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The mammalian target of rapamycin (mTOR) integrates nutrient and mitogen signals to regulate cell growth (increased cell mass and cell size) and cell division. The immunosuppressive drug rapamycin inhibits cell cycle progression via inhibition of mTOR; however, the signaling pathways by which mTOR regulates cell cycle progression have remained poorly defined. Here we demonstrate that restoration of mTOR signaling (by using a rapamycin-resistant mutant of mTOR) rescues rapamycin-inhibited G1-phase progression, and restoration of signaling along the mTOR-dependent S6K1 or 4E-BP1/eukaryotic translation initiation factor 4E (eIF4E) pathways provides partial rescue. Furthermore, interfering RNA-mediated reduction of S6K1 expression or overexpression of mTOR-insensitive 4E-BP1 isoforms that block eIF4E activity inhibit G1-phase progression individually and additively. Thus, the activities of both the S6K1 and 4E-BP1/eIF4E pathways are required for and independently mediate mTOR-dependent G1-phase progression. In addition, overexpression of constitutively active mutants of S6K1 or wild-type eIF4E accelerates serum-stimulated G1-phase progression, and stable expression of wild-type S6K1 confers a proliferative advantage in low-serum-containing media, suggesting that the activity of each of these pathways is limiting for cell proliferation. These data demonstrate that, as for the regulation of cell growth and cell size, the S6K1 and 4E-BP1/eIF4E pathways each represent critical mediators of mTOR-dependent cell cycle control.
Although cell growth (an increase in cell mass and cell size through macromolecular biosynthesis) and cell division are distinct processes and therefore separable under some conditions (13, 22, 32), they are generally tightly coupled such that cell mass and DNA content double during each cell division cycle. Such a mechanism ensures that appropriately sized daughter cells are produced after mitosis. Although the mechanisms by which cell growth and cell cycle division are coordinated are poorly understood, the signaling protein TOR (for target of rapamycin; also known as FRAP, RAFT, or RAPT in mammals) regulates both cell growth and cell cycle progression in species from yeast to flies to mammals and as such is recognized as an evolutionarily conserved central coordinator of these fundamental biological processes (reviewed in references 9, 19, 34, and 46).
TOR belongs to the phosphatidylinositol kinase-related kinase superfamily in which a lipid kinase homology domain functions as a serine/threonine kinase. When complexed with its cellular receptor FK506-binding protein 12 (FKBP12), the immunosuppressive drug rapamycin directly binds to TOR, resulting in inhibition of TOR-dependent downstream signaling (reviewed in references 15 and 46). Rapamycin treatment induces G1-phase arrest in yeast cells and mammalian lymphocytes; in most other cell types, however, the drug delays cell cycle progression rather than inducing an absolute block (reviewed in reference 1). Recently, TOR has also been linked to regulation of cell growth. In flies, larvae null for Drosophila melanogaster TOR (dTOR) are reduced in size, and dTOR-null cells are reduced in size (35, 61). In mammalian cells, rapamycin reduces cell size, and restoration of mammalian TOR (mTOR) signaling rescues this reduced cell size phenotype (13). Thus, TOR signaling regulates both cell cycle progression and cell growth/cell size.
In yeast, TOR monitors and responds to nutrient levels (reviewed in reference 39). In more complex multicellular organisms, however, TOR integrates signals from both nutrients and growth factors (reviewed in reference 46). How nutrients regulate TOR is poorly understood. A novel positive regulator of TOR, the small GTPase Rheb (named for Ras homologue enriched in brain), has recently been shown to function as an integrator of both nutrient and mitogenic signals (14, 43, 49, 54, 62). The tuberous sclerosis complex proteins TSC1/TSC 2 (hamartin/tuberin) negatively regulate TOR by inactivating Rheb through TSC2's (tuberin's) GTPase activating protein (GAP) activity (14, 54, 62; reviewed in reference 31). Mitogens, through direct phosphorylation of TSC2 (tuberin) by Akt/protein kinase B, inhibit the tumor suppressor function of the tuberous sclerosis complex, thereby indirectly promoting TOR-dependent signaling (reviewed in reference 31).
The best-characterized downstream effectors of mTOR include two signaling pathways that act in parallel to control mRNA translation: the 70-kDa ribosomal protein S6 kinase 1 (p70S6K1 or S6K1) pathway and the eukaryotic translation initiation factor 4E (eIF4E)-binding protein 1 (4E-BP1; also known as PHAS-I)/eIF4E pathway. mTOR-dependent signals, in cooperation with phosphatidylinositol 3-kinase-dependent signals, mediate phosphorylation and activation of S6K1 and phosphorylation and inactivation of 4E-BP1 (a repressor of translation initiation) (reviewed in references 15 and 29). S6K1 directly phosphorylates the 40S ribosomal protein S6, which is thought to increase the translation of mRNA species that possess a 5′-terminal oligopyrimidine (5′-TOP) (20, 21, 56). Since ribosomal proteins and translation elongation factors are encoded by 5′-TOP mRNAs, signaling along the S6K1 pathway may promote ribosome biogenesis and therefore enhance protein biosynthetic capacity, although such a model has been questioned (50, 53).
Hypophosphorylated 4E-BP1 binds to and inhibits the rate-limiting translation initiation factor eIF4E, which binds the cap (m7GpppN) at the 5′ ends of mRNA transcripts to initiate cap-dependent translation (reviewed in references 15 and 29). A recently identified mTOR-interacting protein, raptor, links mTOR to substrates such as 4E-BP1 (16, 25) by binding to the TOR signaling (TOS) motif in the C terminus of 4E-BP1 (either directly or indirectly), enabling mTOR-dependent phosphorylation of 4E-BP1 in response to nutrients or growth factors (8, 33, 45, 59). Upon phosphorylation, 4E-BP1 releases from eIF4E, allowing eIF4E to assemble with other translation initiation factors to initiate cap-dependent translation. eIF4E is thought to enhance the translation of transcripts possessing either complex 5′-untranslated region (UTR) secondary structure and/or upstream open reading frames, which often encode proteins associated with a proliferative response (reviewed in reference 48).
Although some experiments have suggested a critical role for S6K1 in S-phase progression (26), mice null for S6K1, although slightly smaller than wild-type (WT) littermates at birth, exhibit few abnormalities (47), and S6K1-null embryonic stem cells exhibit only a mild reduction in cell proliferation rate (23). Thus, although S6K1 is generally thought to play a role in cell cycle progression, its contribution relative to other mTOR-mediated signals is unclear. A potential role for 4E-BP1/eIF4E in cell cycle progression has not been carefully investigated, although eIF4E overexpression transforms rodent fibroblasts (27) and is commonly observed to be overexpressed in human tumors (reviewed in reference 51). We therefore investigated the individual roles of S6K1, 4E-BP1, and eIF4E in mammalian cell cycle progression. We previously reported that the S6K1 and eIF4E pathways function independently to increase cell size downstream of mTOR (13). We now report that these pathways each promote mTOR-dependent cell cycle progression and that the activity of each pathway is rate limiting for cell cycle progression in mammalian cells. The function of these biosynthetic pathways as mediators of cell cycle progression is consistent with a role for these pathways in coupling cell growth with cell cycle progression during cell proliferation.
Rapamycin was kindly provided by S. N. Seghal (Wyeth). RNase A and Fugene 6 transfection reagent were from Roche, and nitrocellulose membrane was from Schleicher & Schuell. X-ray film was from Kodak (X-OMAT LS). All other chemicals were from Sigma.
Anti-AU1 monoclonal antibodies were from Covance, and anti-CD20-fluorescein isothiocyanate (FITC) monoclonal antibodies were from BD Pharmingen. Antihemagglutin in (anti-HA) monoclonal antibodies were kindly provided by Margaret Chou (University of Pennsylvania, Philadelphia), and anti-phospho-S6 antibodies were generously provided by Morris Birnbaum (University of Pennsylvania and Howard Hughes Medical Institute, Philadelphia). Anti-eIF4E antibodies were from Cell Signaling Technology. Anti-S6K1 (6) and anti-mitogen-activated protein kinase (anti-MAPK) (7) antibodies have been described. Anti-green fluorescent protein (anti-GFP) monoclonal antibodies were from Sigma, and anti-chloramphenicol acetyltransferase (anti-CAT) antibodies were from Cortex Biochem (formerly 5-Prime 3-Prime). For immunoblotting, anti-rabbit and anti-mouse horseradish peroxidase-conjugated secondary antibodies were from Amersham and Chemicon, respectively.
pcDNA3/AU1-mTOR eukaryotic expression plasmids encoding WT, rapamycin-resistant (RR; Ser2035Ile), kinase-dead (KD; Asp2338Ala), and double RR/KD alleles of rat mTOR were kindly provided by Robert Abraham (Burnham Institute, San Diego, Calif.) and have been described elsewhere (5). pRK7/HA-S6K1 eukaryotic expression plasmids encoding WT and KD (Lys100Arg) alleles of rat S6K1 (αII) have been described previously (6). The two partial rapamycin-resistant mutants of S6K1, E389D3E (37) and E389ΔCT (44), have been described. E389D3E, containing the point mutations T389E, S411D, S418D, T421E, and S424D, was generated in this lab but was originally described elsewhere (37). pCMV/CD20 was provided by Ed Harlow (Harvard Medical School, Boston, Mass.), and the bicistronic plasmid pGFP/CAT was provided by A. A. M. Thomas (University of Utrecht). The plasmids pMV7/3HA-eIF4E, pACTAG2/HA-WT-4E-BP1, and pACTAG2/HA-AA-4E-BP1 (Thr37/46Ala) were generously provided by Nahum Sonenberg (McGill University, Montreal, Quebec, Canada). Plasmid pACTAG2/3HA-F114A-4E-BP1 has been described (44), and pACTAG2/3HA-AA-Y54A-4E-BP1 (Thr37/46Ala; Tyr54Ala) was generated by using QuickChange (Stratagene) with the AA-4E-BP1 plasmid as a template. pRK7/3HA-eIF4E was generated by PCR amplifying 3HA-eIF4E from pMV7 and subcloning it into pRK7. The hairpin interfering RNA (RNAi) plasmid targeting S6K1 (pBS/U6-S6K1) was generated as follows. The sequence below (the sense strand only is shown) was cloned into pBS/U6 (52) between the ApaI and EcoRI restriction sites to express hairpin RNA from the U6 promoter. In boldface is the targeting DNA sequence (nucleotides 284 to 304 of human S6K1): 5′-GGGTACTTGGTAAAGGGGGCTTTCAAGCTTAGCCCCCTTTACCAAGTACCCTTTTT-3′.
Human U2OS osteosarcoma cells were cultured at 37°C and 5% CO2 in Dulbecco modified Eagle medium (DMEM)-10% fetal bovine serum (FBS). A total of 1.5 × 105 cells were seeded onto 60-mm plates and then transfected 24 h later with Fugene 6 overnight according to the manufacturer's directions by using 5 to 10 μg of total DNA, depending on the experiment. Cells were then washed once with DMEM, deprived of serum for 30 h (DMEM plus 20 mM HEPES [pH 7.2]), and refed with DMEM-10% FBS in the absence or presence of rapamycin at 20 ng/ml for 20 h unless otherwise noted.
Cells were washed once with ice-cold STE (pH 7.2; 150 mM NaCl, 50 mM Tris-HCl, 1 mM EDTA), scraped in lysis buffer (pH 7.2; 10 mM KPO4, 1 mM EDTA, 10 mM MgCl2, 50 mM β-glycerophosphate, 5 mM EGTA, 0.5% NP-40, 0.1% Brij-35, 1 mM sodium orthovanadate, 40 μg of phenylmethylsulfonyl fluoride/ml, 10 μg of leupeptin/ml, 5 μg of pepstatin A/ml), and spun at 15,000 rpm for 5 min. Lysates were subjected to sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE), transferred to nitrocellulose membrane, immunoblotted with primary antibodies followed by horseradish peroxidase-conjugated secondary antibodies, and developed via enhanced chemiluminescence. A Bradford assay was used to determine the protein content (Bio-Rad).
Cell extracts were immunoprecipitated with anti-HA antibodies for 2 h, followed by incubation with protein A-Sepharose CL4B (Pharmacia) for 1 h. Immunoprecipitates were washed with 1 ml each of ice-cold buffer A, buffer B, and buffer ST (salt-Tris) as described previously (30). The kinase activity in washed immunoprecipitates was assayed with recombinant GST-S6 (32 C-terminal amino acids of ribosomal protein S6) as in vitro substrate, as described previously (30). The amount of 32P incorporated into glutathione S-transferase (GST)-S6 was assessed by autoradiography and quantitated on a Bio-Rad phosphorimager with ImageQuant software.
U2OS cells were plated to glass coverslips in 35-mm wells at 0.5 × 105 cells/well and cotransfected 24 h later with Fugene 6 with 0.2 μg of pCMV/CD20 and 2.0 μg of pBS/U6 or pBS/U6-S6K1. After overnight transfection, the cells were serum deprived for 30 h and refed with DMEM-10% FBS for 20 h. Coverslips were washed twice in 2 ml of phosphate-buffered saline plus Ca and Mg (PBS+Ca/Mg), incubated in anti-CD20-FITC antibodies diluted 1:10 in PBS+Ca/Mg 30 min, washed twice in PBS+Ca/Mg, and then fixed 10 min in 3.7% formaldehyde (Polysciences, Inc.) in PBS+Ca/Mg. After fixation, the coverslips were washed, blocked and/or permeabilized for 30 min in PBS+Ca/Mg-0.2% fish skin gelatin (FSG)-0.2% Triton X-100 (TX-100), and incubated for 30 min with anti-phospho-S6 antibodies diluted 1:500 in PBS+Ca/Mg-FSG-TX-100. Coverslips were then washed, incubated for 30 min in donkey anti-rabbit Texas red (1:500; Jackson Laboratories) in PBS+Ca/Mg-FSG-TX-100, washed once in PBS+Ca/Mg-FSG plus DAPI (4′,6′-diamidino-2phenylindole) at 0.1 μg/ml, washed twice in PBS+Ca/Mg-FSG and twice in PBS+Ca/Mg, dipped in distilled H2O, mounted on CitiFluor (Ted Pella), and finally sealed with nail polish. Immunofluorescence images were acquired by using a Nikon Eclipse E800 upright epifluorescence microscope equipped with an Orca 100 camera and Metamorph software.
Cell extracts were incubated in 20 μl of 7-methyl GTP-Sepharose 4B (Amersham Pharmacia) for 2 h at 4°C and then washed twice in lysis buffer. Sepharose beads were resuspended in Laemmli sample buffer with 2% β-mercaptoethanol and resolved by SDS-PAGE.
U2OS cells were cotransfected with 0.5 μg of pACTAG2/3HA-4E-BP1 plasmids and 5 μg of the bicistronic plasmid pGFP/CAT, serum deprived for 30 h, and refed with DMEM-10% FBS in the absence or presence of rapamycin for 20 h. [35S]methionine-cysteine (Translabel; ICN) at 0.25 mCi/ml was included during the last 2 h of serum deprivation or the last 2 h of FBS stimulation. After preparation of cell extracts, GFP and CAT proteins were immunoprecipitated with a mixture of anti-GFP and anti-CAT antibodies, which were then resolved on a SDS-12% polyacrylamide gel. After electrophoresis, the gel was stained, fixed, incubated in 1 M sodium salicylate (pH 6.0) for 30 min, dried, and exposed to X-ray film. The ratio of GFP to CAT was determined by autoradiography and quantification of 35S bands on a Bio-Rad phosphorimager by using ImageQuant software.
To determine DNA content, a Becton Dickinson FACSCalibur flow cytometer with CellQuest software was used. U2OS cells were seeded to 60-mm dishes at 1.5 × 105 cells/plate and transfected in triplicate the next day with 1 μg of CD20 and 10 μg of total plasmid to be assayed. After overnight transfection, followed by serum deprivation and stimulation (as described above), cells were harvested for analysis by flow cytometry as described earlier (13). Briefly, cells were removed from the plates with PBS-EDTA, incubated with anti-CD20-FITC antibodies, fixed in ethanol (80% final), and then incubated in propidium iodide-RNase A. To determine the DNA content of the untransfected cell population, 10,000 propidium iodide+ FITC− cells were collected. To determine the DNA content of the transfected cell population, 3,000 to 5,000 propidium iodide+ FITC+ cells were collected. Single cells were gated away from clumped cells by using an FL2 width versus FL-2 area dot plot. To quantitatively measure the percentage of cells in the various phases of the cell cycle, the marker tool within CellQuest was used to specifically gate on the G0/G1-, S-, and G2/M-phase peaks within an FL2 area histogram. All cell cycle data represent the means ± the standard deviations of triplicate transfections unless otherwise noted.
C2C12 myoblasts were cultured in DMEM-20% FBS and 5% CO2. To generate stable cell lines expressing exogenous HA-tagged S6K1, cells were infected with an S6K1-expressing retrovirus or with a pMV7 control virus and selected in G418. Pools of G418-resistant colonies were collected. To measure the ability of these cell lines to proliferate in low-serum-containing media (i.e., media containing low levels of serum), 104 cells were seeded into triplicate 35-mm wells directly in DMEM-2% FBS and then incubated for 2, 4, 5, and 6 days in this medium. The cell number was determined by counting the cells with a hemocytometer.
Data are presented as the means ± the standard deviations. The statistical significance was determined by using the Student t test (paired two sample for means; two tails) by using Microsoft Excel. P values of <0.05 were considered significant.
While the mTOR inhibitor rapamycin is known to block G1-phase progression, the biochemical signaling pathways regulated by mTOR to control cell cycle progression are poorly defined. We thus set out to investigate the role(s) of the S6K1 and 4E-BP1/eIF4E pathways in cell cycle progression, which also mediate mTOR-dependent cell growth and cell size (13). We chose to perform our analysis in human osteosarcoma U2OS cells, since they are readily synchronized and amenable to transfection. After serum deprivation for 30 h to synchronize the cells in G0 phase, U2OS cells were stimulated with serum in the absence or presence of rapamycin for various times, and DNA content and/or cell cycle profiles were determined by flow cytometry of propidium iodide-stained cells (Fig. (Fig.1).1). This analysis demonstrated that U2OS cells began entering S phase at between 12 and 16 h of serum stimulation and that rapamycin delayed entry into S phase by about 8 h.
Although TOR was initially identified as a gene that, when mutated in budding yeast, conferred resistance to the antiproliferative action of rapamycin (18), the mTOR-dependent nature of the rapamycin-induced effects on cell cycle progression and cellular proliferation in mammalian cells has not been formally demonstrated. We thus assayed the ability of RR-mTOR to rescue rapamycin-inhibited cell cycle progression in U2OS cells. Cells were transiently cotransfected with the cell surface marker CD20 and cDNAs for various mTOR isoforms, serum starved, and stimulated with serum for 20 h in the absence or presence of rapamycin. We chose the 20-h time point for this analysis since there is a marked difference in the percentage of cells that have entered S phase in the absence or presence of drug at this time (Fig. (Fig.1).1). To determine DNA content, cells were stained with anti-CD20-FITC-conjugated antibodies and propidium iodide and then analyzed on a flow cytometer. As expected, expression of RR-mTOR rescued G0-to-S-phase cell cycle progression in the presence of rapamycin, whereas the double RR/KD mutant did not, indicating that inhibition of mTOR is the mechanism by which rapamycin delays cell cycle progression and that the kinase activity of mTOR is required for rescue (Fig. 2A and B). Importantly, only the FITC+ cells transfected with RR-mTOR (Fig. (Fig.2B)2B) (which represent a population enriched in transfected cells) and not the FITC− cells (Fig. (Fig.2C)2C) (which represent a population enriched in untransfected cells) displayed rescue of rapamycin-inhibited cell cycle progression, and all four mTOR constructs were expressed to similar levels (Fig. (Fig.2D).2D). Interestingly, although neither of the mTOR mutants containing the kinase-inactivating mutation (RR/KD or KD) inhibited G1-phase progression in a dominant-negative manner in the absence of rapamycin, both potentiated the inhibitory effect of rapamycin on S-phase entry.
To confirm that expression of RR-mTOR allows signaling to two of its major downstream targets in the presence of rapamycin in this experimental context, we cotransfected U2OS cells with various mTOR plasmids together with either HA-tagged S6K1 (Fig. (Fig.2E)2E) or HA-tagged 4E-BP1 (Fig. (Fig.2F).2F). As reported previously (4), RR-mTOR rescued the inhibitory effect of rapamycin on S6K1 activity, as assayed by mobility shift (anti-HA blot), anti-phospho-S6 immunoblotting, and in vitro kinase assay with recombinant GST-S6 as substrate (Fig. (Fig.2E).2E). Expression of KD-mTOR decreased S6K1 activity slightly in the absence of rapamycin compared to S6K1 activity in WT-mTOR-transfected cells. Since the inhibition of S6K1 activity by rapamycin is complete, no enhanced inhibition by KD-mTOR could be detected.
RR-mTOR also rescued the inhibitory effect of rapamycin on 4E-BP1 phosphorylation, as determined by anti-HA mobility shift (Fig. (Fig.2F),2F), a finding consistent with published results (5, 17). As observed for S6K1 inhibition, the expression of KD-mTOR constructs in the absence of rapamycin produced a dominant-negative effect and increased the intensity of the hypophosphorylated α-band, a finding consistent with decreased 4EBP1 phosphorylation (Fig. (Fig.2F,2F, short exposure). As for cell cycle progression, KD constructs also potentiated the inhibitory effect of rapamycin on 4EBP1 phosphorylation (Fig. (Fig.2F,2F, long exposure). The further decrease in 4EBP1 phosphorylation and cell cycle progression by KD-mTOR isoforms in the presence of rapamycin is consistent with some level of rapamycin-insensitive signaling by mTOR.
To link the S6K1 pathway to mTOR-dependent cell cycle progression, we sought to determine whether expression of RR mutants of S6K1 could rescue cell cycle progression in the presence of rapamycin. U2OS cells were transfected with a panel of S6K1 plasmids, serum deprived, stimulated in the absence or presence of rapamycin, and analyzed by flow cytometry to determine DNA content (Fig. (Fig.3A).3A). Rapamycin inhibited S-phase entry, as expected, whereas cells expressing two different RR-S6K1 constructs (E389D3E and E389ΔCT) displayed modest but significantly enhanced S-phase entry in the presence of rapamycin compared to the vector control and WT- or KD-S6K1-transfected cells (see Fig. Fig.3A3A for P values). Furthermore, cells expressing the two partially rapamycin-resistant mutants of S6K1 (E389D3E and E389ΔCT) also exhibited modest but significant acceleration of S-phase entry in the absence of rapamycin compared to the vector control and WT- or KD-S6K1-transfected cells (Fig. (Fig.3A).3A). When DNA content was determined after serum deprivation, cells expressing the RR-S6K1 constructs displayed cell cycle profiles similar to pRK7- or WT-S6K1-transfected cells, indicating that S-phase entry in cells expressing the RR-S6K1s is not accelerated due to reduced quiescence in G0 phase (Fig. (Fig.3B).3B). The rescue of rapamycin-inhibited G1-phase progression and the acceleration of G1-phase progression in the absence of drug upon expression of RR-S6K1 constructs demonstrates that the S6K1 pathway controls G1-phase cell cycle progression downstream of mTOR. The inability of the RR-S6K1 constructs to completely rescue rapamycin-inhibited cell cycle progression is likely due to the limited resistance of these mutants to rapamycin in U2OS cells (Fig. (Fig.3C)3C) combined with the contribution of other mTOR-dependent pathways to cell cycle control.
Since the activity of the RR-S6K1 mutants was similar to WT-S6K1 when they were assayed after 20 h of serum stimulation, as was the extent of phosphorylation of ribosomal protein S6 (Fig. (Fig.3C),3C), it was interesting that only the RR-S6K1 mutants accelerated serum-stimulated S-phase entry, whereas WT-S6K1 did not. It is important to note that such mutants are known to possess elevated basal kinase activity. We therefore determined whether these mutants (E389D3E and E389ΔCT) exhibit elevated kinase activity after serum deprivation in our cell system (Fig. (Fig.3D).3D). As expected, the RR-S6K1 mutants possessed increased kinase activity (~2-fold) compared to WT-S6K1 after serum deprivation. These data suggest that cells expressing these “constitutively active” mutants of S6K1 mutants accelerate G1-phase progression because they have increased kinase activity at t = 0 and, as a result, are able to progress through G1 phase and enter S phase at a faster rate. We also noted that, although overexpression of WT-S6K1 was not sufficient to drive G1-phase progression, it was sufficient to increase the phosphorylation of endogenous ribosomal protein S6 compared to pRK7 vector control in the absence of serum. Also, whereas E389D3E and E389ΔCT accelerated S-phase entry while WT-S6K1 did not, all three S6K1 isoforms increased the phosphorylation state of ribosomal protein S6 to a similar extent over controls after serum deprivation (Fig. (Fig.3D),3D), suggesting that perhaps S6K1-dependent phosphorylation of ribosomal protein S6 may not be the critical downstream target of S6K1 for cell cycle control.
The data presented above indicate that increased S6K1-dependent signaling is sufficient to accelerate G1-phase progression downstream of mTOR. To determine whether S6K1-dependent signaling is required for G1-phase progression, we reduced the expression of S6K1 by using a plasmid-based RNAi approach (52). A hairpin DNA sequence targeting human S6K1 was subcloned into the RNAi vector pBS/U6 to create pBS/U6-S6K1. We first tested whether this hairpin RNAi plasmid could knock down the expression of transfected, exogenous S6K1. To do this, HA-tagged rat S6K1 was cotransfected with pBS/U6 or pBS/U6-S6K1 at various ratios (Fig. (Fig.4A;4A; human and rat S6K1 are identical in sequence in the targeted region). A 100:1 ratio of RNAi plasmid to transfected HA-S6K1 resulted in almost complete elimination of exogenous S6K1 expression. To confirm that the hairpin RNAi plasmid could also knock down the expression of endogenous S6K1, we performed immunofluorescence with anti-phospho-S6 antibodies as an indirect measure of endogenous S6K1 activity (we have not been able to identify anti-S6K1 antibodies that specifically detect endogenous S6K1 by immunofluorescence). To identify transfected cells, CD20 and either pBS/U6 or pBS/U6-S6K1 were cotransfected, and the cells were stained with anti-CD20 and anti-phospho-S6 antibodies (Fig. (Fig.4B).4B). Cells transfected with the hairpin RNAi plasmid targeting S6K1 exhibited decreased anti-phospho-S6 staining, whereas those transfected with vector control displayed no reduction in anti-phospho-S6 signal. Therefore, the hairpin RNAi plasmid against S6K1 is able to reduce the expression of exogenous S6K1 and reduce the activity of endogenous S6K1. To determine whether S6K1 activity is required for G1-phase cell cycle progression, either vector control or RNAi-S6K1 plasmids were cotransfected with CD20, and the percentage of cells in S-phase after serum deprivation (−FBS) or after serum stimulation (+FBS) was determined (Fig. (Fig.4C).4C). Expression of the RNAi-S6K1 plasmid partially inhibited serum-stimulated S-phase entry, indicating that S6K1 is required for G1-phase progression. The inability of RNAi-mediated reduction of S6K1 expression to completely block S-phase entry suggests that additional mTOR-dependent signaling pathways also contribute to cell cycle control.
In order to link the 4E-BP1/eIF4E pathway to mTOR-dependent cell cycle progression, we determined whether overexpression of eIF4E could drive cell cycle progression in the presence of rapamycin. Since overexpression of eIF4E would be expected to overwhelm endogenous 4E-BP1 and render eIF4E constitutively free even in the presence of hypophosphorylated 4E-BP1, overexpression of WT eIF4E seemed to be a reasonable approach to augment eIF4E function. Cells were transfected with pMV7 vector or eIF4E, deprived of serum, stimulated in the absence or presence of rapamycin, and analyzed by flow cytometry (Fig. (Fig.5A).5A). Rapamycin inhibited S-phase entry of vector-transfected cells, as expected. Similar to cells expressing RR-S6K1, cells overexpressing eIF4E displayed a small but significant acceleration of S-phase entry in the presence of rapamycin compared to the vector control. As observed in cells expressing constitutively active mutants of S6K1 (Fig. (Fig.3A),3A), cells overexpressing eIF4E also showed modest but significant acceleration of G1-phase progression from G0 phase to S phase in the absence of rapamycin. Importantly, the eIF4E-expressing cells quiesced similarly to vector control (−FBS) (Fig. (Fig.5A).5A). Therefore, the eIF4E pathway mediates a component of mTOR-dependent cell cycle control, similar to what was observed with S6K1.
eIF4E overexpression has been reported to activate a negative-feedback loop that results in dephosphorylation of 4E-BP1 and S6K1 (24), a finding that conflicts with our observation that eIF4E accelerates cell cycle progression. On the other hand, eIF4E overexpression has also been reported to transform rodent fibroblasts (27), which is consistent with our results. We therefore sought to determine whether expression level of exogenous eIF4E could account for this discrepancy by highly overexpressing eIF4E from a strong cytomegalovirus (CMV) promoter (vector pRK7) and determining its effect on cell cycle progression. eIF4E expressed from pRK7 expresses to much higher levels than when expressed from the weak retroviral long terminal repeat promoter (vector pMV7), and significantly more eIF4E binds to m7GTP beads when expressed from pRK7 than when expressed from pMV7 (Fig. (Fig.5B).5B). Importantly, in our cell system, when vastly overexpressed from pRK7, eIF4E inhibits G1-phase progression (Fig. (Fig.5C),5C), a finding consistent with activation of the previously reported negative feedback loop (24).
In order to eliminate the possibility that the eIF4E phenotypes observed above were the result of an mTOR-independent mechanism, we investigated the role of 4E-BP1 in cell cycle progression. We took advantage of two different mTOR-insensitive mutants of 4E-BP1 that dominantly bind to and constitutively inhibit eIF4E and therefore cap-dependent translation. AA-4E-BP1 contains alanine substitution mutations at threonines 37 and 46, which are mTOR-dependent priming phosphorylation sites, and thus cannot be phosphorylated by mTOR (reviewed in references 15 and 29). F114A-4E-BP1 contains a mutation in the TOS motif and does not associate with the mTOR/raptor complex; consequently, F114A-4E-BP1 cannot be phosphorylated (8, 33, 45, 59). After serum stimulation, expression of WT-4EBP1 exhibited very little inhibitory effect on S-phase entry, whereas AA-4EBP1 significantly inhibited S-phase entry (Fig. (Fig.6A).6A). Importantly, this effect on S-phase entry was specific for eIF4E since the AA-Y54A mutant of 4EBP1, which cannot bind eIF4E, failed to inhibit S-phase entry. Overexpression of F114A-4E-BP1 inhibited G0-to-S-phase progression to a similar extent as the phosphorylation site mutant AA-4E-BP1 (Fig. (Fig.6B).6B). These results show that the effects of eIF4E on cell cycle progression described above are mTOR-dependent and demonstrate that mTOR-dependent phosphorylation of 4E-BP1 and mTOR/raptor association with 4E-BP1 is required for mTOR-dependent G1-phase progression.
To confirm that these 4E-BP1 constructs behave as expected in our cell system, we assayed the ability of the various 4E-BP1 constructs to bind eIF4E in the absence (−FBS) or presence (+FBS) of serum by m 7 GTP Sepharose affinity chromatography (Fig. (Fig.6C).6C). In this assay, m7GTP, which mimics the cap structure at the 5′ ends of mRNA transcripts, is coupled to Sepharose beads and therefore readily purifies eIF4E and eIF4E-associated proteins, such as 4E-BP1, from cell lysate. As expected, WT-4E-BP1 dissociated from eIF4E upon serum stimulation, whereas AA-4E-BP1 remained constitutively bound to eIF4E and did so at higher levels than WT-4E-BP1. The mutant of 4E-BP1 that lacked the eIF4E-binding motif (Y54A) failed to associate with eIF4E. To further confirm that these mutants of 4E-BP1 produce effects on translation, we cotransfected the various 4E-BP1 constructs with a bicistronic vector containing both a GFP reporter gene downstream from the β-globin 5′-UTR, which monitors cap-dependent translation in vivo, and a CAT reporter gene downstream from the encephalomyocarditis virus internal ribosome entry site (IRES), which monitors cap-independent translation in vivo (Fig. (Fig.6D)6D) (55). As expected, overexpression of WT-4E-BP1 more strongly repressed cap-dependent translation in the absence of serum than in its presence, and rapamycin treatment of serum-stimulated cells expressing WT-4E-BP1 repressed cap-dependent translation. Compared to vector- or WT-4E-BP1-transfected cells, overexpression of AA-4E-BP1 dominantly repressed cap-dependent translation in both the absence or presence of serum, and binding to eIF4E was required for AA-4E-BP1 to repress cap-dependent translation. Therefore, in our cell system, the phosphorylation site mutant AA-4E-BP1 dominantly inhibits cap-dependent translation through its binding to eIF4E, as expected.
To determine whether mTOR signals independently along both the S6K1 and 4EBP1/eIF4E pathways to control G1-phase progression, cells were transfected with the AA-4E-BP1 or RNAi-S6K1 plasmids individually or together, and the ability of the cells to enter S-phase after serum stimulation was assayed. Figure Figure6E6E clearly shows that although downregulation of each pathway individually inhibited S-phase entry, simultaneous downregulation of both pathways more strongly inhibited S-phase entry to an extent approaching that of rapamycin treatment. Since downregulation of both the eIF4E and S6K1 pathways did not completely mimic rapamycin-mediated inhibition of G1-phase progression, it is possible that either there exist other mTOR-dependent pathways yet to be identified that contribute to mTOR-dependent cell cycle control or that we have not completely downregulated these pathways. Importantly, coexpression of the dominant AA-4E-BP1 mutant with RR-mTOR blocked the ability of RR-mTOR to restore G1 progression in the presence of rapamycin, a finding consistent with the idea that 4E-BP1-dependent cell cycle control is me-diated by mTOR (data not shown).
Lastly, we determined whether overexpression of WT-S6K1 could confer a proliferative advantage in low-serum-containing media, a hallmark of transformation. We therefore generated pools of C2C12 myoblasts stably expressing pMV7 vector control or HA-tagged S6K1. We chose C2C12 cells for two reasons: C2C12 proliferation is particularly rapamycin sensitive, sug-gesting that the S6K1 pathway may be important for proliferation control in this cell line, and we were unable to generate U2OS lines that stably overexpressed S6K1. In both full-serum-containing media (20% FBS) and low-serum-containing media (2% FBS), total S6K activity was increased ~3-fold in S6K1-expressing pools compared to parental cells or pMV7-expressing pools (Fig. (Fig.7A).7A). Surprisingly, total S6K1 activity in 2% FBS was increased ~3-fold across the board relative to total S6K1 activity in 20% FBS, suggesting that low serum activates a positive feedback loop in C2C12 cells that ultimately increases S6K1 activity. In 20% FBS, the extent of phosphorylation of ribosomal protein S6 correlated well with the amount of S6K1 activity; in 2% FBS, however, conditions in which overall S6K1 activity is higher, there was no difference in the extent of phosphorylation of ribosomal protein S6 among the various cell lines, most likely due to the fact that this high level of S6K1 activity is sufficient to fully phosphorylate ribosomal protein S6. After characterizing the stable cell lines, we assayed their ability to proliferate over 6 days in low-serum-containing media (2% FBS). Overexpression of S6K1 clearly conferred a proliferative advantage in low-serum-containing media compared to parental cells or the pMV7 control line; no differences in rates of proliferation were observed in full-serum-containing media (20% FBS) (data not shown) (Fig. (Fig.7B).7B). Interestingly, whereas overexpression of WT-S6K1 allowed cells to proliferate in low serum better than controls, this cell line did not exhibit greater S6 phosphorylation. This result may indicate that, similar to what we observed in Fig. Fig.3,3, phosphorylation of ribosomal protein S6 may not be the critical downstream target of S6K1 that is important for S6K1-dependent cell cycle control. It is important to note that the ability of S6K1 overexpression to drive proliferation in low serum likely does not result from an inhibition of C2C12 myoblast differentiation, which is normally induced by serum withdrawal. When C2C12 cells are cultured in 2% FBS, they still proliferate, albeit slowly, and do not show any morphological changes characteristic of muscle differentiation. Furthermore, the activity of S6K1 increases during C2C12 differentiation, although it is not required (12) and, indeed, our data (Fig. (Fig.6A)6A) clearly confirms that S6K1 is more active in low serum (2% FBS) than in high serum (20% FBS). Therefore, it is unlikely that S6K1 enhances apparent proliferation rate by inhibiting differentiation, and thus the observed proliferation effect is likely to be direct.
Although mTOR has been known to control G1-phase progression, the roles of specific mTOR-dependent pathways in cell cycle control have remained poorly defined. We demonstrate that restoration of mTOR function by using rapamycin-resistant mutant rescues rapamycin-inhibited G1-phase progression, as in lower eukaryotes (18). This result is consistent with the reported requirement for mTOR in G1-phase progression; microinjection of the isolated FKBP12-rapamycin-binding (FRB) domain of mTOR inhibited serum-stimulated S-phase entry, presumably by functioning as a dominant-negative (57). Our data furthermore define roles for the mTOR-regulated S6K1 and 4E-BP1/eIF4E pathways in the control of G1-phase progression by mTOR: activation of either the S6K1 or 4E-BP1/eIF4E pathways partially rescues the inhibitory effect of rapamycin on G0-to-S-phase cell cycle progression and modestly accelerates cell cycle progression in the absence of drug. mTOR-dependent phosphorylation of 4E-BP1 is required for mTOR's ability to drive the cell cycle, as is the ability of the mTOR/raptor complex to associate with 4E-BP1 via the TOS motif. Both the S6K1 and the 4E-BP1/eIF4E pathways independently mediate mTOR-dependent cell cycle control in parallel, as simultaneous downregulation of these pathways (using RNAi against S6K1 and overexpression of the phosphorylation site mutant AA-4E-BP1) additively inhibits G1-phase progression compared to downregulation of the pathways individually.
Our finding that RR-S6K1 partially rescues rapamycin-inhibited G1-phase progression is consistent with previous reports in which RR-S6K1 could partially restore rapamycin-suppressed E2F transcriptional responses in Kit225 cells, a human T-cell line (3), and could partially rescue rapamycin-inhibited proliferation of vascular endothelial cells, as assayed by measuring [3H]thymidine incorporation (58). Importantly, we demonstrate that overexpression of S6K1 accelerates serum-stimulated G1-phase progression and increases the proliferation rate in low serum, suggesting that the activity of S6K1 is limiting for G1-phase progression. Since the reduction of S6K1 expression by using RNAi inhibited but did not completely block G1-phase progression, our data suggest that, although S6K1 participates in cell cycle control, it is not absolutely required for serum-stimulated G1-phase progression. This result differs from a previous report in which microinjection of anti-S6K1 antibodies blocked serum-stimulated S-phase entry, as well as total protein synthesis and c-Fos induction (26). Although it is possible that the more modest requirement for S6K1 that we demonstrate here results from our inability to completely inhibit S6K1 signaling with RNAi or results from cell type-specific differences, our data demonstrate complete RNAi-mediated reduction of overexpressed S6K1 and phosphorylation of endogenous ribosomal protein S6. Consistent with our data, deletion of S6K1 in embryonic stem cells reduces, but does not block, the capacity of the cells to proliferate in culture, and rapamycin treatment further reduces proliferation rate of these cells (23). Furthermore, rapamycin completely inhibits S6K1 and S6K2 activity, yet only modestly reduces total protein synthesis, does not block serum-stimulated c-Fos induction, and only delays G1-phase progression. Thus, the formation of intracellular S6K1-containing immune complexes formed by antibody microinjection may disrupt cellular functions differently than the mere absence of S6K1 by RNAi. Collectively, the data suggest that, although it is not absolutely required, S6K1 has a positive influence on cell cycle progression and proliferation and that other mTOR-dependent signaling pathways likely contribute to this as well.
That reduction of S6K1 expression does not induce cell cycle arrest is consistent with our finding that the 4E-BP1/eIF4E pathway operates in parallel to S6K1 downstream of mTOR to control cell cycle progression. We report negative and positive roles for 4E-BP1 and eIF4E, respectively, in control of mTOR-regulated G1-phase progression in mammalian cells. These data are consistent with transformation of rodent fibroblasts by overexpression of eIF4E (27), which is blocked by cooverexpression of 4E-BP1 (41), and consistent with the increased cell division time caused by antisense RNA-mediated reduction in eIF4E expression (10). Our finding that overexpression of eIF4E accelerates G1-phase progression appears at first to conflict with a report that eIF4E overexpression activates a negative-feedback loop, resulting in the dephosphorylation of S6K1 and 4E-BP1 (24). It is important to note that eIF4E accelerates the G1 phase in our cell system when it is expressed at low levels from the weak retroviral long terminal repeat promoter; when eIF4E is expressed to much higher levels with a strong CMV promoter, however, G1-phase progression is inhibited. Therefore, expression level likely determines whether eIF4E activates a negative feedback loop to restrict aberrant cell cycle progression and proliferation. Since simultaneous downregulation of both the S6K1 and 4E-BP1/eIF4E pathways inhibits G1-phase progression to an extent approaching that of rapamycin, it is likely that these represent the major pathways mediating mTOR-dependent cell cycle control.
We also consistently noted that the combination of KD mTOR plus rapamycin more strongly inhibits 4E-BP1 phosphorylation, G1-phase progression, and cell size (13) compared to WT-mTOR plus rapamycin. The trivial explanation for these phenomena is that although rapamycin blocks all mTOR signals, it does so incompletely. A more intriguing possibility is that mTOR also possesses rapamycin-insensitive kinase-dependent functions that, although obviously not inhibited by rapamycin, could be blocked with kinase-inactive and dominant-negative mTOR. In this case, inhibition of the rapamycin-sensitive functions of mTOR by treatment with rapamycin would unmask these rapamycin-insensitive functions of the mTOR kinase. Indeed, the observation that rapamycin completely inhibits S6K1 and endogenous ribosomal prtotein S6 phosphorylation suggests that the blockade of at least some mTOR signals by rapamycin is complete. Furthermore, TOR2 in budding yeast mediates both rapamycin-sensitive and rapamycin-insensitive signals, and one of the proteins (AVO1) found in the rapamycin-insensitive TOR2 complex (TORC2) has a mammalian orthologue of unknown function, mAVO1/hSIN1 (28). Therefore, although the possibility that mTOR may signal in a rapamycin-insensitive manner needs to be investigated much more carefully, there is precedent for the idea.
We reported previously that mTOR controls cell size, which is mediated, at least in part, by the S6K1 and 4E-BP1/eIF4E signaling pathways (13). Since both cell size and cell cycle progression are controlled by mTOR and by the same mTOR-dependent signaling pathways, this nutrient- and mitogen-responsive signaling molecule is centrally positioned to couple cell growth with cell division (Fig. (Fig.8A).8A). These data, together with the known dependence of cell cycle progression on a sufficient level of cell growth (22; for a review, see reference 36), suggest a model in which mTOR primarily drives cell growth (i.e., macromolecular biosynthesis), and as a secondary consequence promotes cell cycle progression (Fig. (Fig.8B,8B, model 1). Although we favor this model, our data do not exclude the possibility that mTOR controls cell cycle progression via a cell growth-independent mechanism (Fig. (Fig.8B8B model 2); for example, mTOR-dependent signaling could directly regulate components of the cell cycle machinery via a fast-acting phosphorylation cascade. Indeed, the phosphatidylinositol 3-kinase/Akt pathway, which coordinates growth factor signaling with mTOR signals, has been reported to have direct effects on the cell cycle machinery (11, 40, 63).
Our data also show that the increased cell size produced by S6K1 or eIF4E overexpression (13) results from augmented cell growth and not from delayed cell cycle progression. Conversely, the decreased cell size observed upon 4E-BP1 overexpression results from decreased cell growth and not from accelerated cell cycle progression. We make this point because cell size phenotypes can result from changes in either cell growth rate or cell cycle progression rate (e.g., when the cell cycle is blocked, cells grow to increased cell size; when the cell cycle accelerates in the face of an unchanged rate of cell growth, cells become smaller). Therefore, whenever a cell size phenotype is observed, it is important to determine whether it results from altered cell growth or altered rate of cell cycle progression.
If mTOR-regulated cell growth influences the rate of cell cycle progression, it seems reasonable to speculate that, since the S6K1 and 4E-BP1/eIF4E pathways control translation, increased expression of critical cell cycle regulatory proteins could represent a mechanism by which cell cycle progession is coupled to cell growth. Indeed, the synthesis of the G1-cyclin CLN3 in budding yeast is controlled by TOR and Cdc33 (an eIF4E orthologue) at the level of translation initiation, effectively coupling the synthesis of a cell cycle regulator to protein biosynthetic rate (2). A short upstream open reading frame in the 5′ leader of the CLN3 transcript functions as a translational control element to repress CLN3 expression when protein synthesis and cell growth rate are low (38). Since eIF4E has also been reported to regulate nucleocytoplasmic transport of mRNA transcripts (reviewed in reference 51), it is also possible that eIF4E drives cell growth and cell cycle progression by increasing the nucleocytoplasmic export rate of mRNAs encoding specific cell cycle regulators instead of or in addition to controlling their translation. Although the identity of the critical eIF4E-controlled cell cycle regulator(s) is not known at this time, eIF4E has been shown to increase the translation of ODC and c-Myc and to increase the expression of cyclin D1 through increased nucleocytoplasmic transport (42; reviewed in reference 48).
The mechanism by which S6K1 drives cell growth and the cell cycle is also unclear. Of course, S6K1-dependent phosphorylation of ribosomal protein S6 may drive ribosome biogenesis and therefore increase protein biosynthetic capacity, resulting in augmented cell growth and promotion of cell cycle progression. Although S6K1 clearly controls cell growth and cell cycle progression, the critical downstream targets responsible for mediating this response are not clearly defined. In two different experiments with two different cell types, we noted that the degree of ribosomal protein S6 phosphorylation was not concordant with the cell cycle phenotype produced by S6K1 overexpression. For example, the S6K1 mutants E389D3E and E389ΔCT accelerated G1-phase progression, whereas WT-S6K1 did not, and yet overexpression of all three S6K1 constructs increased the phosphorylation of ribosomal protein S6 over vector control. In addition, S6K1 overexpression increased proliferation rate in low serum compared to parental or control cells, and yet in low serum ribosomal protein S6 was phosphorylated to similar levels in all three cell lines. The observation that S6K1-regulated G1-phase progression and ribosomal protein S6 phosphorylation can be dissociated suggests the intriguing possibility that S6K1 likely has other targets besides the ribosomal protein S6. Consistent with this idea, we have recently identified a novel S6K1-interacting protein and in vivo substrate, SKAR, which bears homology to the ALY/REF family of proteins that couple transcription, splicing, and RNA export (C. Richardson et al., unpublished data). SKAR also controls cell growth or cell size and also regulates the rate of cell cycle progression (Richardson et al., unpublished), suggesting that S6K1 may control cell growth and cell cycle progression through an RNA processing event (Fig. (Fig.8A).8A). Another in vivo target of S6K1 is the cap-binding protein CBP80, which functions in an early step of RNA splicing (60). Therefore, it is possible that S6K1-regulated cell cycle progression may be due in part to regulation of RNA processing (Fig. (Fig.8A).8A). Alternatively, S6K1 could regulate the cell cycle by a cell growth-independent mechanism via phosphorylation of unidentified targets that would directly regulate the cell cycle machinery (Fig. (Fig.8B,8B, model 2).
In the present study, we identify the S6K1 and 4E-BP1/eIF4E pathways as the major mTOR-dependent downstream signaling pathways that mediate mTOR-regulated G1-phase progression (Fig. (Fig.8A).8A). Since these pathways also mediate mTOR-dependent control of cell growth or cell size (13), mTOR is uniquely positioned to function as a central coordinator of cell growth and cell division. In the future, we hope to better understand the relationship between mTOR-regulated cell growth and cell division and to move further downstream to determine how mTOR couples these fundamental biological processes.
We thank Robert Abraham for his generous donation of mTOR plasmids, Nahum Sonenberg for generously sharing the eIF4E and 4E-BP1 plasmids, Morris Birnbaum for generously providing the anti-phospo-S6 antibodies, and A. A. M. Thomas for sharing the pGFP/CAT plasmid. We are grateful to Martin G. Myers, Jr., for critical reading of the manuscript and members of the Blenis lab for help in general, particularly Stefanie Schalm for generating the pACTAG2/3HA-AA-Y54A-4E-BP1 mutant. We especially thank Jennifer Waters-Shuler and the Nikon Imaging Center at Harvard Medical School for help with immunofluorescence image training and acquisition.
We thank the Tuberous Sclerosis (TS) Alliance and the Rothberg Courage Fund. This study was supported by an NIH award GM51405 to J.B. D.C.F. was supported by NIH NRSA fellowship F32 CA69808 and by the TS Alliance, C.R. was supported by NCI grant T32 CA09361, and A.T. was supported by ALT2002-463.