in glycolytic isoenzymes, such as hexokinase (14), or to an increased expression of glucose transporters (4, 16, 48, 50). Transport of glucose into cells was shown to be mediated by a family of facilitative diffusion glucose transporters (8, 9, 34). The members of this family (GLUT-1 through GLUT-5) comprise 12 transmembrane-spanning domains, with the amino and the carboxy termini exposed to the cytoplasmic face. The expression of the GLUT transporters was investigated in breast cancer cells (1, 52) and surgical specimens (3, 6, 49, 51). GLUT-1 was found to be expressed in the cell membrane, in the cytoplasm of all primary tumors, and in the lymph node metastases. Immunostaining of GLUT-1 in normal mammary epithelium was either negative or weaker than that observed in breast cancer cells of the same patient. GLUT-2 was expressed mainly in the cytoplasm of these tumors and with the same intensity as in normal tissue. GLUT-3 was not expressed in either tumor or healthy tissues. These observations support a dominant role for GLUT-1 in determining glucose transport in breast cancer.

In this study, we investigated the effects of differentiation on the kinetics of glucose transport and metabolism and on the expression of the various glucose transporters in T47D human breast cancer cells. Previous studies have attempted to generalize the differences in glucose metabolism between malignant and normal cells, whereas we present data from cells at their nondifferentiated and differentiated states.

Differentiation was induced by retinoic acid (RA), which belongs to the retinoid family of compounds. These compounds, comprising both natural and synthetic vitamin A derivatives, were shown to be capable of influencing many biological functions, including proliferation, cell cycle arrest, differentiation, apoptosis, and fetal development (19, 27, 28, 35, 42). Retinoid action was shown to be mediated by two classes of nuclear receptors: the retinoic acid receptor (RAR) and the retinoid X receptor (RXR) (30). Both the RARs and the RXRs modulate the expression of their target genes by interacting as either homodimers or heterodimers with RA response elements (30).

The kinetics of glucose transport and phosphorylation, the initial steps of glucose metabolism, were previously investigated with the aid of 2-deoxy-D-glucose (DG) (37). This substrate is transported and phosphorylated with Michaelis-Menten constants similar to those of glucose (5, 9, 34, 47, 52). The phosphorylated form of DG, 2-deoxy-D-glucose 6-phosphate (DGP), is usually not metabolized further and is therefore trapped within the cells (47), although phosphatase activity may yield back DG (20). Most of the studies performed with DG have used a tracer approach with radioactive DG (5, 22, 37, 45). DG transport and phosphorylation were also followed by monitoring DGP accumulation with 31P magnetic resonance spectroscopy (MRS) (11, 24). In addition, the kinetics of glucose consumption and glycolysis can be measured noninvasively by monitoring the metabolism of [1-13C]glucose and the synthesis of [3-13C]lactate with 13C MRS, as was demonstrated previously in studies of breast cancer cells (17, 29, 32).

In this work, the kinetic results derived from the DG and the [1-13C]glucose studies have shown that glycolysis is rate limited by the transport of glucose in control and RA-differentiated cells. The glucose transporters GLUT-1, GLUT-3, and GLUT-4 were found to be present in the cell membrane and in the cytoplasm, whereas GLUT-2 was localized solely in the cytoplasm in both control and differentiated cells. RA-induced differentiation led to a reduction in glycolysis that was associated with a decrease in GLUT-1 expression and a concomitant slowdown of the rate of glucose transport.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
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Cell Culture

T47D-clone 11 human breast cancer cells, kindly provided by Prof. I. Keydar (1985, Tel-Aviv University, Israel), were cultivated in DMEM supplemented with 10% FCS, insulin (0.2 IU/ml), penicillin (200 µg/ml), streptomycin (200 µg/ml), and neomycin (10 µg/ml) (combined antibiotics, BIO-LAB, Jerusalem, Israel). Stock solution of RA (Sigma Chemical, St. Louis, MO) was prepared under subdued light in DMSO at a concentration of 10-2 M and was stored at -70°C in the dark. Cells were treated a day after seeding with RA (1 µM or 10 µM) dissolved in DMSO, and control cultures received the equivalent amount of DMSO. Duplicate cultures were counted after detachment by trypsinization with a cytometer. Percent inhibition of growth was calculated from [(N- Nt)/Nc] × 100, where Nc and Nt represent the number of cells in control and RA-treated samples, respectively.

Flow Cytometry Measurement of Keratin Expression

Cells were washed with PBS, detached with EDTA, and centrifuged at 1,000 rpm for 4 min. The supernatant was removed, and the pellet was washed twice with PBS. Then, 1 × 106 cells were incubated for 40 min at 4°C in 90 µl solution of PBS with 0.05% saponin containing 10 µl of mouse FITC-conjugated monoclonal anti-cytokeratins (K8 and K18) antibody (Novo Castra Laboratory, Newcastle, UK) or with FITC-conjugated mouse IgG1 (PharMingen, San Diego, CA) as negative control. Finally, the cells were washed in PBS, resuspended in 0.6 ml PBS, and transferred to 12 × 75-mm FALCON 2052 FACS tubes (Becton-Dickinson, San Jose, CA). Data from 104 cells were collected on a FACS SCAN flow cytometer (Becton-Dickinson) with an argon-ion laser tuned to 488 nm, and the mean fluorescence was obtained from the recorded data. The measurements of control and RA-treated cells were performed in duplicate, with the samples grown, processed, and analyzed in parallel.

Transmission Electron Microscopy Studies

Cells were fixed in 2.5% glutaraldehyde in 0.1 M cacodylate buffer (pH = 7.4) for 30 min at room temperature. Fixed cells were removed with a rubber policeman and postfixed in 1% OsO4 in cacodylate buffer for 2 h. After dehydration in graded ethanol solutions, the cells were embedded in Epon 812. Ultra-thin sections were contrasted with uranyl acetate and lead citrate. Thin sections were examined with a Jeol JEM 1200 EX II.

Cell Cycle Analysis

Cells were trypsinized and suspended in PBS (1 × 106 cells/ml). Each sample was vortexed with 10 µl of 10% Triton and 5 µl of 15 µM propidium iodide (PI; Sigma Chemical). Analysis was performed immediately after vortexing. Samples were analyzed using a Becton-Dickinson FACS SCAN equipped with an Argon laser (488 nm) and emission filter transmitting at 588 nm (FL2). Gating was applied according to the parameters FL2-area vs. FL2-width to prevent entrance of clumps into the analysis. From each sample, 10,000 cells were collected, excluding the gated cells. Fluorescence of individual cells stained with PI was measured. The percentage of cells in G0/G1, S, and G2/M phases was estimated from the FL2-height histogram and by applying the MOD-FIT software.

Flow Cytometry Measurement of Total GLUT Expression

A procedure described by Ojcius et al. (33) was adapted for T47D cells. Cells were washed with PBS, detached with EDTA-trypsin, resuspended in 2 ml growth medium, and centrifuged at 1,000 rpm for 4 min. The supernatant was removed, and the pellet was washed twice with PBS, fixed in 1 ml of 1% paraformaldehyde, and neutralized in 50 mM NH4Cl. After two additional washes with PBS, 1 × 106 cells were incubated for 45 min at 4°C in 50 µl solution of PBS with 0.05% saponin containing the four rabbit polyclonal anti-GLUT antibodies [Chemicon International, Temecula, CA; 1:50 dilution for GLUT-1 (MYM) and GLUT-2, 1:25 dilution for GLUT-3 (MYG) and GLUT-4] or with 50 µl rabbit IgG (Chemicon International) as negative control. The cells were then washed and incubated for another 30 min at 4°C with 50 µl PE-conjugated donkey anti-rabbit antibody (10 mg/ml; Chemicon International) in PBS with 0.05% saponin. Finally, the cells were washed in PBS, resuspended in 0.6 ml PBS, and transferred to 12 × 75-mm FALCON 2052 FACS tubes (Becton-Dickinson). Data from 104 cells were collected on a FACS SCAN flow cytometer (Becton-Dickinson) with an argon-ion laser tuned to 488 nm, and the mean fluorescence was obtained from the recorded data. The measurements of control and RA-treated cells were performed in duplicate, with the samples grown, processed, and analyzed in parallel.

Flow Cytometry Measurement of Membranal GLUT Expression

Cells were washed with PBS, detached with EDTA-trypsin, resuspended in 2 ml growth medium with soybean trypsin inhibitor (0.2 mg/ml), and centrifuged at 1,000 rpm for 4 min. The supernatant was removed, and the pellet was washed twice with PBS and fixed in 2 ml of 10% formaldehyde. After two additional washes with PBS, 1 × 106 cells were incubated for 45 min at 4°C in 50 µl solution of PBS containing the rabbit polyclonal FITC-anti-GLUT antibodies [Chemicon International; 1:10 dilution for GLUT-1 (FITC-MYH) and GLUT-3 (FITC-MYK)] or with FITC-conjugated rabbit IgG (Jackson Laboratories, West Grove, PA) as negative control. The cells were then washed twice in PBS, resuspended in 0.6 ml PBS, and transferred to 12 × 75-mm FALCON 2052 FACS tubes (Becton-Dickinson). Data from 104 cells were collected on a FACS SCAN flow cytometer (Becton-Dickinson) with an argon-ion laser tuned to 488 nm, and the mean fluorescence was obtained from the recorded data. The measurements of control and RA-treated cells were performed in duplicate, with the samples grown, processed, and analyzed in parallel.

Immunofluorescence Studies of GLUT Localization

Cells grown on coverslips were fixed and permeabilized for 2 min with 0.5% Triton in 3% paraformaldehyde. Cells were further fixed in 3% paraformaldehyde for 20 min, washed with PBS, and incubated for 45 min at room temperature with the various GLUT antibodies, as described above for the flow cytometry analysis of total GLUT expression. Finally, cells were washed three times with PBS and incubated for an additional 30 min with TRITC-labeled goat antibody to rabbit IgGs (Jackson Laboratories). Stained coverslips were mounted in Evanol and examined with a Zeiss microscope.

Cell Culture on Beads

T47D cells were seeded after trypsinization on agarose-polyacrolein beads coated with collagen from calf skin (Type 1, Sigma) at a density of 6 × 106 cells/ml beads. On the following day, the cells were treated with 1 µM RA in DMSO or with DMSO alone (control). The medium with or without RA was replaced every 48 h and then every 24 h before each NMR experiment. Both control and RA-treated cells were grown on beads for 6-7 days. Before the NMR measurements, 2 ml of beads with cells were transferred into a sterile 10-mm NMR tube, and 1 ml of beads with cells remained for determining the number of cells. The tube with the cells was placed in the NMR spectrometer and perfused constantly with 50-200 ml of DMEM-10% FCS saturated with 95% O2-5% CO2 under sterile conditions at 36 ± 1°C (10).

In 13C studies of glucose metabolism, [1-13C]glucose, 99% enriched (Cambridge Isotope Laboratories, Andover, MA) was added to the perfusion medium of control and RA-treated cells at a concentration of 5.6 mM. In the experiments with DG (Sigma Chemical), [1-13C]glucose concentration in the perfusion medium was 8.3 mM, and DG concentration was 0.83 mM. In the kinetic studies of glucose metabolism, increasing concentrations of [1-13C]glucose, in the range of 0.5-13 mM, were added sequentially to the perfusion medium.

NMR Measurements

NMR spectra were recorded with a vertical Bruker AM-500 spectrometer equipped with either a software-controlled quadro nuclear probe or a broad band probe. Proton-decoupled 31P NMR spectra (30-min acquisition) were recorded at 202.5 MHz by applying 45° pulses, 1 s repetition time, and a continuous composite pulse proton decoupling of ~1 W. Concentrations were calculated in reference to the integrated intensity of medium Pi, measured independently (ranging between 1 and 2 mM), with account taken for relaxation effects by referring to results obtained from fully relaxed spectra. 31P signals were assigned in reference to alpha -nucleoside 5'-triphosphate (alpha -NTP) at -10.03 ppm. 13C spectra were recorded at 125.7 MHz by applying 60° pulses, 2 s repetition time, and a continuous composite pulse proton decoupling of ~1 W. The total acquisition time of each spectrum ranged between 10 and 30 min. Spectra were analyzed using XWINNMR (a standard Bruker software package). Studies of glucose transport with DG were performed by alternate recordings of 13C and 31P NMR spectra with the parameters described above. After recording 13C spectra of cells perfused with standard medium containing 8.3 mM [1-13C]glucose, the medium was replaced with fresh medium containing 8.3 mM [1-13C]glucose and 0.83 mM DG, and 31P spectra were recorded, followed by recording again of 13C spectra. Finally, the DG-containing medium was replaced with a standard, DG-free growth medium containing 8.3 mM nonlabeled glucose, and 31P spectra were recorded for several hours.

Data Analysis

Glucose metabolism. The signal of beta -[1-13C]glucose served as a reference for chemical shift assignment at 96.8 ppm. The combined areas of the signals of alpha - and beta -[1-13C]glucose at the beginning of the experiments were proportional to the initial concentration of glucose (5.6 or 8.3 mM) and served as a reference for determining the concentration of the 13C-labeled metabolites, with account taken for relaxation and nuclear Overhauser effects.

The rates of glucose consumption and lactate production were calculated by using a linear fit to the changes in signal area of glucose and lactate, respectively, as previously described (32). Rates were calculated in units of amount (fmol) per cell per time (h). Because the duplication time of these cells was relatively long, 34 h for control cells and 50 h for RA-treated cells, the change in the number of cells in the course of the 13C NMR experiments (~14 h) was small, of the order of 10% or less, and was therefore neglected.

DG transport and phosphorylation. The concentrations of DGP were calculated from 31P spectra in reference to the medium Pi, taking into account saturation effects. Analysis of the time course of DGP accumulation was based on a model developed for brain studies (37) and modified to our system.

In general, glucose transport follows a Michaelis-Menten mechanism. The rate of DG transport in the presence of glucose, (Vt), can be described by the Michaelis-Menten equation by taking into account substrate competition (12)
V<SUB>t</SUB><IT>=</IT><FR><NU><IT>V<SUP>*</SUP></IT><SUB>max</SUB>[DG]<SUB>out</SUB></NU><DE><IT>K<SUP>*</SUP></IT><SUB>m</SUB>(<IT>1+</IT>[Glc]<SUB>out</SUB><IT>/K</IT><SUB>m</SUB>)<IT>+</IT>[DG]<SUB>out</SUB></DE></FR> (1)
V*max and K*m are the Michaelis-Menten constants for DG transport, and Km is the Michaelis-Menten constant for glucose (Glc) transport. When [Glc] > [DG], and K*m approx  Km, the rate becomes proportional to DG concentration according to
V<SUB>t</SUB><IT>=</IT><FR><NU><IT>V<SUP>*</SUP></IT><SUB>max</SUB></NU><DE><IT>K<SUP>*</SUP></IT><SUB>m</SUB>(<IT>1+</IT>[Glc]<SUB>out</SUB><IT>/K</IT><SUB>m</SUB>)</DE></FR> [DG]<SUB>out</SUB> (2)
Thus, when glucose concentration in the medium is approximately constant, the transport can be described as a first-order reaction in [DG]out
<FR><NU>d[DG]<SUB>out</SUB></NU><DE>d<IT>t</IT></DE></FR><IT>=</IT>−<IT>k</IT><SUB>t</SUB>[DG]<SUB>out</SUB> (3)
where kt, the apparent first-order transport rate constant, includes the Michaelis-Menten parameters as shown in Eq. 2.

Under our experimental conditions, Glc concentration was 10-fold higher than DG concentration. Previous studies have shown that the Km values of DG transport and of Glc transport via GLUT-1 through GLUT-4 are similar (5, 9, 34, 52). Thus it follows that the ratio of DG transport rate to Glc transport rate is approximately equal to the concentration ratio of these substrates in the medium.

The rate of DG phosphorylation to DGP can also be described as a first-order reaction (37)
<FR><NU>d[DGP]</NU><DE>d<IT>t</IT></DE></FR><IT>=k</IT><SUB>p</SUB>[DG]<SUB>in</SUB> (4)
where, by assuming negligible back transport of DG from the cells to the medium, we get
<FR><NU>d[DG]<SUB>in</SUB></NU><DE>d<IT>t</IT></DE></FR><IT>=k</IT><SUB>t</SUB>[DG]<SUB>out</SUB><IT>−k</IT><SUB>p</SUB>[DG]<SUB>in</SUB> (5)
Solving Eqs. 4 and 5 yielded the following results
[DG]<SUB>in</SUB>(<IT>T</IT>)<IT>=k</IT><SUB>t</SUB> exp(−<IT>k</IT><SUB>p</SUB><IT>T</IT>) <LIM><OP>∫</OP><LL><IT>0</IT></LL><UL><IT>T</IT></UL></LIM> [DG]<SUB>out</SUB> exp(<IT>k</IT><SUB>p</SUB><IT>·t</IT>)d<IT>t</IT> (6)

[DGP](<IT>&tgr;</IT>) (7)

<IT>=k</IT><SUB>t</SUB><IT>k</IT><SUB>p</SUB> <LIM><OP>∫</OP><LL><IT>0</IT></LL><UL><IT>&tgr;</IT></UL></LIM> <FENCE>exp(−<IT>k</IT><SUB>p</SUB><IT>T</IT>) <LIM><OP>∫</OP><LL><IT>0</IT></LL><UL><IT>T</IT></UL></LIM> [DG]<SUB>out</SUB> exp(<IT>k</IT><SUB>p</SUB><IT>·t</IT>)d<IT>t</IT></FENCE>d<IT>T</IT>
The experimental data of DGP accumulation were fitted to Eq. 7 by using a Levenberg-Marquardt algorithm for nonlinear best fitting, yielding kt and kp. The apparent initial rate of DGP synthesis was calculated from the initial tangent of the fitted curve.

Kinetic studies. The kinetic analysis was based on measurements of the apparent initial rate of glucose consumption as a function of glucose concentrations in the medium, by using the zero trans method for transport measurements (39). The trans face is the internal face of the cell membrane. This method is the most suitable approach for transport measurements of substrates that are rapidly metabolized. The method is based on plots of the apparent initial rate (v), of the rate constant [K = (v/S)], and of the inverse of the rate constant (1/K) of glucose consumption vs. glucose concentrations. A distinct pattern in each of the three plots defines the transport mechanism, namely, simple diffusion, Michaelis-Menten, Michaelis-Menten + diffusion, competition, and so forth. The determination of a transport mechanism is based on the best fit of the experimental data to all three plots, yielding one set of kinetic parameters (39).


    RESULTS
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MATERIALS AND METHODS
RESULTS
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Cell Differentiation

RA inhibited cell growth but did not exert a cytotoxic effect on T47D cells. In the first 3-4 days of treatment with 1 µM or 10 µM RA, no significant growth inhibition was observed, but after 6 days, the percent inhibition by RA reached 40 ± 10%. Longer exposure to RA of up to 11 days resulted in a 60% inhibition of growth. This inhibition led to an increase in the doubling time from 34 h in control cells to 50 h in RA-treated cells. Although growth inhibition was not substantial, it was possible to observe by light microscopy distinct morphological changes induced by RA. These changes included a more rectangular and elongated shape of the cells and a preferential growth of the monolayers in a certain direction.

To further characterize the RA-induced growth inhibition, we measured the cell cycle distribution. The proportion of cells in each phase of the cell cycle is summarized in Table 1. As can be seen, a significant increase in the proportion of cells in the G0/G1 phase was found in RA-treated cells (6 days), with a concomitant small but significant decrease in the S phase and in the G2/M phase.

                              
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Table 1.   Distribution in cell cycle phases of control and RA-treated cells

To characterize the RA effect on cell differentiation, we monitored the presence of keratins with flow cytometry and transmission electron microscopy (TEM). Changes in the keratin expression pattern are considered to be markers of the differentiation status (23). Flow cytometry analyses showed a 70% increase (n = 3, P < 0.003, paired two-tail t-test) in the expression of K8 and K18 cytoplasmatic keratins in cells treated with RA for 5-7 days (Fig. 1A). Inspection of TEM micrographs clearly showed that, in cells treated with RA (1 µM, 6 days), the cytoskeletal organization was characterized by increased levels of keratins (Fig. 1, C, E, and F), whereas in control cells, keratins were hardly visible (Fig. 1, B and D). Furthermore, these micrographs demonstrated augmentation of the presence of desmosomes connecting the keratins of adjacent cells (Fig. 1, C and E).


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Fig. 1.   Induction of differentiation by retinoic acid (RA) in T47D cells. A: flow cytometry histogram of fluorescence intensity expressed in arbitrary units (a.u.) of FITC-conjugated keratins (K8 and K18) in control T47D cells (shaded histogram) and in RA-treated cells, 1 µM, 6 days (open histogram). Nonspecific control antibody in control and RA-treated cells is shown at the low fluorescence intensity (bold lines). B-F: transmission electron micrographs of T47D human breast cancer cells; B and D: control cells; C, E, and F: cells treated with 1 µM RA for 7 days. Arrows, appearance of cytoplasmatic keratins; arrowheads, desmosomes. D and E-F: ×2.5 magnification of areas shown in B and C, respectively. E: keratins attached to a desmosome. F: cytoplasmic keratins.

Phosphate Metabolites

Theprofile and content of the phosphate metabolites in control and RA-differentiated cells were obtained from 31P spectra (Fig. 2B). The signals of the phosphates in these spectra included a phosphocholine (PC) signal at high intensity and other phosphomonoesters (not assigned), intra- and extracellular Pi, signals of glycerophosphoethanolamine (GPE), glycerophosphocholine (GPC), phosphocreatine (PCr), and alpha -, beta -, and gamma -NTP. The steady-state content of the energy-rich metabolites NTP and PCr was not affected by RA treatment (Fig. 2A). The high level of PC, a precursor of phosphatidylcholine, also did not change significantly. Among the other phosphate metabolites, we found that the normally low levels of GPC and GPE, which are metabolic products in the phospholipid breakdown pathway, were significantly elevated in the RA-treated cells (Fig. 2A): GPC level increased from 2.6 to 8.7 fmol/cell (P < 0.0012), and GPE level increased from 1.45 to 8.7 fmol/cell (P < 0.0002).


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Fig. 2.   Phosphate metabolites in control and RA-treated T47D cells perfused at 36 ± 1°C. A: solid columns, control; open columns, RA treated. Results are means ± SE (n = 5), with account taken of saturation effects. * P < 0.0012; ** P < 0.0002. B: 31P spectrum of control T47D cells. Spectrum was recorded as described in MATERIALS AND METHODS and processed using a line broadening of 20 Hz. PC, phosphocholine; Pi, inorganic phosphate; GPE, glycerophosphoethanolamine; GPC, glycerophosphocholine; PCr, phosphocreatine; NTP, nucleoside triphosphate; UDPS, uridine diphosphate sugars.

Metabolism of Glucose

Metabolic modulations associated with differentiation induced by RA were investigated by monitoring glucose metabolism in control and in RA-treated cells with 13C MRS. A typical 13C spectrum of control cells is shown in Fig. 3A. The main signals are due to [1-13C]glucose and [3-13C]lactate in the perfusion medium. A small fraction of 13C-labeled glucose was incorporated to glycerol 3-phosphate and via the tricarboxylic acid cycle into glutamate and glutamine C-4 and also to C-2 and C-3 (Fig. 3B), as previously described (32).


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Fig. 3.   13C studies of glucose metabolism. A: 13C spectrum of control T47D cells perfused for 8 h with medium containing 5.6 mM [1-13C]glucose, at 36 ± 1°C. B: ×32 magnification of A in 25-65 ppm region showing signals due to glutamate, glutamine, and glycerol 3-phosphate (G-3-P). Spectrum was accumulated as described in MATERIALS AND METHODS and processed using a line broadening of 15 Hz. C: time courses of [1-13C]glucose consumption () and of [3-13C]lactate production () in this experiment.

The linear decrease in the 13C signals of the beta - and alpha -anomers of C-1 glucose and the concomitant increase in 13C signal of C-3 lactate were monitored with time (Fig. 3C). The rate of [1-13C]glucose consumption in control cells was 474 ± 60 fmol · cell-1 · h-1 (n = 4) and decreased significantly (P < 0.04 paired two-tailed Student t-test) by ~40% in RA-treated cells to 269 ± 35 fmol · cell-1 · h-1 (n = 4). Similarly, the rate of lactate production in RA-treated cells decreased significantly (P < 0.04 paired two-tailed Student t-test) by ~40% from 321 ± 40 fmol · cell-1 · h-1 (n = 4) in control cells to 195 ± 25 fmol · cell-1 · h-1 (n = 4) in RA-treated cells. No significant changes were observed in the intensities of the small signals of glutamate, glutamine, and glycerol 3-phosphate as a result of RA treatment.

DG Transport and Phosphorylation

The role of glucose transport and phosphorylation in determining glucose metabolism was investigated using DG, an analog of glucose, added at low concentration to the perfusion medium. Synthesis of DGP and the content of the other phosphate metabolites were monitored in living cells by 31P NMR (Fig. 4). The increase with time of DGP concentration (in fmol/cell) was fitted to Eq. 7, with the apparent rate constant of DG transport (kt) and the apparent rate constant of DG phosphorylation (kp) as free parameters (Fig. 5, A and B). This analysis yielded a significantly lower (P < 0.015 paired two-tailed Student t-test) apparent rate constant of DG transport, with an average (±SE) of 0.6 ± 0.2 h-1 (n = 5) relative to the apparent rate constant of DG phosphorylation of 4.6 ± 0.9 h-1 (n = 5) in control and RA-treated cells. These results indicated that in the two differentiated states, transport of DG limited the rate of DGP synthesis. Further support of this conclusion was obtained by comparing the apparent initial rate of DG transport to the rate of glucose consumption. The ratio between these two rates of ~0.1 (n = 4) was similar to the concentration ratio of DG to glucose, as expected, when the rate of glucose consumption is equal to the rate of glucose transport.


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Fig. 4.   31P spectra of cells perfused at 36 ± 1°C for 5 h with medium containing 0.83 mM 2-deoxy-D-glucose (DG) and 8.3 mM [1-13C]glucose. A: control cells; B: RA-treated cells. Spectra were recorded as in Fig. 2. DGP and other metabolites were assigned as in Fig. 2.



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Fig. 5.   Changes in content of DGP and other phosphate metabolites. A-D: after addition of 0.83 mM DG; E and F: after DG removal. In between, 13C spectra were recorded for 7 h. A and B: accumulation of DGP (open circle ) in control and in RA-treated cells, respectively. Curves were obtained by a nonlinear least square fitting to Eq. 7. C,E and D,F: changes in PC (), gamma -NTP (black-triangle), intracellular Pi (), and DGP (open circle ) in control and RA-treated cells, respectively.

The 31P spectra recorded during DGP accumulation also demonstrated a concomitant decrease in NTP and an increase in intracellular Pi, with no change in PC level (Fig. 5, C and D). In addition, no significant changes were observed in the low levels of PCr, GPC, and GPE (data not shown). In control cells, substantial DGP accumulation (~45 fmol/cell) resulted in a decrease of NTP content from 30 to 22 fmol/cell (27%). In parallel, intracellular Pi reached a higher steady-state level, increasing from 22 to 42 fmol/cell (~100%). In RA-treated cells, DGP reached a lower accumulation level of ~24 fmol/cell. The changes in NTP and intracellular Pi levels were lower as well: an ~10% decrease in NTP level and a ~50% increase in intracellular Pi. The decrease in NTP did not result in a concomitant increase in NDP, because both gamma -NTP, which includes also beta -NDP, and beta -NTP decreased by the same extent. Therefore, changes in the ratio of NTP to Pi reflected changes in the phosphorylation potential of the cells. The phosphorylation potential decreased by 2.7-fold and 1.6-fold in control and RA-treated cells, respectively, in agreement with the difference in DGP accumulation at steady state. We have also found that the rate of glucose consumption decreased by two- to threefold in the presence of this analog. The changes in glucose consumption rate correlated with the changes in the phosphorylation potential. Thus it appears that, in the presence of DG, a less favorable but stable energetic steady state has been reached, with a slower rate of glucose metabolism.

After removal of DG from the perfusion medium, DGP slowly decayed exponentially in both control and RA-treated cells, with an apparent rate constant of ~0.1-0.2 h-1, whereas intracellular Pi and NTP levels remained unchanged (Fig. 5, E and F). This indicates an irreversible effect of DG on the energy status of the cells.

Kinetic Studies

Because the initial rate of glucose consumption in the DG experiment was found to be limited by the transport, we determined the kinetics of glucose transport by measuring glucose consumption as a function of glucose concentration. The best fit analysis of the three kinetic plots indicated a Michaelis-Menten mechanism (Fig. 6, left): v = Vmax*[GL]/(Km+[GL]), with a set of best fitted Km and Vmax values (Table 2). In RA-treated cells, the Km of transport, derived from glucose consumption rates, was almost the same as in control cells, but the Vmax of transport was reduced by ~40%, from 753 to 469 fmol · cell-1 · h-1.


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Fig. 6.   Variations in the initial rate (v), rate constant (K = v/S), and 1/K of glucose consumption (left) and lactate production (right) with glucose concentration (S) in RA-treated cells. Results were obtained by analyzing 13C spectra for determining rates of glucose consumption and lactate production after the sequential addition of increasing [1-13C]glucose concentrations. Curves were obtained by fitting data to a Michaelis-Menten equation and were analyzed according to the zero trans method.


                              
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Table 2.   Apparent Km and Vmax of glucose transport and labeled lactate production in control and RA-treated T47D cells

The rates of [3-13C]lactate synthesis at varying concentrations of glucose were analyzed in analogy with the analysis of the glucose data. Although the production of lactate through glycolysis includes many steps, we treated the data by assuming one apparent rate-limiting step. On the basis of this assumption, we have determined apparent Km and Vmax values for lactate synthesis in both control and RA-differentiated cells (Fig. 6, right). The results are summarized in Table 2. The apparent Km of lactate synthesis was the same in control and RA-differentiated cells. Moreover, the apparent Km of lactate synthesis was similar to the Km of glucose transport (2.6 mM). This similarity suggests that glucose transport is also the rate-limiting step of lactate production, namely, glycolysis. The apparent Vmax of lactate production decreased in RA-treated cells by ~35% in accord with the decrease in Vmax of glucose transport. Thus the general conclusion of this study is that, in T47D cells, glycolysis is rate limited by glucose transport, and upon RA differentiation there is ~35-40% reduction in the rate of glycolysis.

Expression of the GLUT Transporters

The rate of glycolysis was found to be limited by the transport of glucose, which in turn is regulated by the presence of glucose transporters in the cell membrane. Regulation of the facilitative diffusion glucose transporters by differentiation was thus investigated by measuring changes in the expression of these transporters. Flow cytometry with anti-glucose transporter antibodies was used in these studies. Further visual assessment of the localization of GLUTs was followed by immunofluorescence studies.

We have used antibodies against the cytoplasmic COOH termini of the transporter, termed cytoplasmic antibody (GLUT-1-MYM, GLUT-3-MYG, and GLUT-4), and against the exofacial loop of the transporter, termed membranal antibody (GLUT-1-MYH, GLUT-2, and GLUT-3-MYK). The MYM and MYG antibodies were characterized and employed previously (33, 49, 50, 51). Experiments in nonpermeabilized and permeabilized cells enabled us to characterize the expression of GLUT-1, GLUT-2, and GLUT-3 in the plasma membrane and in the cytoplasm.

RA treatment resulted in a highly significant (P < 0.0001 paired two-tailed Student's t-test) decrease of 22 ± 4% (n = 7) in the total GLUT-1 expression measured with the MYM antibody. Immunofluorescence with this antibody showed that GLUT-1 was predominantly localized in the cell membrane (Fig. 7, A and B) with the amount of GLUT-1 in the cell membrane of control cells (Fig. 7A) higher than that of treated cells (Fig. 7B). Because nonspecific nuclear staining was observed in the immunofluorescence studies, we further employed the MYH antibody to the GLUT-1 exofacial loop. This antibody demonstrated a 33% (n = 2) decrease in membranal GLUT-1 after RA treatment, confirming the reduction in GLUT-1 expression. GLUT-2 expression increased by 34 ± 6% (n = 2) in permeabilized cells treated with RA. In nonpermeabilized cells, it was not possible to detect any expression of GLUT-2. Thus GLUT-2 appeared to be localized to the cytoplasm and was not present in detectable levels in the plasma membrane. This was further confirmed by immunofluorescence studies (data not shown). The total GLUT-3 expression increased by 25 ± 9% (n = 4) (P < 0.030 paired two-tailed Student's t-test) with RA treatment; however, the membranal expression of GLUT-3 was not affected by RA (n = 2). Immunofluorescence studies have shown that GLUT-3 was distributed in both the plasma membrane and the cytoplasm (Fig. 7, C and D). The increase in GLUT-3 fluorescence in RA-treated cells appeared to be due solely to an increase in cytoplasmic GLUT-3 (Fig. 7D), in accord with the flow cytometry studies. The total GLUT-4 expression was not affected, within experimental error, by the treatment with RA and remained the same as in control cells. GLUT-4 was also found to be localized to both the plasma membrane and the cytoplasm. The antibody staining of the nuclei was regarded as nonspecific on the basis of their previous characterization (50, 51).


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Fig. 7.   Immunofluorescence of GLUT-1 (A, B) and GLUT-3 (C, D) in control cells (A, C) and in cells treated with RA (1 µM) for 7 days (B, D).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Glucose metabolism through glycolysis is known to be enhanced in cancer cells relative to their counter, normal differentiated cells (43). High glycolytic rates were also demonstrated in cultured human breast cancer cells (29, 32). Although the malignant transformation of cancer cells cannot be reversed, it is possible to induce processes that result in a higher differentiation state (38). This study was designed to show that differentiation slows down glycolysis and to characterize the molecular and kinetic changes responsible for this effect. Differentiation was induced by treating T47D human breast cancer cells with RA. RA and other retinoids were shown to inhibit the growth of T47D cells and to cause a cell cycle arrest at G1 phase with a decrease in cell number in the S phase (44, 46). RA was also shown to induce cellular differentiation in T47D cells, as demonstrated by increased levels of two keratins, K8 and K18, which serve as markers for differentiation (23). We have confirmed the RA growth inhibition and cell cycle arrest in the G0/G1 phase and the decrease in the S phase. In addition, we have confirmed the occurrence of morphological changes as well as elevation in the expression of keratins and desmosomes, all indicative of induced differentiation. Several reports have shown that retinoids can also induce apoptosis in breast cancer cells (27, 35, 42). However, neither the TEM nor the flow cytometry studies have shown RA-induced apoptosis in T47D cells. Whether cells will undergo RA-induced apoptosis or differentiation appears to be a cell type-specific process, as was previously demonstrated (42).

The rates of glycolysis in control and differentiated T47D cells were measured using [1-13C]glucose and 13C MRS. The Warburg hypothesis associated impaired respiration and high rates of aerobic glycolysis with malignant transformation (43). However, aerobic glycolysis can occur also in normal highly proliferating cells (13). Indeed, the rates of glucose consumption and of lactate production were found to be similar in T47D cells and in proliferating human mammary epithelial cells (41). Here, however, we have found a substantial difference in the rate of aerobic glycolysis in the same cells but at two distinct differentiation states. In general, changes in glycolysis were attributed to variations in the expression and activity of the isoenzymes participating in the glycolytic pathway (14) and in the expression of the glucose transporters (4, 6, 7, 16, 48, 50). We investigated the latter alternative by characterizing the kinetics of glucose transport and metabolism and determining the expression of the glucose transporters.

Glucose transport was measured by using the tracer DG method (37), but instead of the usual radioactive procedure, we monitored DGP synthesis directly by 31P NMR. Although we have used a higher DG concentration than in the classical radioactive measurements, a tracer kinetic approach could be applied. The model-based kinetic analysis of DG transport and phosphorylation to DGP yielded an approximately tenfold lower apparent transport rate constant relative to the apparent phosphorylation rate constant. This result indicated that the rate-limiting step of DGP synthesis is the transport of DG. A similar conclusion was reached for colon adenocarcinoma Caco-2 cells by use of radioactive DG (5). The congruency between the ratio of [DG] to [glucose] and the ratio of apparent initial rate of DGP synthesis to glucose consumption rate further indicated that glucose transport limits glucose consumption. The apparent initial rate of DGP synthesis, governed by the transport, was reduced by ~40% as a result of differentiation, indicating that the decreased rates of glucose consumption and of lactate production were a result of a similar decrease in glucose transport.

The parallel monitoring of DGP and the other soluble phosphate metabolites showed that accumulation of DGP led to a concomitant decrease in the phosphorylation potential and in the rate of glycolysis. This inhibition of glycolysis, beyond competitive substrate inhibition, appeared to be associated with the decrease in the phosphorylation potential. Similarly, a reduction in ATP in the presence of DG was previously suggested to account for the inhibition of glycolysis (26, 22). The removal of DG was followed by a slow and partial reduction of DGP but did not result in reversal of the phosphorylation potential. Similarly, it was previously shown that DG blocking of glucose consumption was not reversed despite a decrease in plasma DG (45). These results suggest a permanent DGP inhibition, presumably of glucose phosphoisomerase and hexokinase (22, 45). After removal of DG, DGP level decreased; the decay constant of this decrease (0.1-0.2 h-1) was considerably lower than the apparent rate constant of DG phosphorylation (~5 h-1) and was therefore neglected in the kinetic analysis of DGP synthesis.

Because glucose consumption was found to be rate limited by glucose transport, we monitored this process using the zero trans method for transport kinetic measurements (39). The application of this method using MRS was demonstrated for choline transport (25) and for cyclocreatine transport (31) in human breast cancer cells. In these studies as well as in our study, substrate phosphorylation was much faster than the transport, thus enabling the application of this kinetic method. Analysis of the kinetic data indicated a Michaelis-Menten mechanism of transport with distinctive Km and Vmax of transport. In control and differentiated cells, the Km was similar (2.9 mM), whereas Vmax was ~40% lower in the differentiated cells.

Analysis of the rates of lactate production as a function of increasing glucose concentration in the medium with a Michaelis-Menten mechanism yielded an apparent Km of ~2.7 mM in both control and differentiated cells. This value was similar to the Km value found for glucose transport in these cells, suggesting that the rate of lactate production via glycolysis was also limited by the rate of glucose transport.

Measurements of the changes in expression of the GLUTs as a result of differentiation by RA showed a small but consistent and highly significant decrease in the expression of total and membranal GLUT-1. The nonspecific staining in the total GLUT-1 measurements could mask the actual changes. However, the use of the exofacial directed antibody in nonpermeabilized cells, which ensured specific staining of membranal GLUT-1, yielded similar results. The 35-40% decrease in glycolysis in the presence of RA appears to be of the same order as the membranal reduction in GLUT-1 expression. However, we cannot rule out the possibility that deactivation of preexisting membranal GLUT-1 transporters is also responsible for the decrease in glucose transport rates, because changes in activation of preexisting membranal GLUT-1 in other cell types accounted for an increase in glucose transport (15, 36). The expression of GLUT-2, GLUT-3, and GLUT-4 appeared to be unrelated to the changes in glucose transport induced by RA. A correlation between GLUT-1 expression and glucose transport was previously observed in pancreatic tumors and in breast carcinoma (6, 21). However, a recent study in the human epidermoid carcinoma cell line (A431) and in T47D cells failed to show this correlation (1). In addition, in this work, T47D cells were found to express only mRNA of GLUT-1. Resolving these discrepancies requires further comprehensive kinetic studies and characterization of the GLUT expression in the systems described above.

Previous studies of MCF7 and MDA-MB-231 cells showed that GLUT-1, GLUT-2, and GLUT-5 were localized in the cytoplasm adjacent to the nucleus and in the plasma membrane. GLUT-3 and GLUT-4 were not detected in these cells (52). In contrast, a recent study indicated the presence of GLUT-3 in MCF7 cells but did not specify the localization of this transporter (18). Conflicting results on the localization of GLUT-1 in breast biopsies were also reported (3, 7, 49), whereas GLUT-3 was not detected in these biopsies (3, 7, 49). Thus, whether there is a specific presence and localization of the GLUT transporters in breast cancer is still an open question.

In the kinetic studies presented here, a single Km value of glucose transport was determined, of ~2.9 mM, which was not affected by RA. Because GLUT-1, GLUT-3, and GLUT-4 have a similar Km value, which differs from the Km of GLUT-2 (34, 40), it was not possible to distinguish the contributions of these three transporters. The lack of a high Km value for glucose transport is in accord with the absence of GLUT-2 in the cell membrane of T47D cells, as was found by immunofluorescence and by flow cytometry analysis. In two other human breast cancer cell lines, MCF7 and MDA-MB-468, two Km values of glucose transport were found, 2 and 10 mM, indicating the presence of GLUT-1 and GLUT-2, respectively (52).

Finally, because 31P NMR spectra provide a profile of all soluble phosphate metabolites, we have found a distinct increase in the levels of both GPC and GPE in the RA-differentiated cells. GPC and GPE are breakdown products of their corresponding phospholipids, phosphatidylcholine and phosphatidylethanolamine, respectively. The degradation of the phospholipids to GPC and GPE is catalyzed by phospholipases (A1 or A2) and lysophospholipase. Further degradation of these two metabolites to their free metabolites (choline and ethanolamine) is catalyzed by glycerophosphocholine phosphodiesterase. The increase in GPC and GPE levels can be explained by an increase in the activity of the phospholipases. Alternatively, inhibition of the phosphodiesterase can also lead to GPC and GPE accumulation. It appears that the former alternative is more favorable, because inhibition of the GPC and GPE breakdown should have resulted in a decrease in intracellular choline and ethanolamine and a concomitant decrease in PC and PE levels. However, such a decrease in PC and PE levels was not observed.

In summary, RA was shown to induce differentiation in T47D human breast cancer cells. This differentiation was accompanied by a reduction in the expression of the glucose transporter GLUT-1, as well as by a decrease in the rate of glucose metabolism via glycolysis, which was found to be rate limited by the transport of glucose. Thus we suggest that the reduction in the expression of membranal GLUT-1 led to a decrease in Vmax of glucose transport in the more differentiated cells. Because the expression of GLUT-1 varied according to the degree of cellular differentiation, it might correlate with the prognosis of breast cancer.


    ACKNOWLEDGEMENTS

We thank B. Geiger and A. Tchausovsky for their help with the immunofluorescence studies and W. Mueller-Klieser from Mainz University and Dr. F. Kohen for helpful discussions in the course of this work.


    FOOTNOTES

This work was supported by the German-Israel Foundation and by Sir David Alliance CBE, UK.

H. Degani is the incumbent of the Fred and Andrea Fallek Professorial Chair for Breast Cancer Research.

Address for reprint requests and other correspondence: H. Degani, Dept. of Biological Regulation, Weizmann Institute of Science, Rehovot 76100, Israel (E-mail: hadassa.degani{at}weizmann.ac.il).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.

Received 6 December 1999; accepted in final form 11 April 2000.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

1.   Aloj, L, Caraco C, Jagoda E, Eckelman WC, and Neumann RD. Glut-1 and hexokinase expression: relationship with 2-fluoro-2-deoxy-D-glucose uptake in A431 and T47D cells in culture. Cancer Res 59: 4709-4714, 1999[Abstract/Free Full Text].

2.   Baggetto, LG. Deviant energetic metabolism of glycolytic cancer cells. Biochimie 74: 959-974, 1992[ISI][Medline].

3.   Binder, C, Binder L, Marx D, Schauer A, and Hiddemann W. Deregulated simultaneous expression of multiple glucose transporter isoforms in malignant cells and tissues. Anticancer Res 17: 4299-4304, 1997[ISI][Medline].

4.   Birnbaum, MJ, Haspel HC, and Rosen OM. Transformation of rat fibroblasts by FSV rapidly increases glucose transporter gene transcription. Science 235: 1495-1498, 1987[ISI][Medline].

5.   Bissonnette, P, Gagne H, Blais A, and Berteloot A. 2-Deoxyglucose transport and metabolism in Caco-2 cells. Am J Physiol Gastrointest Liver Physiol 270: G153-G162, 1996[Abstract/Free Full Text].

6.   Brown, RS, Leung JY, Fisher SJ, and Frey KA. Intratumoral distribution of tritiated-FDG in breast carcinoma: correlation between GLUT-1 expression and FDG uptake. J Nucl Med 37: 1042-1047, 1996[Abstract].

7.   Brown, RS, and Wahl RL. Overexpression of Glut-1 glucose transporter in human breast cancer. Cancer 72: 2979-2984, 1993[ISI][Medline].

8.   Burant, CF, Sivitz WI, Fukumoto H, Kayano T, Magamatsu S, Seino S, Pessin JE, and Bell GI. Mammalian glucose transporters: structure and molecular regulation. In: Recent Progress in Hormone Research. New York: Academic, 1991, p. 349-387.

9.   Carruthers, A. Facilitated diffusion of glucose. Physiol Rev 70: 1135-1176, 1990[Free Full Text].

10.   Degani, H, Ronen SM, and Furman E. Breast cancer: spectroscopy and imaging of cells and tumors. In: NMR in Physiology and Biomedicine, edited by Gillies RJ. San Diego, CA: Academic, 1994, p. 329-351.

11.   Deuel, RK, Geneviene MY, Sherman WR, Schickner DJ, and Ackerman JJH Monitoring the time course of cerebral deoxyglucose metabolism by 31P nuclear magnetic spectroscopy. Science 228: 1329-1331, 1985[ISI][Medline].

12.   Dixon, M, and Webb EC. Enzymes. New York: Academic, 1964, p. 84-87.

13.   Eigenbrodt, E, Fister P, and Reinacher M. New perspectives on carbohydrate metabolism in tumor cells. In: Regulation of Carbohydrate Metabolism. Cleveland, OH: Chemical Rubber, 1985, p. 141-179.

14.   Eigenbrodt, E, Gerbracht U, Mazurek S, Presek P, and Friis R. Carbohydrate metabolism and neoplasia: new perspectives for diagnosis and therapy. In: Biochemical and Molecular Aspects of Selected Cancers. San Diego, CA: Academic, 1994, p. 311-385.

15.   Fisher, MD, and Frost SC. Translocation of GLUT1 does not account for elevated glucose transport in glucose-deprived 3T3-L1 adipocytes. J Biol Chem 271: 11806-11809, 1996[Abstract/Free Full Text].

16.   Flier, JS, Mueckler MM, Usher P, and Lodish HF. Elevated levels of glucose transport and transporter messenger RNA are induced by ras or src oncogenes. Science 235: 1492-1495, 1987[ISI][Medline].

17.   Furman, E, Rushkin E, Margalit R, Bendel P, and Degani H. Tamoxifen induced changes in MCF7 human breast cancer: in vitro and in vivo studies using nuclear magnetic resonance spectroscopy and imaging. J Steroid Biochem Molec Biol 43: 189-195, 1992[ISI][Medline].

18.   Grover-McKay, M, Walsh SA, Seftor EA, Thomas PA, and Hendrix MJC Role of glucose transporter 1 protein in human breast cancer. Phatol Oncol Res 4: 115-120, 1998.

19.   Gudas, LJ, Sporn MB, and Roberts AB. Cellular biology and biochemistry of the retinoids. In: The Retinoids., edited by Sporn MB, Roberts AB, and Goodman DS. New York: Raven, 1994, p. 443-520.

20.   Hawkins, RA, and Miller AL. Loss of radioactive 2-deoxy-D-glucose-6-phosphate from brains of conscious rats: implications for quantitative autoradiographic determination of regional glucose utilization. Neuroscience 3: 251-258, 1978[ISI][Medline].

21.   Higashi, T, Tamaki N, Honda T, Torizuka T, Kimura T, Inokuma T, Ohshio G, Hosotani R, Imamura M, and Konishi J. Expression of glucose transporters in human pancreatic tumors compared with increased FDG accumulation in PET study. J Nucl Med 38: 1337-1344, 1997[Abstract].

22.   Horton, RW, Meldrum BS, and Bachelard HS. Enzymic and cerebral metabolic effects of 2-deoxy-D-glucose. J Neurochem 21: 507-520, 1973[ISI][Medline].

23.   Jing, Y, Zhang J, Waxman S, and Mira-y-Lopaz R. Upregulation of cytokeratins 8 and 18 in human breast cancer T47D cells is retinoid-specific and retinoic acid receptor-dependent. Differentiation 60: 109-117, 1996[ISI][Medline].

24.   Kaplan, O, Navon G, Lyon RC, Faustino PJ, Straka EJ, and Cohen JS. Effects of 2-deoxyglucose on drug-sensitive and drug-resistant human breast cancer cells: toxicity and magnetic resonance spectroscopy studies of metabolism. Cancer Res 50: 544-551, 1990[Abstract].

25.   Katz-Brull, R, and Degani H. Kinetics of choline transport and phosphorylation in human breast cancer cells; NMR application of the zero trans method. Anticancer Res 16: 1375-1380, 1996[ISI][Medline].

26.   Koobs, DH. Phosphate mediation of the Crabtree and Pasteur effects. Science 178: 127-133, 1972[ISI][Medline].

27.   Liu, Y, Lee MO, Wang HG, Li Y, Hashimoto Y, Klaus M, Reed JC, and Zhang X. Retinoic acid receptor beta mediates the growth-inhibitory effect of retinoic acid by promoting apoptosis in human breast cancer cells. Mol Cell Biol 16: 1138-1149, 1996[Abstract].

28.   Lotan, R. Effects of vitamin A and its analogous (retinoids) on normal and neoplastic cells. Biochim Biophys Acta 605: 33-91, 1981[ISI].

29.   Lyon, RC, Cohen JS, Faustino PJ, Megnin F, and Myers CE. Glucose metabolism in drug-sensitive and drug-resistant human breast cancer cells monitored by magnetic resonance spectroscopy. Cancer Res 15: 870-877, 1988.

30.   Mangelsdorf, DJ, Umesono K, and Evans RM. The retinoid receptors. In: The Retinoids: Biology, Chemistry and Medicine, edited by Sporn MD, Roberts AB, and Goodman DS. New York: Raven, 1994, p. 319-349.

31.   Maril, N, Degani H, Rushkin E, Sherry AD, and Cohn M. Kinetics of cyclocreatine and Na(+) cotransport in human breast cancer cells: mechanism of activity. Am J Physiol Cell Physiol 277: C708-C716, 1999[Abstract/Free Full Text].

32.   Neeman, M, and Degani H. Metabolic studies of estrogen- and tamoxifen-treated human breast cancer cells by nuclear magnetic resonance spectroscopy. Cancer Res 49: 589-594, 1989[Abstract].

33.   Ojcius, DM, Degani H, Mispelter J, and Dautry-Varsat A. Enhancement of ATP levels and glucose metabolism during an infection by chlamydia. J Biol Chem 273: 7052-7058, 1998[Abstract/Free Full Text].

34.   Pessin, JE, and Bell GI. Mammalian facilitative glucose transporter family: structure and molecular regulation. Annu Rev Physiol 54: 911-930, 1992[ISI][Medline].

35.   Seewaldt, VL, Johnson BS, Parker MB, Collins SJ, and Swisshelm K. Expression of retinoic acid receptor beta mediates retinoic acid-induced growth arrest and apoptosis in breast cancer cells. Cell Growth Differ 6: 1077-1088, 1995[Abstract].

36.   Shetty, M, Loeb JN, Vikstrom K, and Ismail-Beigi F. Rapid activation of GLUT1 glucose transporter following inhibition of oxidative phosphorylation in clone 9 cells. J Biol Chem 268: 17225-17232, 1993[Abstract/Free Full Text].

37.   Sokoloff, L, Reivich M, Kennedy C, Des Rosiers MH, Patlak CS, Pettigrew KD, Sakurada O, and Shinohara M. The [14C]deoxyglucose method for the measurement of local cerebral glucose utilization: theory, procedure and normal values in the conscious and anesthetized albino rat. J Neurochem 28: 897-916, 1977[ISI][Medline].

38.   Sporn, MB, Dunlop NM, Newton DL, and Smith JM. Prevention of chemical carcinogenesis by vitamin A and its synthetic analogs (retinoids). Fed. Proc. 35: 1332-1338, 1976[ISI][Medline].

39.   Stein, WD. Kinetics of transport: analyzing, testing and characterizing models using kinetic approach. In: Methods in Enzymology, edited by Fleischer S, and Fleischer B. New York: Academic, 1989, p. 23-62.

40.   Thorens, B. Facilitated glucose transport in epithelial cells. Annu Rev Physiol 55: 591-608, 1993[ISI][Medline].

41.   Ting, Y-LT, Sherr D, and Degani H. Variations in energy and phospholipid metabolism in normal and cancer human mammary epithelial cells. Anticancer Res 16: 1381-1388, 1996[ISI][Medline].

42.   Toma, S, Isnardi L, Raffo P, Dastoli G, De Francisci E, Riccardi L, Palumbo R, and Bollag W. Effects of all-trans-retinoic acid and 13-cis-retinoic acid on breast-cancer cell lines: growth inhibition and apoptosis induction. Int J Cancer 70: 619-627, 1997[ISI][Medline].

43.   Warburg, O. On the origin of cancer cells. Science 123: 309-314, 1956[ISI].

44.   Wetherall, NT, and Taylor CM. The effects of retinoid treatment and antiestrogens on the growth of T47D human breast cancer cells. Eur J Cancer Clin Oncol 22: 53-59, 1986[ISI][Medline].

45.   Wick, AN, Drury DR, Nakada HI, and Wolfe JB. Localization of the primary metabolic block produced by 2-deoxyglucose. J Biol Chem 224: 963-969, 1957[Free Full Text].

46.   Wilcken, NRC, Sarcevic B, Musgrove EA, and Sutherland RL. Differential effects of retinoids and antiestrogens on cell cycle progression and cell cycle regulatory genes in human breast cancer cells. Cell Growth Differ 7: 65-74, 1996[Abstract].

47.   Wohlhueter, RM, and Plagemann PGW The roles of transport and phosphorylation in nutrient uptake in cultured animal cells. Int Rev Cytol 64: 171-240, 1980[ISI].

48.   Yamamoto, T, Seino Y, Fukumoto H, Koh G, Yano H, Inagaki N, Yamada Y, Inoune K, Manabe T, and Imura H. Over expression of facilitative glucose transporter genes in human cancer. Biochem Biophys Res Commun 170: 223-230, 1990[ISI][Medline].

49.   Younes, M, Brown RW, Mody DR, Fernandez L, and Laucirica R. GLUT1 expression in human breast carcinoma: correlation with known prognostic markers. Anticancer Res 15: 2895-2898, 1995[ISI][Medline].

50.   Younes, M, Lechago LV, Somoano JR, Mosharaf M, and Lechago J. Wide expression of human erythrocyte glucose transporter GLUT1 in human cancers. Cancer Res 56: 1164-1167, 1996[Abstract].

51.   Younes, M, Lechago LV, Somoano JR, Mosharaf M, and Lechago J. Immunohistochemical detection of GLUT3 in human tumors and normal tissues. Anticancer Res 17: 2747-2750, 1997[ISI][Medline].

52.   Zamora-Leon, SP, Golde DW, Concha II, Rivas CI, Delgado-Lopez F, Baselga J, Nualart F, and Vera JC. Expression of the fructose transporter GLUT5 in human breast cancer. Proc Natl Acad Sci USA 93: 1847-1852, 1996[Abstract/Free Full Text].


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