Department of Pediatric Surgery, Osaka University Graduate School of Medicine, Suita, Osaka 565-0871, Japan
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ABSTRACT |
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This study characterized the
Na+-dependent transport of L-glutamine by a
human neuroblastoma cell line, SK-N-SH. The Na+-dependent
component represented >95% of the total glutamine uptake. Kinetic
studies showed a single saturable high-affinity carrier with a
Michaelis constant (Km) of 163 ± 23 µM
and a maximum transport velocity (Vmax) of
13,713 ± 803 pmol · mg
protein1 · min
1. Glutamine uptake
was markedly inhibited in the presence of L-alanine, L-asparagine, and L-serine. Li+ did
not substitute for Na+. These data show that
L-glutamine is predominantly taken up through system
ASC. Glutamine deprivation resulted in the decrease of glutamine transport by a mechanism that decreased
Vmax without affecting
Km. The expression of the system ASC subtype
ASCT2 decreased in the glutamine-deprived group, whereas glutamine
deprivation did not induce changes in system ASC subtype ASCT1 mRNA
expression. Adaptive increases in Na+-dependent glutamate,
Na+-dependent 2-(methylamino)isobutyric acid, and
Na+-independent leucine transport were observed under
glutamine-deprived conditions, which were completely blocked by
actinomycin D and cycloheximide. These mechanisms may allow cells to
survive and even grow under nutrient-deprived conditions.
amino acid; system ASC; adaptive upregulation
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INTRODUCTION |
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AMINO ACID TRANSPORT across the plasma membrane is essential for supplying cells with amino acids for cellular metabolism (27). Transport is mediated via membrane proteins called carriers or transporters, which are responsible for the translocation of amino acid substrates from one side of the membrane to the other. Malignant cells display uncontrolled rates of cellular proliferation and require an increased supply of precursor amino acids to support key biosynthetic pathways (23). As a result, these cells have very efficient transport systems and can transport amino acids across the plasma membrane faster than normal cells (23). Human hepatoma cells, for example, transport glutamine at a rate 10-20 times faster than normal hepatocytes (5).
Glutamine provides nitrogen for a number of important precursors for macromolecule synthesis including purines, pyrimidines, amino sugars, and some amino acids (11). In addition, it is an important fuel for cancer cells. Glutamine metabolism may also supply precursors for the synthesis of glutathione, which serves as a major store of cellular reducing equivalents (3). In cancer cells, cell-growth rates and the biosynthesis of DNA and protein correlate directly with glutamine concentration in culture media (32). A significant correlation has also been found between glutamine transport and its disappearance rates (9). These findings suggest that glutamine transport is rate limiting for glutamine utilization.
Neuroblastomas are childhood tumors that are derived from neural crest cells. They are biologically remarkable in that some neuroblastomas regress spontaneously without chemotherapy, and spontaneous and induced maturation is seen with significant frequency (29). In contrast, many other neuroblastoma tumors show invasive and progressive growth behavior. The prognosis for this tumor can be determined using genetic markers such as a short-arm deletion on chromosome 1p (7), increased ploidy (1), and amplification of the N-myc oncogene (25). Although many tumor markers have been investigated with respect to the biology of neuroblastomas, a definitive and consistent causal pattern for the diverse behavior and variable biology of these tumors is still unexplained. Hannuniemi et al. (16) showed that the maximal velocities of the saturable influx of leucine, lysine, and glycine were greater in cultured neuroblastoma cells than in glioma cells. The contribution of the amino acid transport system (system A) is greater in a neuroblastoma cell line with no N-myc amplification (8). Because of the potential role of glutamine as a mediator of tumor growth, glutamine transport across the plasma membrane may be rate limiting for subsequent metabolic events in neuroblastomas. However, glutamine uptake by neuroblastoma cells has not been previously investigated.
Several studies have shown the adaptive regulation of amino acid transport in cells subjected to amino acid starvation (4, 10, 22, 30). For example, glutamine deprivation causes a significant increase in glutamine and glutamate transport velocity (22, 30). In neuroblastoma cells, the removal of glutamine from the culture medium resulted in a 10-fold increase in the specific activity of the enzyme glutamine synthetase compared with the basal level, which suggests a high capacity for glutamine utilization (20). It remains unknown how neuroblastoma cells regulate amino acid transport activities when the availability of key nutrients such as glutamine is limited.
The purpose of this study was to characterize glutamine transport across the plasma membrane of human neuroblastoma cells. In addition, we examined the effects of glutamine deprivation on the transport of glutamine and other amino acids. We used an SK-N-SH human neuroblastoma cell line because it provided a well-characterized in vitro model system in which to study the growth mechanism.
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METHODS |
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Materials. The radiolabeled amino acids L-[3H]glutamine, L-[3H]glutamate, and L-[3H]leucine and the system A-specific substrate 2-[3H](methylamino)isobutyric acid (MeAIB) were obtained from Amersham (Arlington Heights, IL). Cell culture media were from GIBCO-BRL (Gaithersburg, MD). Amino acids and all biochemicals were purchased from Sigma Chemical (St. Louis, MO) and fetal bovine serum (FBS) was from JRH Biosciences (Lenexa, KS). Tissue-culture plates were obtained from Corning (New York, NY). Neuroblastoma cell line SK-N-SH was provided by Dr. Tadao Ohno (RIKEN Cell Bank, Tsukuba, Japan).
Cell culture. Neuroblastoma cells were cultured at 37°C in a humidified atmosphere of 5% CO2-95% air. Cells were maintained in DMEM supplemented with 2 mM L-glutamine, 10% FBS, 1,000 U/ml penicillin, and 1,000 U/ml streptomycin. The culture medium was changed every 3 days until cells were confluent. Cells were then used for experiments.
Amino acid transport measurement. Transport of L-[3H]glutamine was measured by the cluster-tray method of Gazzola et al. (13). Before use in transport assays, cells were rinsed twice with warm Na+-free Krebs-Ringer phosphate buffer (CholKRP, made by replacing corresponding Na+ salts with choline chloride and choline phosphate) to remove extracellular Na+ and amino acids. Radiolabeled glutamine (5 µCi of [3H]glutamine/ml) transport assays were performed for 1 min at 37°C with 10 µM unlabeled glutamine in both Na+-Krebs-Ringer phosphate (NaKRP) and CholKRP buffers. The transport reaction was terminated by discarding the uptake buffer and rinsing the cells three times with ice-cold CholKRP buffer (2 ml per well per rinse). The wells containing the cells were allowed to dry and were then solubilized in 200 µl of 0.2 N NaOH-0.2% SDS solution. The cell extract (100 µl) was neutralized with 10 µl of 2 N HCl and subjected to scintillation spectrophotometry. The remaining 100 µl in each well was used for a bicinchoninic acid protein assay (28). Na+-dependent glutamine transport values were obtained by subtracting the transport values in CholKRP from those in NaKRP. Saturable Na+-independent transport values were determined in CholKRP by subtracting the values in the presence of excess (10 mM) unlabeled glutamine from those in its absence. Transport of L-[3H]leucine, L-[3H]glutamate, and [3H]MeAIB was measured by the same method. Transport velocities were expressed in picomoles per milligram of protein per minute. Data (means ± SD) were analyzed and compared with Student's t-test or one-way ANOVA. P < 0.05 was considered statistically significant.
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RESULTS |
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Time course of [3H]glutamine transport. Na+-dependent glutamine uptake was found to be linear for at least 3 min, and the Na+-dependent component was shown to account for >95% of the total glutamine uptake at all time points. Therefore, 1 min was chosen for the measurement of the initial rate of Na+-dependent glutamine transport for subsequent experiments.
Kinetics of [3H]glutamine transport.
Na+-dependent glutamine uptake was determined at glutamine
concentrations between 10 µM and 1 mM in both Na+ and
choline buffers. A saturation plot of uptake velocity as a function of
glutamine concentration is shown in Fig.
1A. When an Eadie-Hofstee plot
was created for each kinetic study, the data were found to fit a single
linear regression line. A representative plot is shown in Fig.
1B. When the data from three separate kinetics studies were
averaged, they demonstrated a single high-affinity transport carrier
for glutamine with a Michaelis constant (Km) of
163 ± 23 µM and a maximum transport velocity
(Vmax) of 13,713 ± 803 pmol · mg
protein1 · min
1.
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Amino acid inhibition of [3H]glutamine transport.
To characterize the Na+-dependent glutamine transport
system, glutamine transport was measured in the presence of 5 mM
concentrations of selected amino acids. The osmotic effects of
inhibitors were compensated for by the addition of 5 mM sucrose to
control assays. The data are expressed as a percentage of the control
rate of glutamine uptake (Fig. 2).
Na+-dependent glutamine transport was significantly
inhibited by L-glutamine, L-alanine,
L-serine, and L-asparagine (substrates for
system ASC; P < 0.001) but was unaffected by
L-glutamate and L-arginine. Neither MeAIB, a
highly specific substrate for system A, nor
2-aminobicyclo-(2,2,1)-heptane-2-carboxylic acid, a
specific substrate for the Na+-independent system L,
demonstrated significant inhibition of glutamine uptake.
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Effects of external pH on [3H]glutamine transport.
Na+-dependent glutamine transport was measured at several
pH values between 6.0 and 8.0. The pH of the extracellular medium was
adjusted by adding varying amounts of HCl or NaOH.
Na+-dependent glutamine transport significantly decreased
at pH 6.0 (492 ± 89 pmol · mg
protein1 · min
1) and at pH 6.5 (552 ± 14 pmol · mg
protein
1 · min
1) compared with that
observed at pH 7.5 (992 ± 51 pmol · mg
protein
1 · min
1;
P < 0.01). Transport was insensitive to
extracellular pH change between 7.0 (927 ± 50 pmol · mg
protein
1 · min
1) and 8.0 (820 ± 68 pmol · mg
protein
1 · min
1).
Effects of cations on [3H]glutamine transport.
To examine the ability of the transporter to exchange Li+
for Na+, Na+-dependent glutamine transport was
measured in the presence of 119 mM NaCl, LiCl, and choline chloride
(Table 1). Na+-dependent
glutamine transport significantly decreased in the presence of
Li+ and choline compared with Na+
(P < 0.001).
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Effects of glutamine concentration on cell growth.
The effect of glutamine concentration on cell growth was determined.
Cells were seeded at a density of 1 × 105 cells/ml (1 ml/well) into 12-well tissue-culture plates. After 24 h, the
culture medium was removed and changed to glutamine-free DMEM
supplemented with 10% FBS plus various concentrations of glutamine (2 mM and 400, 200, 100, and 0 µM). Cells were detached from the plate
with trypsin and counted at days 0, 1, 2, and 3 with a hemocytometer. Cell growth in 2 mM glutamine was chosen as the
control. As shown in Fig. 3, cell-growth
rates were dependent on glutamine concentrations. In 200 µM
glutamine, cells grew about half as fast as controls. Cells showed slow
growth even in 0 µM glutamine, and this glutamine concentration was
used for subsequent experiments as the glutamine-deprived condition.
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Time-dependent effects of glutamine deprivation on
[3H]glutamine transport.
Cells were seeded into 24-well tissue-culture plates (0.5 ml/well). At
100% cell confluence, the culture medium was removed and changed to
glutamine-free DMEM supplemented with 10% FBS plus 2 (control) or 0 (glutamine deprivation) mM glutamine. Na+-dependent
glutamine transport was measured at 0, 8, 16, and 24 h. As shown
in Fig. 4A,
Na+-dependent glutamine transport in the glutamine-deprived
cells decreased significantly compared with control at 8 h
(control, 1,320 ± 14; glutamine deprivation, 1,159 ± 186 pmol · mg
protein1 · min
1;
P < 0.05), 16 h (control, 1,638 ± 26;
glutamine deprivation, 1,185 ± 77 pmol · mg
protein
1 · min
1; P < 0.01), and 24 h (control, 1,424 ± 33; glutamine
deprivation, 983 ± 154 pmol · mg
protein
1 · min
1; P < 0.01). To determine the kinetics of the glutamine-deprived effects
on glutamine transport, the transport of glutamine from 10 µM to 1 mM
was determined in both control and glutamine-deprived cells (Fig.
4B). There was a significant decrease in maximum transport velocity in the glutamine-deprived group compared with control (Vmax: control, 13,713 ± 803; glutamine
deprivation, 9,553 ± 646 pmol · mg
protein
1 · min
1; P < 0.01), but no significant change was observed in transport affinity
(Km: control, 163 ± 23; glutamine
deprivation, 181 ± 18 µM).
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Effects of glutamine deprivation on expression of system ASC
subtypes.
The effect of glutamine deprivation on the expression of different
subtypes of system ASC (ASCT1 and ASCT2) was analyzed using RT-PCR
(Fig. 5). Cells were seeded at a density
of 1 × 105 cells (2 ml/well) into six-well
tissue-culture plates. After 100% cell confluence was attained, the
culture medium was removed and replaced with glutamine-free DMEM
supplemented with 10% FBS plus 2 (control) or 0 (glutamine
deprivation) mM glutamine. At 0, 12, and 24 h after the medium was
changed, total RNA was extracted from the cells using a RNAce Total
Pure Purifications kit (Bioline, London, UK). The amount of extracted
RNA was calculated from optical density measurements at 260 nm. RNA
(1.0 µg) was used to generate first-strand complementary DNA using an
Advantage RT for PCR kit (Clontech, Palo Alto, CA) according to the
manufacturer's recommended procedures. RT-PCR was performed using the
same complementary DNA samples with primer pairs specific for human
ASCT1, ASCT2, and glyceraldehyde-3-phosphate dehydrogenase (GAPDH)
using a Takara PCR Thermal Cycler. The primer pairs used were (forward)
5'-GAG CTC AAC GCA GGA CAG ATT-3', (reverse) 5'-AGG ATC AGA GGC AGG TCA TGA-3' for ASCT1; (forward) 5'-CAA GGA GGT GCT CGA TTC GT-3', (reverse)
5'-ACC CTG GTT CCG GTG ATA TTC-3' for ASCT2; and (forward) 5'-GAA GGT
GAA GGT CGG-3', (reverse) 5'-GAA GAT GGT GAT GGG-3' for GAPDH.
Amplification was initiated by a 10-min denaturation at 95°C followed
by 40 cycles at 95°C for 15 s and 60°C for 60 s. After
the last cycle, the samples were incubated for 10 min at 72°C. The
PCR products were then visualized by ultraviolet illumination after
electrophoresis through 1.6% agarose gels containing 0.5 µg/ml
ethidium bromide. The gel photographs were scanned with a computerized
densitometer. Semiquantitative analysis of ASCT1 and ASCT2 was
performed by comparison with the housekeeping gene GAPDH.
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Effects of glutamine deprivation on amino acid transport and
effects of actinomycin D and cycloheximide treatment on induction of
amino acid transport by glutamine deprivation.
Cells were seeded into 24-well tissue-culture plates (0.5 ml/well). At
100% cell confluence, the culture medium was removed and changed to
glutamine-free DMEM supplemented with 10% FBS plus 2 (control) or 0 (glutamine deprivation) mM glutamine in the absence or presence of
either actinomycin D (4 µM) or cycloheximide (20 µM). After 24 h, uptake of [3H]glutamine, [3H]glutamate,
[3H]leucine, and [3H]MeAIB was measured
using the same method as described for glutamine transport measurement.
The transport of these amino acids was linear for at least 3 min, and
Na+-dependent glutamate (system X
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DISCUSSION |
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Our study is the first to characterize the details of the glutamine transport system in a human neuroblastoma cell line. Ninety-five percent of glutamine transport by SK-N-SH cells occurred via carrier-mediated Na+-dependent transport pathways. Kinetic analysis demonstrated a single high-affinity transport carrier for glutamine with a Km of 163 µM. This transporter has high affinity, because the circulating concentration of glutamine is 600-700 µM or about four times greater than the Km. Thus even when circulating concentrations of glutamine are very low, such as in a poorly vascularized portion of a solid tumor, the glutamine carrier should still be saturated. This ability to utilize glutamine may be essential to support proliferation, energy metabolism, and protein synthesis.
In the past, a large number of mammalian amino acid transport systems were studied using membrane vesicle preparations or cultured cells (6, 12, 17, 19). These included groups of Na+-dependent amino acid transporters that utilize free energy stored as a Na+ electrochemical potential gradient across plasma membranes for the uphill transport of amino acids. The Na+-dependent transporters play a central role in amino acid mobilization in animals because of their ability to transport amino acids against a concentration gradient. Criteria used to differentiate amino acid transport systems in mammalian cells to include substrate specificity, analog cross-inhibition patterns, transport kinetics, and ion dependency (27). Because MeAIB did not interfere with glutamine transport, system A is not an important transport system for glutamine. Another Na+-dependent transporter, which is named system N for the nitrogen-containing side chain of its substrates, is characterized by histidine transport inhibition of glutamine (19). In our studies, histidine only partially blocked glutamine transport, and Li+ did not substitute for Na+ as a cation (as it does for system N). The inability of Li+ to support glutamine transport in conjunction with the inhibitor profile indicates that system N is not a major component of glutamine transport. In summary, based on the inability of this high-affinity carrier to transport glutamine in the presence of alanine, serine, or asparagine, its intolerance to N-methylated substrate, and the failure of Li+ to substitute for Na+, we conclude that glutamine is predominantly taken up through system ASC in SK-N-SH cells.
The relative pH insensitivity of system ASC has been a useful parameter in differentiating it from systems A and N (27). However, in our studies, glutamine transport decreased at pH 6.0 and 6.5 compared with pH 7.5. Handlogten et al. (15) obtained similar results and reported that system ASC was inhibited when the pH was lowered from 6.5 to 5 in a hepatoma cell line. In the brush-border membrane vesicles of rabbit ileum, system ASC increased at lower pH (24). Therefore, the pH sensitivity of system ASC is not specific to the SK-N-SH neuroblastoma cell line. Our finding suggests that pH changes may serve as important regulatory steps in subsequent intracellular metabolism in SK-N-SH cells.
Glutamine deprivation resulted in the decrease of glutamine transport
by a mechanism that decreased Vmax without
affecting Km (see Fig. 4). This indicates that
the number of active glutamine transporters in the cell membrane
decreased without affecting the affinity of the transporter. An
increase in the number of active amino acid transporter proteins in the
cell membrane usually involves an increase in de novo protein synthesis
of the carrier itself (27). Therefore, the decrease in
glutamine transporters may occur secondary to a decrease in the rate of
de novo carrier biosynthesis or an increase in the rate of carrier
breakdown. In contrast to glutamine transport, SK-N-SH cells responded
to glutamine deprivation by increasing the transport activities of leucine, glutamate, and MeAIB. Na+-independent leucine,
Na+-dependent glutamate, and Na+-dependent
MeAIB are transported via systems L, X
The first mammalian glutamine transporter gene was isolated in 1996 from a mouse testis cDNA library; it encoded a 553-amino acid protein with functional properties of system ASC (31). It was called ASCT2 to distinguish it from ASCT1, a system ASC isoform isolated in 1993 that does not transport glutamine (2, 26). ASCT2, which exhibits 57% sequence identity to ASCT1, takes up glutamine with high affinity and transports a wide panel of other amino acids including serine, threonine, cysteine, alanine, and asparagine as well as branched-chain amino acids (leucine, valine, and isoleucine) to a lesser degree (31). There have been no studies of the molecular regulation of system ASC transporter genes in amino acid-deprived conditions. In this study, ASCT2 mRNA expression decreased in the glutamine-deprived group, whereas glutamine deprivation did not induce changes in ASCT1 mRNA expression, which shows that the decrease in ASCT2 mRNA expression is comparable to that observed for the glutamine transport activity in the glutamine-deprived group. Therefore, we conclude that glutamine is taken up via a system ASC subtype, ASCT2, and downregulation of glutamine transport induced by glutamine deprivation is due to modifications in the expression of ASCT2-related genes.
The increased amino acid transport that was induced by
glutamine deprivation was completely blocked by the RNA synthesis
inhibitor actinomycin D and by the protein synthesis inhibitor
cycloheximide. In our unpublished data, leucine transport increased in
the glutamine-deprived group by a mechanism that increased
Vmax. Amino acid starvation increased the
expression of the ATA2 system A transporter gene in human fibroblasts
(14) and rat glioma cells (21). Therefore, adaptive upregulation of amino acid transport may result from the
emergence of new carriers after enhanced transcriptional activity, possibly associated with increased gene expression of each amino acid
transporter. The enhanced activity of systems A, L, and
X
As shown in Fig. 3, the growth of SK-N-SH cells was less sensitive to glutamine deprivation than has been reported for other cancer cell lines (32). The SK-N-SH cells survived and even grew in glutamine concentrations lower than normal circulating levels (500-600 µM). The centers of solid tumors are generally poorly vascularized, and intercellular amino acid concentrations can be much lower than the normal circulating levels (27). Decreased extracellular amino acid levels encountered by tumors in vivo may elicit similar adaptive responses in amino acid transport that contribute to the maintenance of cytoplasmic amino acid levels that are essential for growth.
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FOOTNOTES |
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Address for reprint requests and other correspondence: M. Wasa, Dept. of Pediatric Surgery, Osaka Univ. Graduate School of Medicine, 2-2 Yamadaoka, Suita, Osaka 565-0871, Japan (E-mail: wasa{at}pedsurg.med.osaka-u.ac.jp).
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. Section 1734 solely to indicate this fact.
First published January 9, 2002;10.1152/ajpcell.00324.2001
Received 17 July 2001; accepted in final form 2 January 2002.
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