In recent years, the intracellular oxidation–reduction (redox) state has gained increasing attention as a critical mediator of cell signaling, gene expression changes and proliferation. This review discusses the evidence for a redox cycle (i.e., fluctuation in the cellular redox state) regulating the cell cycle. The presence of redox-sensitive motifs (cysteine residues, metal co-factors in kinases and phosphatases) in several cell cycle regulatory proteins indicate periodic oscillations in intracellular redox state could play a central role in regulating progression from G0/G1 to S to G2 and M cell cycle phases. Fluctuations in the intracellular redox state during cell cycle progression could represent a fundamental mechanism linking oxidative metabolic processes to cell cycle regulatory processes. Proliferative disorders are central to a variety of human pathophysiological conditions thought to involve oxidative stress. Therefore, a more complete understanding of redox control of the cell cycle could provide a biochemical rationale for manipulating aberrant cell proliferation.
Cellular proliferation encompasses tightly regulated biochemical and genetic pathways, the loss of which can lead to aberrant proliferation. In recent years, the role of the intracellular redox state as a growth regulator is increasingly becoming appreciated (Burdon et al., 1989; Burdon and Rice-Evans, 1989). Cellular redox state is a delicate balance between the levels of reactive oxygen species (ROS) produced during metabolism and the antioxidant system that scavenges them. In general, ROS contain one or more unpaired electrons in their highest occupied orbital. The partial reduction of molecular oxygen results in the production of superoxide (O2•−) and hydrogen peroxide (H2O2) (Halliwell and Gutteridge, 1999). O2•− and H2O2 react with transition metal ions (e.g. cuprous and ferrous ions) through Fenton and Haber–Weiss chemistry, promoting further radical generation, including the highly reactive hydroxyl radical (•OH) (Halliwell and Gutteridge, 1992). Cells can generate ROS from exogenous sources as well as endogenously. Endogenously, ROS are largely produced as oxidative by-products of cellular metabolism; mitochondria, accounting for approximately 1–2% of molecular oxygen consumed, are the major site of O2•− production. Other endogenous sources of ROS include the superoxide-generating nicotinamide adenine dinucleotide phosphate (reduced form) (NADPH) oxidase complex, peroxisomes that generate H2O2, metabolism of fatty acid chains, cytochrome P450 reductase, xanthine oxidase, myeloperoxidase and nitric oxide synthase (Massey et al., 1969; Bredt et al., 1991; Bokoch and Knaus, 2003; Schrader and Fahimi, 2004; Zangar et al., 2004; Klebanoff, 2005). Additionally, recent evidence suggests interaction between specific receptor–ligands generates H2O2 (DeYulia et al., 2005). ROS, diverse and abundant in biological systems, were traditionally thought of as toxic products leading to damage of cellular components, including nucleic acids, proteins and lipids resulting in cell death (apoptosis and necrosis). However, recent evidence suggests ROS could be beneficial and function as signaling molecules regulating proliferation and growth arrest (Rhee, 1999). This dual function of ROS (signaling molecules and toxins) could be due to the differences in their concentrations, pulse duration and sub–cellular localization. Such a threshold concept of ROS regulating multiple biological processes is evident from a recent report of H2O2 regulating both proliferation and cell death (Laurent et al., 2005). NIH3T3 fibroblasts treated with 0.02–0.13 μ M H2O2 enhanced proliferation, whereas treatment with 0.25–2 μ M H2O2 or compounds (buthionine-(S, R)-sulfoximine (BSO), superoxide dismutases (SOD) mimics) known to increase intracellular H2O2 levels resulted in growth arrest and cell death. Therefore, while higher levels of ROS could be toxic, low levels of ROS are necessary for proper functioning of cellular processes including proliferation.
Cells protect themselves against the reactive and damaging effects of ROS by expressing antioxidant enzymes like SOD, catalase (CAT), glutathione peroxidase (GPx) and other small molecular ROS scavengers. Of the enzymatic antioxidants, SOD converts O2•− into H2O2 (McCord and Fridovich, 1969). There are three forms of SOD: CuZnSOD (SOD1), the cytoplasmic and nuclear form; MnSOD (SOD2), the mitochondrial form (Fridovich, 1989); and extracellular SOD (EcSOD, SOD3), found in the plasma membrane (Folz and Crapo, 1994). CAT specifically reduces H2O2, whereas GPx reduces both H2O2 and organic hydroperoxides, utilizing reduced glutathione (GSH) as an electron donor. While CAT is localized primarily in the peroxisomes and cytoplasm, different isozymes of GPx are found in most sub–cellular compartments (Yamamoto et al., 1988; Oberley et al., 1990). Other cellular peroxidase scavenging enzyme systems include thioredoxin, glutaredoxins and the six-member family of peroxiredoxins (Rhee et al., 1999; Tanaka et al., 2000; Fernandes and Holmgren, 2004). Cysteine, GSH, vitamins C and E (ascorbic acid and α-tocopherol), among others, constitute a pool of small molecular weight non-enzymatic antioxidants (Di Mascio et al., 1991). GSH is ubiquitously found within the cell in both reduced (GSH) and oxidized (GSSG) forms (Schafer and Buettner, 2001). GSSG can be reduced back to GSH by glutathione reductase, at the cost of NADPH, and also by the thioredoxin or glutaredoxin systems. For this reason, changes in the GSH/GSSG ratio are routinely used as indicators of perturbation in the intracellular redox state.
Intracellular redox state and cell cycle progression
At the heart of every organism's development and growth process is its ability to divide. In normal cells, growth and division is a highly ordered, regulated series of events. When signaled by various mitogenic stimuli (growth factors), cells give rise to two daughter cells by transiting through the different cell cycle phases (G0 to G1 to S to G2 to M). In the S phase, the cellular DNA is duplicated, in the following gap phase (G2), cells prepare for mitosis and, upon entering the M phase, the DNA gets segregated and the cell divides into two. G1, however, is the critical phase of the cycle; it determines if cells continue in the cycle or if proliferation is arrested. Temin (1971) first proposed the presence of a G1-phase decision point, after which cells became independent of mitogenic signaling. Later, Pardee (1974) named this point the ‘restriction point’ and defined it as the point after which a cell is committed to enter the S phase, more or less independent of external conditions.
The first evidence of a redox cycle within the cell cycle came in 1931 from Rapkine (1931), who demonstrated the presence of soluble thiols that fluctuated cyclically. Using sea urchin eggs, he observed levels of soluble thiols decreased until metaphase and then increased toward mitosis. Later, in 1960, Kawamura et al. also used sea urchin eggs to show increased protein thiol (–SH) staining in prophase as the mitotic spindle was assembling. This SH staining remained high as cells moved into metaphase, gradually decreased in anaphase and was almost undetectable in telophase (Kawamura, 1960). Using synchronous HeLa cells, Mauro et al. (1969) further dissected the changes occurring in protein-bound and non-protein sulfhydryl (–SH) and disulfide (–SS–) groups in each phase of the cell cycle. The authors in this study observed the concentration of non-protein –SH dropped 10-fold from early G1 to late G1 and then rose almost 30-fold by the end of S phase. In contrast, the protein–bound –SH groups rose during G1 and dropped during the S phase (Mauro et al., 1969).
The concept of a redox cycle regulating the cell cycle is further supported by a recent report from Tu et al. (2005). Budding yeast exhibit a metabolic redox cycle consisting of a reductive non-respiratory phase and an oxidative respiratory phase. This metabolic redox cycle coordinates with periods of gene expression regulating essential cellular and metabolic events (Tu et al., 2005). Many of the genes regulating DNA replication and cell cycle progression are expressed during its reductive phase with cell cycle initiation occurring very late during the oxidative phase. Consistent with these results, we have shown previously that a transient increase in pro-oxidant levels early in G1 is required for the cells to transit from G1 into the S phase (Menon et al., 2003). Inhibiting this pro-oxidant event using an antioxidant like N-acetyl L-cysteine (NAC) arrests the cells in the G1 phase. Additionally, we found the fluorescence of a pro-oxidant sensitive dye was maximal in late S and G2+M phases of the HeLa cell cycle compared to G1 (Goswami et al., 2000). Additional reports from other laboratories have shown sub–lethal doses of ROS (O2•− and H2O2) added exogenously stimulated proliferation in cultured hamster fibroblasts (Burdon et al., 1989, 1990; Stirpe et al., 1991; Burdon and Gill, 1993). Interestingly, the effect of metabolic redox reactions on growth was not exclusive to mammalian cells, but can also be seen in Dictyostelium discoideum, one of the simplest organisms capable of forming multicellular structures. When conditions are favorable, D. discoideum survive as individual amoeba, but in response to starvation, the individual cells form spore–containing aggregates. During this process, O2•− generation is required for transition to a multicellular developmental phase. Furthermore, scavenging O2•− can prevent the formation of multicellular aggregates (Bloomfield and Pears, 2003). Thus, the use of O2•− as a signaling messenger by a lower eucaryote suggests this mode of signaling arose quite early during the evolutionary process and has been indispensable to every organism's growth and development.
The hypothesis of a redox cycle regulating the cell cycle is also supported by reports of antioxidant enzymes influencing cellular growth stage. MnSOD enzyme activity in NIH3T3 cells decreased during the S phase compared to G0. Cells possessing the ability of contact inhibition show higher SOD levels when proliferation halts; those not demonstrating density-dependent growth inhibition do not exhibit increased SOD levels (Oberley et al., 1995; Li and Oberley, 1998). Earlier, we have shown quiescent (G0) normal human fibroblasts cultured for a prolonged period of time lose the capacity to replicate DNA after sub–culture (Sarsour et al., 2005). However, G0 cells overexpressing MnSOD maintained their proliferative capacity, even after prolonged culture (Sarsour et al., 2005). MnSOD overexpression inhibited the age-associated increase in p16 (cyclin-dependent kinase inhibitor (CKI)) protein levels (Sarsour et al., 2005). It is believed this p16 inhibition protects the fibroblasts capacity to replicate DNA and divide (Sarsour et al., 2005). These results support the idea that O2•− is required for signal transduction in actively proliferating cells, whereas growth inhibition requires a reduction in mitogenic signaling, possibly by decreasing O2•− levels and increasing SOD activity. These observations not only underscore the importance of O2•− signaling but also highlight the importance of SOD regulation during growth. Furthermore, because mitochondria are the major source of cellular ROS (O2•−), and SOD dismutases O2•−, these results also support the hypothesis that mitochondrial oxidants could function as signaling molecules regulating cell cycle progression (Figure 1).
Modulation of other antioxidant enzymes has also been shown to affect proliferation rates. Catalase overexpression in vascular smooth muscle cells inhibits proliferation and increases the apoptotic rate, indicating H2O2 not only regulates proliferation but also survival (Brown et al., 1999). We have shown overexpression of GPx4 (PhGPx) inhibits progression from G1 to S, suggesting the possible involvement of lipid peroxidation-mediated signaling in cell cycle progression (Wang et al., 2003). In addition to antioxidant enzymes, cellular thiols also function as redox buffers and help maintain cellular redox potential. The major contributors to total thiol pools within the cell include the GSH/GSSG, thioredoxin/thioredoxin reductase and the cysteine/cystine couples, among others (Schafer and Buettner, 2001; Jones et al., 2004). Recent observations by Conour et al. (2004) indicate GSH content was significantly higher in the G2+M phase compared to G1, suggesting cells in the G2+M phase are at a more reduced state compared to the G1 phase. S-phase cells showed an intermediate redox state (Conour et al., 2004). Our laboratory, as well as others, has shown modulating intracellular redox state using NAC inhibits proliferation in mouse embryonic fibroblasts, hepatic stellate cells and vascular smooth muscle cells (Kim et al., 2001; Menon et al., 2003; Kyaw et al., 2004). These results further support the hypothesis that intracellular redox state is a critical regulator of cell cycle progression.
It has also been observed that altering extracellular redox environment changes proliferation rates. DNA synthesis, measured by the incorporation of 5-bromo-2-deoxyuridine, was at its minimum when extracellular redox was kept in an oxidizing state. In contrast, DNA synthesis significantly increased as the reduction potential (Eh) shifted to a more reducing state (Jonas et al., 2002). Hutter et al. (1997) measured changes in redox potential as cells progressed from an actively dividing to a contact inhibited growth state. The authors observed that actively dividing normal fibroblasts showed a reduction potential of −222 mV, this gradually increased and contact inhibited cell displayed a redox potential of −188 mV. Furthermore, to prove redox state fluctuations are the cause rather than the effect of proliferation, they used BSO (an inhibitor of γ-glutamylcysteine synthase) to inhibit GSH synthesis. Treating cells with BSO resulted in a less reduced state and decreased proliferation. Conversely, it has been shown that fibroblasts cultured sparsely in the presence of ROS scavengers show reduced protein tyrosine phosphorylation after epidermal growth factor (EGF) stimulation, mimicking the behavior of contact inhibited fibroblasts (Pani et al., 2000). Limoli et al. (2004) showed higher ROS levels in neuronal precursor cells when grown at low density compared to those grown at high density. Higher ROS levels were also associated with increased proliferation rates and metabolic activity, which could be inhibited with the antioxidant, α-lipoic acid (Limoli et al., 2004). These reports further support the concept that ROS could act like a dual-edged sword. Although proliferation occurs within a range of ROS levels, concentrations below or above this range result in growth arrest or cell death. The above results also suggest both the intra- (primarily mitochondria derived) and extracellular (receptor–ligand interaction) redox environments regulate cellular proliferation (Figure 1).
Intracellular redox state and cell cycle proteins
The influence of the cellular redox state on cell cycle regulatory proteins is not clearly understood. The tightly regulated progression through the cell cycle is brought about by periodic activation of cyclin and cyclin-dependent kinases (CDKs). The independent discoveries of cyclins in sea urchin oocytes, along with the simultaneous identification of maturation-promoting factors in frog oocytes and of CDC proteins in Saccharomyces cerevisiae, converged to give rise to the present–day concepts of cyclin–CDK complexes (Hartwell et al., 1970; Masui and Markert, 1971; Evans et al., 1983). The seminal discovery of cyclin/CDK complexes is believed to be at the core of understanding cellular proliferation controls. Cell cycle progression requires precise signal integration arising from mitogenic activation of transduction cascades. This is then followed by efficient transcription, activity of cell cycle phase-specific regulatory proteins and timely rapid degradation of critical cell cycle regulators. Thus, the redox control of cell cycle progression can act on any, or all, of the stages in this molecular network.
The generation of ligand-stimulated ROS plays an important role in mitogenic and growth factor signaling required for cellular progression through the cell cycle. Administering ROS activates a wide variety of proteins in various pathways; for example, platelet-derived growth factor (PDGF), EGF, mitogen–activated protein kinase (MAPK), protein kinase C (PKC) and protein kinase B (PKB/Akt) signal–transduction pathways. H2O2 generation during the activation of PDGF, transforming growth factor and EGF receptor signaling has also been well established (Sundaresan et al., 1995; Bae et al., 1997; Brar et al., 1999; Junn et al., 2000; Okuyama et al., 2001). The H2O2 generated results in tyrosine phosphorylation of the receptor, which subsequently activates MAPK (Gamou and Shimizu, 1995). The H2O2-dependent activation of the MAPK pathway can also activate antioxidant enzymes or other detoxifying genes like GCLC (glutamate-cysteine ligase, catalytic subunit) via the antioxidant response elements found in the promoter regions of these enzymes (Go et al., 2004). PKC enzyme activity can also be increased by exposure to O2•−, inducing disulfide bond formation and release of zinc from the cysteine-rich region of the enzyme (Knapp and Klann, 2000). In addition to the redox state altering kinase activity, regulation of protein tyrosine phosphatases and low molecular weight phosphotyrosine phosphatases by ROS has been extensively studied (Chiarugi et al., 2003). Recently, a bioinformatic approach to identify the redox-sensitive cell cycle proteins revealed almost 92 candidate proteins that could influence cellular progression in a redox-sensitive manner, of which only 24% have been identified experimentally (Conour et al., 2004). Of note, about 20% of the 92 proteins function in the G1 phase, only 10% in the S phase, but as much as 50% of the major redox motif-containing proteins were present in the G2+M phase (Conour et al., 2004).
Redox regulation of G1 and S cell cycle regulatory proteins
Following mitogenic stimulation, the first cyclins to be activated in G1 are the D-type cyclins (D1, D2 and D3) that associate with its CDK partner, CDK4 or CDK6 (Sherr, 1995). H2O2 exposures activate cyclin D1 accumulation in Her14 fibroblasts by inhibiting cyclin D1 protein degradation (Martinez Munoz et al., 2001). Degradation of cyclins by the ubiquitin–proteasome pathway is a key step in the regulation of cell cycle progression. The role of redox environment in regulating this pathway has also been reported. The activities of ubiquitin-activating enzyme (E1) and ubiquitin-conjugating enzyme (E2) were both inhibited with increased GSSG:GSH ratios (Jahngen-Hodge et al., 1997). In addition, it was shown that reversible S-thiolation of E1 and E2 enzymes regulates the ubiquitin-dependent proteolysis in response to altered redox environment (Obin et al., 1998). Furthermore, the 20S proteasome activity in S. cerevisiae was also reported to be inhibited by S-glutathionylation following H2O2 treatment (Demasi et al., 2003). These observations suggest redox regulation of the proteasome degradation machinery could regulate cyclin D1 levels.
Alternatively, redox modification of cyclin D1 itself could regulate its protein levels. Cyclin D1 contains two phosphorylation sites, on residues Thr286 and Thr288, which regulate its degradation. During late G1, cyclin D1 is proteasomally degraded following its phosphorylation at Thr286 by glycogen synthase kinase (GSK-3β) (Diehl et al., 1998). Degradation can also be mediated independent of GSK-3β via phosphorylation at the Thr288 residue by Mirk/dyrk kinase (Zou et al., 2004). We propose that redox regulation of cyclin D1 can occur through changes in redox state of critical cysteines, most likely the cysteine residue at position 285. Redox modification at cysteine 285 (–SH reduced to -S–S- oxidized state, or vice versa) could result in conformational change, which could then affect phosphorylation at Thr286 and/or -288 affecting its degradation. As cyclins are positive regulators of the cyclin/CDK kinase activities, redox modifications of cyclin D1 protein levels would be expected to significantly affect cyclin D1/CDK4/6 kinase activities.
A resultant effect from post-translational modification in one amino acid affecting the post-translational modification in a neighboring amino acid was originally suggested for the ‘binary switches’ concept for the histone code hypothesis (Fischle et al., 2003). Histone modifications are both highly reversible (e.g. lysine acetylation and serine/threonine phosphorylation) and stable (e.g. lysine and arginine methylation). Thus, methyl/phos and acetyl/phos switching mechanisms have been suggested to play a key role in controlling the genomes’ capacity to inherit, store and release biological information. Likewise, a redox-dependent phosphorylation (redox–phos switch) has been shown to regulate protein kinase A and protein kinase B (AKT) activities (Murata et al., 2003; Humphries et al., 2005). The C-subunit of protein kinase A contains two cysteine residues (Cys199 and Cys343) located close to two major phosphorylation sites (Thr197 and Ser338). Thiol–redox modification of Cys199 and Cys343 results in disulfide bond formation that enhances Thr197 and Ser338 dephosphorylation. These redox modifications of protein kinase A phosphorylation decreased its activity (Humphries et al., 2005). Likewise, oxidative stress-induced disulfide bond formation between Cys297 and Cys311 decreased AKT kinase activity by activating protein phosphatase 2A-dependent dephosphorylation of Thr308 and Ser473 (Murata et al., 2003). It will be of interest to determine if a redox–phos switch regulates cyclin D1 protein levels.
A redox-dependent transcriptional response of cyclin D1 gene expression to mitogenic signaling has also been reported in cells re-entering the cell cycle from the G0 phase (Burch and Heintz, 2005). The promoter region of cyclin D1 gene contains binding sites for redox-sensitive transcription factors including cyclic adenosine monophosphate response element-binding protein, nuclear factor-κB (NF-κB), activator protein 1 and Sp1. Several reports show redox modifications of critical cysteines in these transcription factors regulate their DNA-binding activities (Abate et al., 1990; Toledano and Leonard, 1991; Manome et al., 1993; Ohba et al., 1994; Janssen et al., 1995; Lo and Cruz, 1995). Hence, redox-state-dependent modulation of the activity of these transcription factors could significantly influence cyclin D1 transcription.
Earlier work of Rainwater et al. (1995) shows that cysteine redox modifications regulate p53 DNA-binding activity. The tumor suppressor protein p53 is a transcription factor regulating transcription of numerous genes following genotoxic insult and is frequently mutated in many cancer types. The DNA-binding domain of p53 contains four conserved cysteine residues (135, 141, 275 and 277); site-directed mutagenesis of residues 275 and 277 demonstrates these cysteines play a role in redox regulation of p53 binding to DNA in vitro (Rainwater et al., 1995). Downstream targets of p53 are numerous and include p21, GPx and many other redox enzymes involved in multiple cellular processes (Di Leonardo et al., 1994; Polyak et al., 1997; Tan et al., 1999). Adenosine diphosphate ribosylation factor (ARF), another tumor-suppressing protein encoded by the inhibitors of CDK4 (INK4)/ARF locus, inhibits Mdm2-mediated p53 degradation. In the presence of oxidizing agents like H2O2, ARF reversibly forms oligomers involving three critical cysteine residues (Menendez et al., 2003). These reports suggest that fluctuations in cellular redox environment could regulate the activity of multiple proteins that are known to control numerous biological processes including proliferation.
As cells transit through G1, cyclin E is the next cyclin to be synthesized. Cyclin E kinase activity, in association with CDK2, peaks in late G1. Once cells enter the S phase, cyclin E is degraded and CDK2 then associates with cyclin A. It is currently unknown if fluctuations in intracellular redox state influence cyclin E protein levels and activity.
Retinoblastoma and E2F
Once assembled and activated, cyclin D–CDK4/6 kinase can phosphorylate the retinoblastoma (Rb) protein (Pan et al., 2001). Expression and phosphorylation of the Rb protein is redox regulated in human natural killer cells and fibroblasts treated with thiol antioxidants (Yamauchi and Bloom, 1997; Menon et al., 2003). Rb phosphorylation releases the transcription factor E2F (Nevins, 1992), which subsequently upregulates E2F-responsive genes required to enter the S phase. As discussed earlier, entry into the S phase is preceded by a transient increase in pro-oxidant levels (Menon et al., 2003), which is consistent with a decrease in MnSOD activity in the S phase (Li and Oberley, 1998). However, persistent increase in E2F is deleterious and induces apoptosis in serum-starved NIH3T3 fibroblasts (Tanaka et al., 2002). E2F overexpression-induced apoptosis is associated with increased ROS levels and inactivation of MnSOD transcription. E2F competes with the p50 subunit of NF-κB resulting in NF-κB inactivation. Inactivation of NF-κB decreased MnSOD transcription, which could then result in increased ROS levels (Tanaka et al., 2002). These results further support the threshold concept of ROS function; lower levels of ROS are growth stimulatory, whereas higher levels of ROS are detrimental. Furthermore, these results also suggest that a feedback mechanism could link cell cycle regulatory proteins and antioxidant enzymes activity, thereby maintaining an appropriate redox environment conducive for proliferation. Additionally, MnSOD being a mitochondrial ROS-detoxifying enzyme, these results also suggest regulated levels of mitochondrial oxidants could influence cell cycle regulatory protein levels and activity, which subsequently regulates progression from one cell cycle phase to the next (Figure 1).
Redox regulation of mitotic cyclins
Once in the G2 phase of the cycle, cyclin A binds and forms an active kinase complex with CDK1 that help cells to transit into mitosis. Progression through the M phase requires the activity of another phase-specific complex, cyclin B-CDK1 (Pines and Hunter, 1989). A recent report shows CDK1 phosphorylates peroxiredoxin I inhibiting its oxidase activity. It is hypothesized that transient inhibition in peroxiredoxin I oxidase activity by mitotic cyclin/CDK phosphorylation would accumulate H2O2, which then stimulates progression from G2 to M (Chang et al., 2002). Another protein critical to G2 checkpoint control and DNA repair is topoisomerase IIα (Topo IIα). We have shown previously the expression of Topo IIα is cell cycle regulated, primarily via changes in its mRNA stability (Goswami et al., 1996). Topo IIα mRNA stability is regulated by redox-dependent protein binding to specific mRNA sequence in its 3′-untranslated region. Furthermore, our results also show increased fluorescence of a pro-oxidant dye in the G2 and M phases of the HeLa cell cycle, indicative of an increased oxidation state and correlative with enhanced accumulation of Topo IIα mRNA levels. In contrast, the decreased oxidation state in G1 correlates with decrease in Topo IIα mRNA levels (Goswami et al., 2000). These results further support the hypothesis that periodic oscillations in ROS levels owing to changes in cellular metabolic redox reactions (redox cycle) could regulate the protein and activity of cyclin/CDKs, which in turn regulates progression from G0/G1 to S to G2 and M phases (Figure 1).
Redox regulation of CKIs
Whereas cyclin–CDK kinase complex activities act as positive regulators of the cell cycle, there is also a negative regulation in the form of CKIs, essentially functioning as the brakes during cellular transit (Grana and Reddy, 1995). CKIs play a critical role in regulating the assembly and activity of the cyclin–CDK kinase complexes and are broadly classified into two families: INK4 and Cip/Kip (CDK inhibitory protein)/(kinase inhibitory protein). The INK4 family, consisting of four members (p16 (INK4A), p15 (INK4B), p18 (INK4C) and p19 (INK4D)), specifically binds and inhibits CDK4/6. Alternatively, the Cip/Kip family (p21cip, p27kip1 and p57kip2) has a broader specificity in inhibiting CDK activity. Although, comparatively, little is known about the redox regulation of most of the CKIs, several studies have established the effect of various redox manipulations on p21 protein expression. p21 protein is induced in response to the antioxidant epigallocatechin gallate; it binds and inhibits cyclin D1, thereby inhibiting proliferation of MCF10A cells (Liberto and Cobrinik, 2000). p21 and p16 are involved in mediating NAC-induced G1 arrest, which is p53 independent (Liu et al., 1999). p21 is also regulated in a p53-independent manner by a post-transcriptional mechanism in cells exposed to diethymaleate, which depletes cellular GSH (Esposito et al., 1997). We have shown previously that overexpression of MnSOD altered p21 and p16 protein levels in normal human skin fibroblasts (Sarsour et al., 2005). These results suggest that p21 levels are sensitive to both antioxidants and pro-oxidants, indicating a pro-oxidant–antioxidant balance is required for the maintenance of appropriate levels of p21 during the cell cycle. Although the mechanisms regulating redox control of CKIs are unknown, it is speculated that redox-sensitive protein binding to the 3′-untranslated region mRNA could influence CKIs mRNA turnover and/or translation. We have previously proposed such a mechanism for cell cycle phase-dependent and redox regulation of Topo IIα expression, a G2 checkpoint protein (Goswami et al., 2000).
Redox regulation of Cdc25 phosphatase
Cdc25 is a family of dual-specific phosphatases that dephosphorylate pThr14 and pTyr15 on CDK and activate cyclin–CDK complex kinase activity (Sebastian et al., 1993). Dunphy and Kumagai (1991) showed in vitro that phosphatase activity of cdc25 protein can be inhibited using N-ethylmaleimide (thiol-alkylating agent) or mutating a single conserved cysteine residue. Recently, Savitsky and Finkel (2002) reported redox modification of Cdc25c protein in HeLa cells exposed to H2O2. H2O2 caused the formation of an intramolecular disulfide bond between two critical site cysteines (Cys377 and Cys330) on the protein that resulted in its enhanced binding to 14-3-3 and subsequent degradation. Cdc25c harboring double mutants of Cys377 and Cys330 were resistant to H2O2-induced degradation (Savitsky and Finkel, 2002). The consequence of these mutations on redox-sensitive progression during the cell cycle will require additional studies. However, these results do support the hypothesis that a feedback mechanism links metabolic redox reactions (ROS signaling) to the redox-sensitive cell cycle proteins, regulating normal cell cycle progression (Figure 1).
In conclusion, compelling evidence from numerous observations supports our hypothesis that periodic oscillations in metabolic redox reactions (a redox cycle) represents a fundamental mechanism linking oxidative metabolic processes to cell cycle regulatory processes (Figure 1). The presence of redox-sensitive motifs (cysteine residues, metal co-factors in kinases and phosphatases) in several cell cycle-regulatory proteins indicate fluctuations in intracellular redox state could play a central role in regulating progression through the cell cycle. The periodicity in intracellular redox state can be regulated by a delicate balance between production of ROS (cellular metabolism) and subsequent removal by antioxidants (both non-enzymatic and enzymatic pathways). Therefore, defects in ROS production and their removal could perturb the redox cycle, which in turn could lead to aberrant proliferation. Proliferative disorders are central to a variety of human pathophysiological conditions, for example, cancer, neurodegenerative disorders, atherosclerosis, aging, hyperplasia-induced fibrotic responses and wound healing. Interestingly, many of these proliferative disorders are also associated with defects in the antioxidant pathways, presumably affecting redox regulation of cellular proliferation. In fact, our earlier results demonstrate a differential redox response between non-malignant and malignant cells exposed to NAC. Although NAC exposure induced G1 delay in non-malignant human breast epithelial cells, the same treatment did not perturb cell cycle progression in malignant breast epithelial cells (Menon et al., 2005). Modulation of the redox state with NAC is also known to induce apoptosis selectively in transformed and tumor-derived cells compared to normal cells (Havre et al., 2002). These results strongly support the hypothesis that loss of a redox control of the cell cycle could lead to aberrant proliferation. Therefore, it is hypothesized that re-establishing the redox cycle by manipulating the antioxidant pathways could reverse many aspects of aberrant cellular proliferation.
Abate C, Luk D, Gagne E, Roeder RG, Curran T . (1990). Fos and jun cooperate in transcriptional regulation via heterologous activation domains. Mol Cell Biol 10: 5532–5535.
Bae YS, Kang SW, Seo MS, Baines IC, Tekle E, Chock PB et al. (1997). Epidermal growth factor (EGF)-induced generation of hydrogen peroxide. Role in EGF receptor-mediated tyrosine phosphorylation. J Biol Chem 272: 217–221.
Bloomfield G, Pears C . (2003). Superoxide signalling required for multicellular development of Dictyostelium. J Cell Sci 116 (Part 16): 3387–3397.
Bokoch GM, Knaus UG . (2003). NADPH oxidases: not just for leukocytes anymore!. Trends Biochem Sci 28: 502–508.
Brar SS, Kennedy TP, Whorton AR, Murphy TM, Chitano P, Hoidal JR . (1999). Requirement for reactive oxygen species in serum-induced and platelet-derived growth factor-induced growth of airway smooth muscle. J Biol Chem 274: 20017–20026.
Bredt DS, Hwang PM, Glatt CE, Lowenstein C, Reed RR, Snyder SH . (1991). Cloned and expressed nitric oxide synthase structurally resembles cytochrome P-450 reductase. Nature 351: 714–718.
Brown MR, Miller Jr FJ, Li WG, Ellingson AN, Mozena JD, Chatterjee P et al. (1999). Overexpression of human catalase inhibits proliferation and promotes apoptosis in vascular smooth muscle cells. Circ Res 85: 524–533.
Burch PM, Heintz NH . (2005). Redox regulation of cell-cycle re-entry: cyclin D1 as a primary target for the mitogenic effects of reactive oxygen and nitrogen species. Antioxid Redox Signal 7: 741–751.
Burdon RH, Gill V . (1993). Cellularly generated active oxygen species and HeLa cell proliferation. Free Radical Res Commun 19: 203–213.
Burdon RH, Gill V, Rice-Evans C . (1989). Cell proliferation and oxidative stress. Free Radical Res Commun 7: 149–159.
Burdon RH, Gill V, Rice-Evans C . (1990). Oxidative stress and tumour cell proliferation. Free Radical Res Commun 11: 65–76.
Burdon RH, Rice-Evans C . (1989). Free radicals and the regulation of mammalian cell proliferation. Free Radical Res Commun 6: 345–358.
Chang TS, Jeong W, Choi SY, Yu S, Kang SW, Rhee SG . (2002). Regulation of peroxiredoxin I activity by Cdc2-mediated phosphorylation. J Biol Chem 277: 25370–25376.
Chiarugi P, Pani G, Giannoni E, Taddei L, Colavitti R, Raugei G et al. (2003). Reactive oxygen species as essential mediators of cell adhesion: the oxidative inhibition of a FAK tyrosine phosphatase is required for cell adhesion. J Cell Biol 161: 933–944.
Conour JE, Graham WV, Gaskins HR . (2004). A combined in vitro/bioinformatic investigation of redox regulatory mechanisms governing cell cycle progression. Physiol Genom 18: 196–205.
Demasi M, Silva GM, Netto LE . (2003). 20S proteasome from Saccharomyces cerevisiae is responsive to redox modifications and is S-glutathionylated. J Biol Chem 278: 679–685.
DeYulia Jr. GJ, Carcamo JM, Borquez-Ojeda O, Shelton CC, Golde DW . (2005). Hydrogen peroxide generated extracellularly by receptor–ligand interaction facilitates cell signaling. Proc Natl Acad Sci USA 102: 5044–5049.
Di Leonardo A, Linke SP, Clarkin K, Wahl GM . (1994). DNA damage triggers a prolonged p53-dependent G1 arrest and long-term induction of Cip1 in normal human fibroblasts. Genes Dev 8: 2540–2551.
Di Mascio P, Murphy ME, Sies H . (1991). Antioxidant defense systems: the role of carotenoids, tocopherols, and thiols. Am J Clin Nutr 53 (Suppl 1): 194S–200S.
Diehl JA, Cheng M, Roussel MF, Sherr CJ . (1998). Glycogen synthase kinase-3beta regulates cyclin D1 proteolysis and subcellular localization. Genes Dev 12: 3499–3511.
Dunphy WG, Kumagai A . (1991). The cdc25 protein contains an intrinsic phosphatase activity. Cell 67: 189–196.
Esposito F, Cuccovillo F, Vanoni M, Cimino F, Anderson CW, Appella E et al. (1997). Redox-mediated regulation of p21(waf1/cip1) expression involves a post-transcriptional mechanism and activation of the mitogen-activated protein kinase pathway. Eur J Biochem 245: 730–737.
Evans T, Rosenthal ET, Youngblom J, Distel D, Hunt T . (1983). Cyclin: a protein specified by maternal mRNA in sea urchin eggs that is destroyed at each cleavage division. Cell 33: 389–396.
Fernandes AP, Holmgren A . (2004). Glutaredoxins: glutathione-dependent redox enzymes with functions far beyond a simple thioredoxin backup system. Antioxid Redox Signal 6: 63–74.
Fischle W, Wang Y, Allis CD . (2003). Binary switches and modification cassettes in histone biology and beyond. Nature 425: 475–479.
Folz RJ, Crapo JD . (1994). Extracellular superoxide dismutase (SOD3): tissue-specific expression, genomic characterization, and computer-assisted sequence analysis of the human EC SOD gene. Genomics 22: 162–171.
Fridovich I . (1989). Superoxide dismutases. An adaptation to a paramagnetic gas. J Biol Chem 264: 7761–7764.
Gamou S, Shimizu N . (1995). Hydrogen peroxide preferentially enhances the tyrosine phosphorylation of epidermal growth factor receptor. FEBS Lett 357: 161–164.
Go YM, Gipp JJ, Mulcahy RT, Jones DP . (2004). H2O2-dependent activation of GCLC-ARE4 reporter occurs by mitogen-activated protein kinase pathways without oxidation of cellular glutathione or thioredoxin-1. J Biol Chem 279: 5837–5845.
Goswami PC, Roti Roti JL, Hunt CR . (1996). The cell cycle-coupled expression of topoisomerase IIalpha during S phase is regulated by mRNA stability and is disrupted by heat shock or ionizing radiation. Mol Cell Biol 16: 1500–1508.
Goswami PC, Sheren J, Albee LD, Parsian A, Sim JE, Ridnour LA et al. (2000). Cell cycle-coupled variation in topoisomerase IIalpha mRNA is regulated by the 3′-untranslated region. Possible role of redox-sensitive protein binding in mRNA accumulation. J Biol Chem 275: 38384–38392.
Grana X, Reddy EP . (1995). Cell cycle control in mammalian cells: role of cyclins, cyclin dependent kinases (CDKs), growth suppressor genes and cyclin-dependent kinase inhibitors (CKIs). Oncogene 11: 211–219.
Halliwell B, Gutteridge JM . (1992). Biologically relevant metal ion-dependent hydroxyl radical generation. An update. FEBS Lett 307: 108–112.
Halliwell B, Gutteridge JMC . (1999). Free Radicals in Biology and Medicine 3rd edn. Oxford University Press: New York.
Hartwell LH, Culotti J, Reid B . (1970). Genetic control of the cell-division cycle in yeast. I. Detection of mutants. Proc Natl Acad Sci USA 66: 352–359.
Havre PA, O’Reilly S, McCormick JJ, Brash DE . (2002). Transformed and tumor-derived human cells exhibit preferential sensitivity to the thiol antioxidants, N-acetyl cysteine and penicillamine. Cancer Res 62: 1443–1449.
Humphries KM, Deal MS, Taylor SS . (2005). Enhanced dephosphorylation of cAMP-dependent protein kinase by oxidation and thiol modification. J Biol Chem 280: 2750–2758.
Hutter DE, Till BG, Greene JJ . (1997). Redox state changes in density-dependent regulation of proliferation. Exp Cell Res 232: 435–438.
Jahngen-Hodge J, Obin MS, Gong X, Shang F, Nowell Jr TR, Gong J et al. (1997). Regulation of ubiquitin-conjugating enzymes by glutathione following oxidative stress. J Biol Chem 272: 28218–28226.
Janssen YM, Heintz NH, Mossman BT . (1995). Induction of c-fos and c-jun proto-oncogene expression by asbestos is ameliorated by N-acetyl-L-cysteine in mesothelial cells. Cancer Res 55: 2085–2089.
Jonas CR, Ziegler TR, Gu LH, Jones DP . (2002). Extracellular thiol/disulfide redox state affects proliferation rate in a human colon carcinoma (Caco2) cell line. Free Radical Biol Med 33: 1499–1506.
Jones DP, Go YM, Anderson CL, Ziegler TR, Kinkade Jr JM, Kirlin WG . (2004). Cysteine/cystine couple is a newly recognized node in the circuitry for biologic redox signaling and control. FASEB J 18: 1246–1248.
Junn E, Lee KN, Ju HR, Han SH, Im JY, Kang HS et al. (2000). Requirement of hydrogen peroxide generation in TGF-beta 1 signal transduction in human lung fibroblast cells: involvement of hydrogen peroxide and Ca2+ in TGF-beta 1-induced IL-6 expression. J Immunol 165: 2190–2197.
Kawamura N . (1960). Cytochemical and quantitative study of protein-bound sulfhydryl and disulfide groups in eggs of Arbacia during the first cleavage. Exp Cell Res 20: 127–138.
Kim KY, Rhim T, Choi I, Kim SS . (2001). N-acetylcysteine induces cell cycle arrest in hepatic stellate cells through its reducing activity. J Biol Chem 276: 40591–40598.
Klebanoff SJ . (2005). Myeloperoxidase: friend and foe. J Leukocyte Biol 77: 598–625.
Knapp LT, Klann E . (2000). Superoxide-induced stimulation of protein kinase C via thiol modification and modulation of zinc content. J Biol Chem 275: 24136–24145.
Kyaw M, Yoshizumi M, Tsuchiya K, Izawa Y, Kanematsu Y, Fujita Y et al. (2004). Antioxidant effects of stereoisomers of N-acetylcysteine (NAC), L-NAC and D-NAC, on angiotensin II-stimulated MAP kinase activation and vascular smooth muscle cell proliferation. J Pharmacol Sci 95: 483–486.
Laurent A, Nicco C, Chereau C, Goulvestre C, Alexandre J, Alves A et al. (2005). Controlling tumor growth by modulating endogenous production of reactive oxygen species. Cancer Res 65: 948–956.
Li N, Oberley TD . (1998). Modulation of antioxidant enzymes, reactive oxygen species, and glutathione levels in manganese superoxide dismutase-overexpressing NIH/3T3 fibroblasts during the cell cycle. J Cell Physiol 177: 148–160.
Liberto M, Cobrinik D . (2000). Growth factor-dependent induction of p21(CIP1) by the green tea polyphenol, epigallocatechin gallate. Cancer Lett 154: 151–161.
Limoli CL, Rola R, Giedzinski E, Mantha S, Huang TT, Fike JR . (2004). Cell-density-dependent regulation of neural precursor cell function. Proc Natl Acad Sci USA 101: 16052–16057.
Liu M, Wikonkal NM, Brash DE . (1999). Induction of cyclin-dependent kinase inhibitors and G(1) prolongation by the chemopreventive agent N-acetylcysteine. Carcinogenesis 20: 1869–1872.
Lo YY, Cruz TF . (1995). Involvement of reactive oxygen species in cytokine and growth factor induction of c-fos expression in chondrocytes. J Biol Chem 270: 11727–11730.
Manome Y, Datta R, Taneja N, Shafman T, Bump E, Hass R et al. (1993). Coinduction of c-jun gene expression and internucleosomal DNA fragmentation by ionizing radiation. Biochemistry 32: 10607–10613.
Martinez Munoz C, Post JA, Verkleij AJ, Verrips CT, Boonstra J . (2001). The effect of hydrogen peroxide on the cyclin D expression in fibroblasts. Cell Mol Life Sci 58: 990–996.
Massey V, Strickland S, Mayhew SG, Howell LG, Engel PC, Matthews RG et al. (1969). The production of superoxide anion radicals in the reaction of reduced flavins and flavoproteins with molecular oxygen. Biochem Biophys Res Commun 36: 891–897.
Masui Y, Markert CL . (1971). Cytoplasmic control of nuclear behavior during meiotic maturation of frog oocytes. J Exp Zool 177: 129–145.
Mauro F, Grasso A, Tolmach LJ . (1969). Variations in sulfhydryl, disulfide, and protein content during synchronous and asynchronous growth of HeLa cells. Biophys J 9: 1377–1397.
McCord JM, Fridovich I . (1969). Superoxide dismutase. An enzymic function for erythrocuprein (hemocuprein). J Biol Chem 244: 6049–6055.
Menendez S, Khan Z, Coomber DW, Lane DP, Higgins M, Koufali MM et al. (2003). Oligomerization of the human ARF tumor suppressor and its response to oxidative stress. J Biol Chem 278: 18720–18729.
Menon SG, Coleman MC, Walsh SA, Spitz DR, Goswami PC . (2005). Differential susceptibility of nonmalignant human breast epithelial cells and breast cancer cells to thiol antioxidant-induced G(1)-delay. Antioxid Redox Signal 7: 711–718.
Menon SG, Sarsour EH, Spitz DR, Higashikubo R, Sturm M, Zhang H et al. (2003). Redox regulation of the G1 to S phase transition in the mouse embryo fibroblast cell cycle. Cancer Res 63: 2109–2117.
Murata H, Ihara Y, Nakamura H, Yodoi J, Sumikawa K, Kondo T . (2003). Glutaredoxin exerts an antiapoptotic effect by regulating the redox state of Akt. J Biol Chem 278: 50226–50233.
Nevins JR . (1992). E2F: a link between the Rb tumor suppressor protein and viral oncoproteins. Science 258: 424–429.
Oberley TD, Oberley LW, Slattery AF, Lauchner LJ, Elwell JH . (1990). Immunohistochemical localization of antioxidant enzymes in adult Syrian hamster tissues and during kidney development. Am J Pathol 137: 199–214.
Oberley TD, Schultz JL, Li N, Oberley LW . (1995). Antioxidant enzyme levels as a function of growth state in cell culture. Free Radical Biol Med 19: 53–65.
Obin M, Shang F, Gong X, Handelman G, Blumberg J, Taylor A . (1998). Redox regulation of ubiquitin-conjugating enzymes: mechanistic insights using the thiol-specific oxidant diamide. FASEB J 12: 561–569.
Ohba M, Shibanuma M, Kuroki T, Nose K . (1994). Production of hydrogen peroxide by transforming growth factor-beta 1 and its involvement in induction of egr-1 in mouse osteoblastic cells. J Cell Biol 126: 1079–1088.
Okuyama H, Shimahara Y, Kawada N, Seki S, Kristensen DB, Yoshizato K et al. (2001). Regulation of cell growth by redox-mediated extracellular proteolysis of platelet-derived growth factor receptor beta. J Biol Chem 276: 28274–28280.
Pan W, Cox S, Hoess RH, Grafstrom RH . (2001). A cyclin D1/cyclin-dependent kinase 4 binding site within the C domain of the retinoblastoma protein. Cancer Res 61: 2885–2891.
Pani G, Colavitti R, Bedogni B, Anzevino R, Borrello S, Galeotti T . (2000). A redox signaling mechanism for density-dependent inhibition of cell growth. J Biol Chem 275: 38891–38899.
Pardee AB . (1974). A restriction point for control of normal animal cell proliferation. Proc Natl Acad Sci USA 71: 1286–1290.
Pines J, Hunter T . (1989). Isolation of a human cyclin cDNA: evidence for cyclin mRNA and protein regulation in the cell cycle and for interaction with p34cdc2. Cell 58: 833–846.
Polyak K, Xia Y, Zweier JL, Kinzler KW, Vogelstein B . (1997). A model for p53-induced apoptosis. Nature 389: 300–305.
Rainwater R, Parks D, Anderson ME, Tegtmeyer P, Mann K . (1995). Role of cysteine residues in regulation of p53 function. Mol Cell Biol 15: 3892–3903.
Rapkine L . (1931). Su les processus chimiques au cours de la division cellulaire. Ann Physiol Physiochem Biol 7: 382–418.
Rhee SG . (1999). Redox signaling: hydrogen peroxide as intracellular messenger. Exp Mol Med 31: 53–59.
Rhee SG, Kang SW, Netto LE, Seo MS, Stadtman ER . (1999). A family of novel peroxidases, peroxiredoxins. Biofactors 10: 207–209.
Sarsour EH, Agarwal M, Pandita TK, Oberley LW, Goswami PC . (2005). Manganese superoxide dismutase protects the proliferative capacity of confluent normal human fibroblasts. J Biol Chem 280: 18033–18041.
Savitsky PA, Finkel T . (2002). Redox regulation of Cdc25C. J Biol Chem 277: 20535–20540.
Schafer FQ, Buettner GR . (2001). Redox environment of the cell as viewed through the redox state of the glutathione disulfide/glutathione couple. Free Radical Biol Med 30: 1191–1212.
Schrader M, Fahimi HD . (2004). Mammalian peroxisomes and reactive oxygen species. Histochem Cell Biol 122: 383–393.
Sebastian B, Kakizuka A, Hunter T . (1993). Cdc25M2 activation of cyclin-dependent kinases by dephosphorylation of threonine-14 and tyrosine-15. Proc Natl Acad Sci USA 90: 3521–3524.
Sherr CJ . (1995). Mammalian G1 cyclins and cell cycle progression. Proc Assoc Am Physicians 107: 181–186.
Stirpe F, Higgins T, Tazzari PL, Rozengurt E . (1991). Stimulation by xanthine oxidase of 3T3 Swiss fibroblasts and human lymphocytes. Exp Cell Res 192: 635–638.
Sundaresan M, Yu ZX, Ferrans VJ, Irani K, Finkel T . (1995). Requirement for generation of H2O2 for platelet-derived growth factor signal transduction. Science 270: 296–299.
Tan M, Li S, Swaroop M, Guan K, Oberley LW, Sun Y . (1999). Transcriptional activation of the human glutathione peroxidase promoter by p53. J Biol Chem 274: 12061–12066.
Tanaka H, Matsumura I, Ezoe S, Satoh Y, Sakamaki T, Albanese C et al. (2002). E2F1 and c-Myc potentiate apoptosis through inhibition of NF-kappaB activity that facilitates MnSOD-mediated ROS elimination. Mol cell 9: 1017–1029.
Tanaka T, Nakamura H, Nishiyama A, Hosoi F, Masutani H, Wada H et al. (2000). Redox regulation by thioredoxin superfamily; protection against oxidative stress and aging. Free Radical Res 33: 851–855.
Temin HM . (1971). Stimulation by serum of multiplication of stationary chicken cells. J Cell Physiol 78: 161–170.
Toledano MB, Leonard WJ . (1991). Modulation of transcription factor NF-kappa B binding activity by oxidation–reduction in vitro. Proc Natl Acad Sci USA 88: 4328–4332.
Tu BP, Kudlicki A, Rowicka M, McKnight SL . (2005). Logic of the yeast metabolic cycle: temporal compartmentalization of cellular processes. Science 310: 1152–1158.
Wang HP, Schafer FQ, Goswami PC, Oberley LW, Buettner GR . (2003). Phospholipid hydroperoxide glutathione peroxidase induces a delay in G1 of the cell cycle. Free Radical Res 37: 621–630.
Yamamoto K, Volkl A, Hashimoto T, Fahimi HD . (1988). Catalase in guinea pig hepatocytes is localized in cytoplasm, nuclear matrix and peroxisomes. Eur J Cell Biol 46: 129–135.
Yamauchi A, Bloom ET . (1997). Control of cell cycle progression in human natural killer cells through redox regulation of expression and phosphorylation of retinoblastoma gene product protein. Blood 89: 4092–4099.
Zangar RC, Davydov DR, Verma S . (2004). Mechanisms that regulate production of reactive oxygen species by cytochrome P450. Toxicol Appl Pharmacol 199: 316–331.
Zou Y, Ewton DZ, Deng X, Mercer SE, Friedman E . (2004). Mirk/dyrk1B kinase destabilizes cyclin D1 by phosphorylation at threonine 288. J Biol Chem 279: 27790–27798.
We thank Ms Kellie Bodeker with editorial assistance. This work was supported by funding from NIH CA 111365 and The University of Iowa Carver Trust.
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Menon, S., Goswami, P. A redox cycle within the cell cycle: ring in the old with the new. Oncogene 26, 1101–1109 (2007). https://doi.org/10.1038/sj.onc.1209895
- antioxidant enzymes
- cell cycle
- cyclin D1
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