Activation of extracellular signal‐regulated kinase (ERK) in G₂ phase delays mitotic entry through p21 ^CIP1

Abstract

Abstract.  Extracellular signal‐regulated kinase activity is essential for mediating cell cycle progression from G₁ phase to S phase (DNA synthesis). In contrast, the role of extracellular signal‐regulated kinase during G₂ phase and mitosis (M phase) is largely undefined. Previous studies have suggested that inhibition of basal extracellular signal‐regulated kinase activity delays G₂‐ and M‐phase progression. In the current investigation, we have examined the consequence of activating the extracellular signal‐regulated kinase pathway during G₂ phase on subsequent progression through mitosis. Using synchronized HeLa cells, we show that activation of the extracellular signal‐regulated kinase pathway with phorbol 12‐myristate 13‐acetate or epidermal growth factor during G₂ phase causes a rapid cell cycle arrest in G₂ as measured by flow cytometry, mitotic indices and cyclin B1 expression. This G₂‐phase arrest was reversed by pre‐treatment with bisindolylmaleimide or U0126, which are selective inhibitors of protein kinase C proteins or the extracellular signal‐regulated kinase activators, MEK1/2, respectively. The extracellular signal‐regulated kinase‐mediated delay in M‐phase entry appeared to involve de novo synthesis of the cyclin‐dependent kinase inhibitor, p21 ^CIP1, during G₂ through a p53‐independent mechanism. To establish a function for the increased expression of p21 ^CIP1 and delayed cell cycle progression, we show that extracellular signal‐regulated kinase activation in G₂‐phase cells results in an increased number of cells containing chromosome aberrations characteristic of genomic instability. The presence of chromosome aberrations following extracellular signal‐regulated kinase activation during G₂‐phase was further augmented in cells lacking p21 ^CIP1 . These findings suggest that p21 ^CIP1 mediated inhibition of cell cycle progression during G₂/M phase protects against inappropriate activation of signalling pathways, which may cause excessive chromosome damage and be detrimental to cell survival.

INTRODUCTION

The extracellular signal‐regulated kinases 1 and 2 (ERK1/2) are members of the mitogen‐activated protein (MAP) kinase family and are involved in a variety of cellular functions including proliferation, differentiation, apoptosis and motility (Lewis et al. 1998; Pearson et al. 2001). ERK protein activation most often occurs sequentially through Ras G‐proteins, Raf kinases, and MAP/ERK kinases 1 and 2 (MEK1/2), which are the only known activators of ERK1/2 (Lewis et al. 1998). Upon activation, the ERK pathway promotes cell proliferation by indirect regulation of the activity of cyclin‐dependent kinases (Cdk) at specific times during the cell cycle. For example, Ras G‐protein activity couples growth factor receptor signalling to ERK proteins, which promote cyclin D1 expression, Cdk2 activity, and G₁/S‐phase progression (Liu et al. 1995; Aktas et al. 1997). Ras also promotes progression through G₁‐phase by increasing cyclin E/Cdk2 activity in cooperation with the ERK substrate, c‐Myc (Leone et al. 1997). Activated Cdk proteins phosphorylate the retinoblastoma (Rb) tumour‐suppressor protein, which acts as a cell cycle repressor of transcription during G₁‐phase in its hypophosphorylated form (Hatakeyama & Weinberg 1995). Thus, Ras, indirectly through the activation of Cdk proteins, causes Rb phosphorylation and inhibition of Rb repression of genes required for entry into S phase (Mittnacht et al. 1997; Peeper et al. 1997). As expected, proteins downstream of Ras, including Raf, MEK, and ERK, have also been shown to play a central role in promoting entry into S‐phase progression by regulating the expression of cyclins and activation of Cdk proteins (Weber et al. 1997; Cheng et al. 1998; Lents et al. 2002).

The ERK pathway may also promote cell cycle progression by regulating the expression of Cdk inhibitors such as p21 ^CIP1 (Sewing et al. 1997; Kivinen & Laiho 1999; Coleman et al. 2003). For example, many growth factors that require Ras signalling through ERK to increase cell proliferation also induce p21 ^CIP1 expression (Leone et al. 1997; Kivinen & Laiho 1999). Similarly, pharmacological inhibition of MEK1/2 with PD98059 blocks growth factor‐mediated G₁/S‐phase progression as well as p21 ^CIP1 expression (Kivinen & Laiho 1999). Although it may appear counter productive for the Ras/Raf/MEK/ERK pathway to promote the expression of both cyclin D proteins and the p21 ^CIP1 inhibitor, studies using cells that do not express the p21 ^CIP1 gene show a reduced ability to activate cyclin D/Cdk complexes and promote G₁/S‐phase progression (Cheng et al. 1999). Given that increased p21 ^CIP1 expression is observed in more that 90% of breast cancer tissues that also over‐express cyclin D proteins (Russell et al. 1999), elevated p21 ^CIP1 levels have been proposed to function in delaying cell cycle progression in order to protect cancer cells against DNA damaging agents that would otherwise promote apoptosis (Gartel & Radhakrishnan 2005). Thus, tumour cells containing oncogenic Ras or Raf proteins may regulate the expression of cyclins and cell cycle inhibitors, which may simultaneously provide cells with a proliferative advantage and protection against damaging agents during cell cycle progression.

Despite its important role in regulating G₁/S phase, the function of the ERK pathway in regulating somatic cell cycle progression through G₂ phase and mitosis (M phase) is still under investigation. Inhibition of basal ERK pathway activity, with pharmacologic agents, RNA interference, or by over‐expression of dominant negative mutants of MEK1, causes a delay in G₂ and M‐phase progression (Wright et al. 1999; Roberts et al. 2002; Liu et al. 2004). Several proteins of the ERK pathway have been shown to have increased catalytic activity during the mitotic phase. For example, a cytoplasmic form of Raf‐1 has been shown to be activated through a Ras‐independent mechanism and is uncoupled from MEK1/2 in total protein lysates isolated from cells arrested in mitosis with nocodazole (Laird et al. 1995; Ziogas et al. 1998; Laird et al. 1999). However, Raf‐1–dependent activation of MEK1 has been linked to the regulation of mitotic Golgi apparatus fragmentation (Colanzi et al. 2003). Thus, it is possible that localized Raf‐1/MEK/ERK signalling modules may be more relevant to cells progressing through G₂ and M phases. Localized ERK pathway activation is supported by several studies demonstrating the presence of active forms of MEK and ERK proteins that are localized to the nucleus, chromosome kinetochores, centrosomes, microtubules, and the Golgi complex during G₂ and M‐phase transitions (Acharya et al. 1998; Shapiro et al. 1998; Zecevic et al. 1998; Cha & Shapiro 2001; Willard & Crouch 2001). However, regulatory functions of the ERK pathway proteins and of the mitotic substrates at these intracellular locations remain largely unknown.

Extracellular signal‐regulated kinase pathway regulation during G₂/M‐phase progression may involve protein kinase C (PKC) isoforms. Earlier studies have suggested that PKC proteins activate Raf‐1 through direct phosphorylation of serines 259 and 499 (Kolch et al. 1993). However, others have suggested that these phosphorylation sites are not required for Raf‐1 activation (Barnard et al. 1998; Schonwasser et al. 1998). Alternatively, PKC may indirectly activate Raf‐1 by phosphorylating and inactivating the Raf kinase inhibitory protein (RKIP) (Corbit et al. 2003). These studies proposed that phosphorylated RKIP dissociates from Raf‐1, which allows Raf‐1 activation to occur.

The classical PKC isoforms (α, βI, βII, and γ) are activated by diacylglycerol (DAG) and calcium, whereas activation of the novel PKC isoforms (δ, ɛ, η, and θ) only requires DAG (Ron & Kazanietz 1999). DAG analogues such as phorbol esters, are potent activators of the classical and novel PKC isoforms (Ron & Kazanietz 1999). Long‐term exposure (> 12 h) to phorbol 12‐myristate‐13‐acetate (PMA) has been reported to induce G₂‐phase arrest through a mechanism involving phospholipid metabolites (Kaszkin et al. 1991) or G₁‐phase arrest through inhibition of cyclin dependent kinases and activation of p21 ^CIP1 and p27 ^KIP1 expression (Hamada et al. 1996; Frey et al. 1997). Depending on the cell line, PKC isoforms may promote or inhibit G₂/M‐phase progression. For example, PKC activity has been reported to decrease in mitotic glioma cells (Soma et al. 1994), but is elevated in promyelocytic leukaemia cells and has been involved in mitotic nuclear envelope breakdown (Goss et al. 1994). MCF‐7 breast cancer cells treated with PMA immediately after release from aphidicolin‐induced S‐phase arrest have shown increased p21 ^CIP1 expression and subsequent accumulation of cells in the G₂ phase (Barboule et al. 1999). However, the signalling events that mediate the G₂‐phase arrest in response to PMA have not been determined.

The goal of the current study was to examine the consequence of ERK activation in regulating G₂/M‐phase progression. Our findings demonstrate that ERK activation during the G₂ phase delays cell cycle progression into mitosis through a mechanism involving de novo synthesis of p21 ^CIP1 . In addition, ERK activation during G₂ phase causes an increase in the hallmarks of chromosome instability, a phenomenon that is exacerbated in cells lacking p21 ^CIP1 . These data suggest that ERK‐mediated induction of p21 ^CIP1 during G₂ phase and cell cycle arrest may protect cells from excessive DNA damage that might occur during mitotic transitions and result in compromised cell survival.

MATERIALS AND METHODS

Cell culture and reagents

HeLa cells (#CCL‐2, ATCC, Manassas, VA), human retina epithelial cells that stably express human telomerase reverse transcriptase (hTERT‐RPE cells, #CRL‐4000; ATCC, Manassas, VA), HCT116 parental, or HCT116 p21^−/– cells (kindly provided by Dr Bert Vogelstein, Johns Hopkins University) were cultured in a complete medium consisting of Dulbecco's modified Eagle's medium supplemented with 10% foetal bovine serum and antibiotics (penicillin, 100 U/ml; streptomycin, 100 µg/ml) from Invitrogen (Carlsbad, CA). Epidermal growth factor (EGF) and PMA were purchased from Sigma (St. Louis, MO) and used at final concentrations of 0.01–100 ng/ml and 0.01–0.1 µm, respectively. The MEK1/2 inhibitor, U0126, and the PKC inhibitor, bisindolylmaleimide (Bis) I, were purchased from Calbiochem (La Jolla, CA) and used at final concentrations of 10 µm and 1 µm, respectively. The proteasome inhibitor, MG115, and protein synthesis inhibitor, cyclohexamide, were purchased from Calbiochem and used at final concentrations of 10 µg/ml and 10 µm, respectively.

Antibodies specific for cyclin B1 (sc‐245) and p21 ^CIP1 (sc‐397) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Antibodies specific for phosphorylated ERK1/2 (M‐8159) and α‐tubulin (T‐6557) were purchased from Sigma. Immunoblotting analysis was performed as previously described (Dangi et al. 2003).

Cells were synchronized at the G₁/S‐phase boundary using a double thymidine block as previously described (Dangi et al. 2003). Briefly, cells (approximately 50% confluent) were treated with 2 mm thymidine in complete medium for 16 h. The cells were released back into the cell cycle by washing with Hanks buffered saline solution (HBSS, Invitrogen) and were incubated for an additional 8 h in complete medium in the absence of thymidine. Cells were treated a second time with 2 mm thymidine in complete medium for 16 h, which resulted in an average of 85–90% of them synchronized at the G₁/S‐phase boundary of the cell cycle (Dangi et al. 2003). Synchronized cells were washed with HBSS, were released back into the cell cycle and were harvested at various times after release with or without the indicated treatments. Protein lysates were collected from synchronized cells after two washes with cold phosphate buffered saline (PBS, pH 7.2; Invitrogen) by scraping with 300 µl of cold tissue lysis buffer (20 mm Tris‐base, pH 7.4, 137 mm NaCl, 2 mm ethylenediamenetetraacetic acid (EDTA), 1% Triton X‐100, 25 mmβ‐glycerophosphate, 2 mm sodium pyrophosphate, 10% glycerol, 1 mm sodium orthovanadate, 1 mm phenylmethylsulphonyl fluoride (PMSF), 1 mm benzamidine). The lysates were centrifuged at 20 000 (Xg) to remove insoluble material, were diluted with an equal volume of 2X sodium dodecyl sulphate (SDS)‐sample buffer, and the proteins were separated on SDS‐polyacrylamide gel electrophoresis for immunoblot analysis.

Immunofluorescence and mitotic index assays

HeLa or HCT116 cells were grown on round coverslips (No. 1, 18 mm; VWR, West Chester, PA) in 6‐cm tissue culture plates and then were synchronized at the G₁/S boundary as described in the previous section. Cells were released back into the cell cycle for 7 h, which corresponded to G₂ phase, and were treated with PMA or EGF. In some cases, 20–30 min prior to PMA or EGF treatment, cells were pre‐treated in the presence or absence of Bis or U0126. Immunofluorescence was performed as previously described (Cha & Shapiro 2001). Briefly, at varying times after release, coverslips were fixed with 4% paraformaldehyde diluted in PBS, from a 16% stock (Electron Microscopy Sciences, Hatfield, PA) for 5 min and were then permeabilized with 0.1% triton X‐100 for 3 min. Alternatively, cells were fixed in cold methanol (−20 °C) for 10 min prior to staining. Following blocking with bovine serum albumin, cells were incubated with antibodies against γ‐tubulin or p62 nucleoporin followed by fluoroscein or Texas red‐conjugated secondary antibodies. Cellular DNA was stained with 4′, 6‐diamidino‐2‐phenylindole (DAPI, 0.2 µg/ml in PBS) for 5 min. Mitotic chromosomes were identified, based on their characteristic condensed structure, using a Nikon E800 fluorescence microscope, and photographs were taken using a Hamamatsu CCD camera (ORCA‐ER‐285; Biovision Technologies, Exton, PA, USA). Images of mitotic cells, which included cells in prophase, pro‐metaphase, metaphase, anaphase and telophase, were processed using iplab software (Scanalytics, Inc. Fairfax, VA). Mitotic cells were expressed as a fraction of total cells counted to determine the mitotic index (MI). Cells containing multiple nuclei (two or more) were counted and expressed as a percentage of the total cell number. Centrosomes were identified by γ‐tubulin staining and cells containing greater that two centrosomes were expressed as a percentage of the total cell number. Approximately 250–300 cells were counted for each condition.

Flow cytometry

Synchronized cells were trypsinized, washed with cold PBS, fixed with cold (−20 °C) 70% ethanol and stored at 4 °C overnight. Cells were then incubated with 100 µg/mL propidium iodide (Sigma) dissolved in 0.2 M Tris pH 7.5, 20 mm EDTA, 1 mg/ml RNase A (Sigma) for 1 h at room temperature and were then diluted with an equal volume of PBS. DNA content was measured by flow cytometry (FACScan Analyser; Becton Dickinson, Franklin Lakes, NJ) and was analysed using the Sync Wizard Model, ModFit LT software (Becton Dickinson). Fifteen thousand cells were counted under each condition. To determine G₁, S, and G₂/M‐phase cell populations, the settings for 2 N and 4 N DNA content peaks were obtained within each experiment using the G₁/S arrested cells (2N) as the reference and applied to all samples within a given experiment.

RESULTS

PKC mediates PMA‐induced G₂ arrest

Protein kinase C proteins are linked to the regulation of the ERK pathway and cell cycle progression. Using PMA as a potent activator of PKC, we first addressed the requirement for PKC and ERK signalling in mediating PMA‐induced cell cycle delay during G₂ phase. HeLa cells synchronized at the G₁/S phase boundary were released back into the cell cycle, pre‐treated in the absence or presence of the PKC inhibitor, Bis during G₂ phase, and were then stimulated with or without PMA. Cells harvested at various times after release were analysed by flow cytometry for cell‐cycle progression and ERK activation. Fluorescence‐activated cell sorter (FACS) analysis of DNA content showed that untreated cells enter G₂/M phase after 7 h release from G₁/S, exit mitosis, and return to G₁ phase 13 h after release from the initial G₁/S‐phase block (Fig. 1a). In contrast, PMA‐treated cells remain primarily in G₂/M phase up to 13 h after release from G₁/S block (Fig. 1a). The PMA‐induced arrest in G₂/M phase was reversed by the PKC inhibitor, Bis I (Fig. 1a). Delays in G₂/M‐phase progression were also observed in non‐transformed telomerase immortalized human retinal pigment epithelial (hTERT‐RPE) cells (Fig. 1b) or HEK293 cells (data not shown) treated with PMA during G₂ phase.

Figure 1.

Fluorescence‐activated cell sorting (FACS) analysis of PMA‐mediated G₂‐phase arrest. HeLa cells were synchronized at G₁/S boundary and released back into the cell cycle. (a) Cells were left untreated (control) or treated with PMA (0.1 µm) at 7 h after G₁/S release, with the bisindolylmaleimide (Bis, 1 µm) at 6.5 h after G₁/S release, or with PMA and Bis together. (b) FACS analysis of synchronized hTERT cells treated in the absence (control) or presence of PMA at 7 h after G₁/S‐phase release. Cells were harvested at the times indicated after release from G₁/S and the number of cells (Y axis) containing 2N or 4N DNA content (X axis) were determined by FACS.

In agreement with previous studies (Roberts et al. 2002), basal ERK activity was elevated in G₂/M phase and PMA treatment caused sustained ERK activation for up to 6 h (Fig. 2a). Bis had little effect on basal ERK activity but it did inhibit PMA‐induced ERK activity (Fig. 2a). In control cells, cyclin B1 expression showed a typical transient expression as cells progressed through G₂ phase (7 and 7.3 h after G₁/S release), mitosis (9 h), and back into G₁ phase (11 and 13 h). In contrast, cyclin B1 expression in PMA‐treated cells was sustained for up to 13 h and supported the FACS data, indicating that these cells were arrested in the G₂ or M phase (Fig. 2a). However, pre‐treatment with Bis before PMA restored the transient expression of cyclin B1 indicate that PKC activity is required for maintaining PMA‐induced G₂/M‐phase arrest (Fig. 2a).

Figure 2.

PMA‐induced G₂‐phase arrest requires PKC activity. Cells synchronized at the G₁/S boundary were released back into the cell cycle for 6.5 h and pre‐treated with or without Bis (1 µm). At 7 h post‐G₁/S release, cells were treated in the absence or presence of PMA (0.1 µm) and harvested at the times indicated. (a) Immunoblot analysis of active ERK1/2 (ppERK1/2, top panel) and cyclin B1 (middle panel) as the marker of G₂/M progression. The expression of α‐tubulin (lower panel) is shown for a protein loading control. (b) The MI at various times after G₁/S release in controls (open bars), PMA‐treated (black bars), Bis‐treated (checkered bars), or Bis‐ and PMA‐treated (grey bars) cells. Data represent the mean and standard error from three independent experiments.

To determine whether the PMA‐induced cell cycle delay was occurring in the G₂ or the M phase, the MI was evaluated under various conditions. As shown in Fig. 2b, the MI of PMA‐treated cells at 9 and 11 h after G₁/S release was greatly reduced compared to untreated control cells. By 13 h after release, the MI of PMA‐treated cells began to increase, indicating a more than 4 h delay in mitotic entry (Fig. 2b). The delay in mitotic entry was reversed by pre‐treatment with Bis (Fig. 2b), supporting the requirement for PKC in mediating PMA‐induced G₂‐phase arrest.

ERK mediates PMA‐induced G₂‐arrest

We next examined the requirement for ERK signalling in mediating PMA‐induced G₂‐phase arrest. G₂‐phase cells generated as described in Fig. 1 were pre‐treated in the absence or presence of the MEK1/2 inhibitor, U0126, followed by PMA stimulation. Similar to inhibition of PKC, inhibition of ERK activation by U0126 also reversed most of the PMA‐induced G₂‐arrest (Fig. 3). As expected, U0126 blocked PMA‐induced ERK activation (Fig. 4a) and reversed the PMA‐induced G₂‐phase arrest as indicated by MI measurements (Fig. 4b). In addition, U0126 treatment also enhanced the MI at 13 h post‐G₁/S‐phase release (Fig. 4b) and did not reverse elevated cyclin B1 expression (Fig. 4a). These findings indicated a delay in mitotic progression and are consistent with our previous studies that have demonstrated a requirement for ERK during metaphase to anaphase transitions (Roberts et al. 2002). Thus, these data indicate that PMA‐induced G₂‐phase arrest requires PKC activation of the ERK pathway.

Figure 3.

FACS analysis of ERK dependency in mediating PMA‐induced G₂‐phase arrest. HeLa cells were synchronized at the G₁/S boundary and released back into the cell cycle. Cells were left untreated (control), treated with PMA (0.1 µm) at 7 h after release, treated with U0126 (10 µm) at 6.5 h after G₁/S release or treated with PMA and U0126 together. Cells were harvested at the times indicated after release from G₁/S and the number of cells (Y axis) containing 2N or 4N DNA content (X axis) were determined by FACS.

Figure 4.

PMA‐induced G₂‐phase arrest requires ERK activity. Cells synchronized at the G₁/S boundary were released back into the cell cycle and treated in the presence of absence of PMA (0.1 µm, 7 h after G₁/S release), U0126 (10 µm, 6.5 h after G₁/S release), or PMA and U0126 together. (a) Immunoblot analysis of ppERK1/2 (top panel), cyclin B1 (middle panel), and α‐tubulin (lower panel) as a protein loading control. (b) The MI at varying times after G₁/S release in untreated (white bars), PMA‐treated (black bars), U0126‐treated (checkered bars), or PMA‐ and U0126‐treated cells (grey bars). Data represent the mean and standard error from three independent experiments.

Activation of ERK during G₂ phase by EGF delays M‐phase progression

We next tested whether activating the ERK pathway by EGF during G₂ also affected G₂/M progression. Cells synchronized in the G₂ phase were pre‐treated with or without U0126 and then were treated in the presence or absence of EGF. FACS analysis indicated that EGF mediated an approximately 2 h delay in G₂/M progression compared to controls (Fig. 5, 11 h time point) and that this delay was reversed by pre‐treatment with U0126 (Fig. 5). As expected, EGF activation of ERK was inhibited by U0126 (Fig. 6a). Cyclin B1 degradation and mitotic entry as determined by MI were also delayed by approximately 2 h in cells treated with EGF in the G₂ phase and this was reversed by the presence of U0126 (Fig. 6a,b). As in Fig. 5b, U0126‐treated cells showed an elevated MI at the 13‐h time point (Fig. 6b), indicative of a delay in metaphase to anaphase transitions (Roberts et al. 2002). These data provide further support that the level of ERK activity during G₂ phase regulates mitotic entry and progression.

Figure 5.

FACS analysis of ERK dependency in mediating EGF‐induced delay in G₂/M progression. HeLa cells were synchronized at the G₁/S boundary and released back into the cell cycle. Cells were left untreated (control), treated with EGF (100 ng/mL) at 7 h after G₁/S release, treated with U0126 (10 µm) at 6.5 h after G₁/S release, or treated with EGF and U0126 together. Cells were harvested at the times indicated after release from G₁/S and the number of cells (Y axis) containing 2N or 4N DNA content (X axis) were determined by FACS.

Figure 6.

EGF‐induced G₂‐phase arrest requires ERK activity. Cells synchronized at the G₁/S boundary were released back into the cell cycle and treated with EGF (100 ng/ml, 7 h after G₁/S release), U0126 (10 µm, 6.5 h after G₁/S release), or both EGF and U0126. (a) Immunoblots show the levels of ppERK1/2 (top panel), cyclin B1 (middle panel) and α‐tubulin (lower panel) as a protein loading control. (b) The MI at varying times after G₁/S release in untreated (white bars), EGF‐treated (black bars), U0126‐treated (checkered bars), or EGF‐ and U0126‐treated cells (grey bars). Data represent the mean and standard error from three independent experiments.

An ERK activation threshold determines M‐phase progression

Our data suggest that PMA or EGF‐mediated activation of the ERK pathway during the G₂ phase delays mitotic entry. We next determined whether an ERK activity threshold was needed to be reached before a delay in G₂/M‐phase progression could be observed. To test this, cells synchronized at the G₁/S boundary were released into G₂ phase and were treated with varying doses of EGF or PMA. Treatment with 0.1 ng/ml of EGF caused a transient activation of the ERK pathway but had little effect on cyclin B1 expression (Fig. 7a). Increasing the EGF dose further enhanced the magnitude and duration of ERK activity and the expression of cyclin B1 at 11 h after G₁/S release as compared to untreated controls (Fig. 7a). As another measure of cell cycle progression, the expression of the Cdk inhibitor p21 ^CIP1 was measured. Whereas the lowest dose of EGF that caused ERK activation had little effect on p21 ^CIP1, higher EGF doses increased p21 ^CIP1 expression in a manner that correlated with ERK activity and cyclin B1 expression (Fig. 7a).

Figure 7.

Cyclin B1 and p21^CIP1 expression during G₂/M‐phase correlate with ERK activity. Cells synchronized in G₁/S were released back into the cell cycle and treated with the indicated dose of EGF (a) or PMA (b). Immunoblot analysis shows the level of ppERK1/2 (top panel), cyclin B1, and p21 ^CIP1 (middle panels) expression. The expression of α‐tubulin (lower panel) is shown for a protein loading control.

Similarly, the low doses of PMA (0.01 nm) activated ERK but had little effect on cyclin B1 expression as compared to controls (Fig. 7b). When PMA concentrations were increased by 10‐fold or more, maximal ERK activation was observed and a corresponding increase in cyclin B1 and p21 ^CIP1 expression consistent with G₂‐phase arrest (Fig. 7b). Thus, the level of ERK activity during G₂ phase appears to regulate cell cycle progression through regulation of p21 ^CIP1 expression.

ERK regulates p21 ^CIP1 expression during G₂ phase

Several studies support a role for the CIP/KIP family of Cdk inhibitors in regulating cell cycle progression during G₂ phase in response to various stress conditions (Niculescu et al. 1998; Coqueret 2003; Di Gennaro et al. 2003). Thus, the requirement for PKC and ERK proteins in regulating the expression of p21 ^CIP1 was examined in G₂‐phase cells treated with PMA. Increased p21 ^CIP1 expression was observed within 2 h of PMA treatment (9 h after G₁/S release) and continued to increase after 4 and 6 h treatment with PMA (11 and 13 h after G₁/S release, respectively) (Fig. 8a,b). The increased p21 ^CIP1 expression corresponded to the accumulation of G₂/M‐phase cells analysed previously by FACS analysis (1, 3) and the accumulation of cyclin B1 protein (2, 4). Inhibition of PKC or ERK activation during G₂ phase with Bis or U0126, respectively, inhibited PMA‐induced p21 ^CIP1 expression (Fig. 8a,b). These data suggest that elevated PKC and ERK activities delay G₂/M progression by increasing the expression of p21 ^CIP1 during G₂ phase.

Figure 8.

De novo synthesis of p21^CIP1 during G₂‐phase requires PKC and ERK activity. HeLa cells were synchronized at the G₁/S boundary, released, and harvested at the times indicated. (a) Immunoblot analysis of p21 ^CIP1 expression in control cells or cells treated with PMA, Bis, or PMA plus Bis as described in 1, 2. (b) Immunoblot of p21 ^CIP1 in control cells or cells treated with PMA, U0126, or PMA plus U0126 as described in 3, 4. (c) Immunoblot analysis of p21 ^CIP1 in untreated or cells treated with PMA in the presence or absence of cyclohexamide (10 µm) or MG115 (10 µg/mL) at 7 h after G₁/S‐phase release. The arrows and asterisks highlight the expression of p21 ^CIP1 at 9 or 11 h after G₁/S release in cells treated with PMA minus or plus cyclohexamide or MG115. Τhe expression of α‐tubulin (lower panel) is shown for a protein loading control in a, b, and c.

It was next investigated whether PMA‐induced expression of p21 ^CIP1 during G₂ phase was the result of increased protein synthesis or to chromosome stability. Synchronized cells were treated with cyclohexamide at 7.5 h after G₁/S release to block protein synthesis. After 30 min, cells were treated with PMA (8 h after G₁/S release) and harvested after an additional 1 or 3 h incubation (9 and 11 h post‐G₁/S‐phase release). As shown, cyclohexamide blocked basal and PMA‐induced p21 ^CIP1 expression at 9 and 11 h post‐G₁/S release (Fig. 8c). In contrast, inhibition of the proteasome and protein degradation with MG115 had little effect on PMA‐induced p21 ^CIP1 expression (Fig. 8c). MG115 treatment in the absence of PMA caused a small increase in p21 ^CIP1 expression, suggesting a role for the proteasome in p21 ^CIP1 turnover during G₂ and M phase (Fig. 8c). These data indicate that one mechanism for PMA‐mediated G₂‐phase arrest is through ERK‐mediated de novo synthesis of p21 ^CIP1 .

p21 ^CIP1 is required for PMA‐induced G₂‐phase arrest

To demonstrate that p21 ^CIP1 expression is required for PMA‐induced G₂‐phase arrest, cell cycle progression was monitored in HCT116 colorectal carcinoma cells lacking p21 ^CIP1 . FACS analysis showed that greater than 85% of both parental and p21 ^CIP1 knockout cells synchronized at the G₁/S‐phase boundary with 2N DNA content using excess thymidine (data not shown). Treatment with PMA at 6 h after release, when most cells have entered G₂/M‐phase, caused a delay in mitotic progression as measured by FACS in the parental but not the cells lacking p21 ^CIP1 (Fig. 9a,b). Examination of mitotic indices demonstrated that PMA induced a 2‐h delay in mitotic entry in parental cells but had no effect on mitotic entry in cells lacking p21 ^CIP1 (Fig. 9c,d). Although the data also show that cells lacking p21 ^CIP1 progress through G₂/M‐phase at different rates than the parental cells, these findings nevertheless indicate that p21 ^CIP1 is required for PMA‐induced G₂‐phase arrest.

Figure 9.

Expression of p21^CIP1 is required for PMA‐induced G₂‐phase arrest. HCT116 parental (a and c) or p21 ^CIP1 knockout cells (b and d) were synchronized at the G₁/S‐phase boundary, treated in the absence (closed bars) or presence (open bars) of 0.1 µm PMA at 6 h after release, and harvested at the times indicated after G₁/S release for FACS analysis of G₂/M‐phase cells (a and b) or mitotic indices (c and d).

A role for ERK‐mediated p21 ^CIP1 expression during G₂ phase in regulating chromosome stability

The last set of experiments described here examined the consequence of ERK activation during G₂ phase and p21 ^CIP1 expression, on hallmark characteristics of chromosome stability. Defects in mitosis and cytokinesis are manifested by cells that are binucleate and/or contain multiple centrosomes; both of which are indicators of chromosome instability (Saavedra et al. 1999; , Fenech 2002), among the cells described in these experiments. HeLa cells synchronized in G₂ were treated with EGF in the presence or absence of U0126, and then were incubated for 17 h to allow completion of one cell cycle, and then were processed for immunofluorescence. Approximately 15% of the cells treated with EGF were bi‐nucleate compared to approximately 3% in untreated cells (Fig. 10a,b). The increase in bi‐nucleate cells caused by EGF could be prevented with U0126 treatment (Fig. 10a). This response was unique to ERK activation during the G₂ phase, as asynchronous cells treated with EGF showed only a slight increase in bi‐nucleate cells as compared to G₂‐phase synchronized cells treated with EGF (Fig. 10c). Similarly, G₂‐phase cells treated with EGF also exhibited an increase in the number of cells containing greater than two centrosomes, which also was partially prevented with U0126 (Fig. 10d). These findings suggest that ERK activation during G₂ phase promotes cellular changes that are consistent with chromosome instability.

Figure 10.

ERK activation during G₂‐phase promotes hallmarks of chromosome instability; protective function of p21^CIP1 expression. (a) Cells synchronized at the G₁/S‐phase boundary were released for 7 h and treated with or without EGF (10 ng/mL) in the absence or presence of U0126 (10 µm). After 17 h, cells were immunostained for p62 nucleoporin as a marker of the nuclear envelope and counter‐stained with DAPI. The graph shows the percentage of bi‐nucleated cells under each condition. (b) Representative image of DAPI and p62 nucleoporin staining in control and EGF treated cells showing bi‐nucleated cells. (c) Percentage of bi‐nucleated cells in asynchronous or G₂‐phase synchronized cells treated for 17 h in the absence or presence of EGF. (d) Percentage of cells containing greater than two centrosomes in control, EGF or EGF plus U0126 treated cells as described for (a). HCT116 parental (e) or p21 ^CIP1 knockout cells (f) were synchronized at G₁/S phase, treated in the absence or presence of PMA at 6 h after release, and the percentage of bi‐nucleated cells were determined at the times indicated.

Finally, a function for p21 ^CIP1 expression during G₂ phase in regulating the generation of bi‐nucleate cells was examined in HCT116 parental and p21 ^CIP1 knockout cells. Synchronized cells were treated during G₂ phase in the absence or presence of PMA. At times after PMA treatment, cells were fixed and stained with DAPI for nuclear analysis. Similar to EGF, PMA treatment increased the number of bi‐nucleate cells in both parental and p21 ^CIP1 knockout cells (Fig. 10e,f). However, in the absence of p21 ^CIP1, there were substantially more bi‐nucleate cells in untreated or PMA‐treated conditions, as compared to the parental cells (Fig. 10e,f). The ratio of PMA to unstimulated cell was also expressed using the basal levels of bi‐nucleate cells in parental or p21 ^CIP1 knockout cells. When five time points for the unstimulated conditions were averaged, the p21 ^CIP1 knockout cells showed approximately 2.5‐fold more bi‐nucleate cells compared to the parental cells in the absence of PMA treatment. When this difference is taken into account, PMA still induced approximately threefold more bi‐nucleate cells in p21 ^CIP1 knockout cells compared to the parental cells at the 11‐h time point (data not shown). However, at the 25‐ or 33‐h time points, this analysis showed no difference in the PMA‐induced bi‐nucleate cells between the p21 ^CIP1 knockout cells and the parental cells (data not shown). Thus, although the magnitude in number of bi‐nucleate cells in p21 ^CIP1 knockout cells treated with or without PMA are greater than the parental cells (Fig. 10e,f), the fold increase as a result of PMA is only observed during the first cell cycle. Nevertheless, these data support a role for p21 ^CIP1 expression during G₂ phase in providing protection against a phenotype characteristic of chromosome instability.

DISCUSSION

Using synchronized cells, these studies demonstrate that activation of the ERK pathway during G₂ phase with PMA or EGF will initiate a rapid delay in G₂ to M‐phase progression. The ERK‐mediated delay during the G₂ phase appears to occur, in part, through the de novo synthesis of the cell‐cycle inhibitor p21 ^CIP1 in p53 defective cells. PMA or EGF dose–response assays indicated that some ERK activity above basal levels is tolerated during G₂ phase. However, once an ERK activity threshold was reached, then the delay in G₂‐phase progression could be observed. A potential consequence of activated ERK during G₂ phase is an increased number of bi‐nucleate cells that, in this case, are an indicator of defects in mitosis and cytokinesis and a characteristic of chromosome instability. Although a role for the ERK pathway during PMA or EGF‐induced G₁ or G₂‐phase arrest was implied previously (Kinzel et al. 1990; Kaszkin et al. 1991; Kosaka et al. 1996; Barboule et al. 1999), the current studies are to our knowledge, the first to show a link between ERK activity and the regulation of p21 ^CIP1 expression during G₂ phase.

The function of the ERK pathway in modulating G₂ and M‐phase transitions remains largely unknown. Recent studies using pharmacological inhibitors, catalytically inactive mutants of ERK pathway proteins, and RNA interference suggests that a basal ERK activity is important for G₂ and M‐phase progression in several transformed and non‐transformed cell lines (Abbott & Holt 1999; Wright et al. 1999; Hayne et al. 2000; Roberts et al. 2002; Liu et al. 2004). In addition, activation of the ERK pathway caused by over‐expression of constitutively active Ras, Raf‐1, or MEK1 mutants can induce a p53‐dependent cell cycle arrest and senescence in non‐transformed cells (Lin et al. 1998; Zhu et al. 1998; Ferbeyre et al. 2002). Moreover, over‐expression of the Mos kinase and subsequent ERK activation caused a p53‐dependent cell‐cycle arrest characterized by an increased number of bi‐nucleate cells (Fukasawa & Vande Woude 1997). Thus, although some basal level of ERK activity is needed during G₂/M phase, ERK activation above basal levels may also be detrimental at this time of the cell cycle.

Our studies used primarily p53 defective cells, which have suggested that G₂‐phase arrest mediated by ERK activation and p21 ^CIP1 induction is regulated through p53‐independent mechanisms. This is in agreement with others that used conditional activation of Raf kinase and ERK to cause a G₁‐phase arrest in mouse fibroblasts through p53‐independent induction of p21 ^CIP1 (Woods et al. 1997). Thus, p53‐independent mechanisms for cell cycle control as previously suggested (Taylor & Stark 2001) and p53‐dependent mechanisms appear to function in ERK‐mediated G₁ or G₂‐phase arrest.

One role for G₂‐phase arrest as a result of ERK activation may be to protect cells from signals that may interfere with mitotic transitions. A major function for the ERK pathway is to mediate transcription events in response to extracellular signals. However, the structural constraints imposed by condensed chromosomes result in transcription inhibition in mitotic cells. Thus, elevated ERK activity throughout the cell during G₂ phase might be sensed as an inappropriate activation of transcription events that would detract from the high energy requirements needed for the dynamic structural changes that occur during mitosis. To compensate, our findings suggest that ERK may negatively regulate the cell cycle by imposing a G₂‐phase checkpoint through induction of p21 ^CIP1 until ERK activity returns to levels that are compatible with proper mitotic transitions.

Increased expression of p21 ^CIP1, which is found in some cancer cells, may act to protect cells against apoptosis‐inducing agents (Gartel & Radhakrishnan 2005). Similarly, activation of the ERK pathway in cancer cells may be involved in the development of drug resistance (Davis et al. 2003). ERK‐induced G₂‐phase arrest following treatment with genotoxic agents, such as doxorubicin, or microtubule‐interfering drugs, such as paclitaxel, may provide a protective mechanism that prevents cells with extensive DNA damage from entering mitosis and generating additional damage that would initiate an apoptotic response. Thus, G₂‐phase arrest may allow the cells to repair DNA damage induced by chemotherapeutic drugs. As such, efforts towards simultaneous inhibition of ERK or other signalling pathways may be a more effective approach to treating drug‐resistant cancer cells. For example, simultaneous inhibition of the PKC, Chk1, and the ERK pathway with UCN‐01 and MEK1/2 inhibitors has been shown to be effective in killing leukaemia cells that harbour the BCR‐Abl fusion protein and are resistant to the anticancer drug ST1571, also known as Gleevec (Yu et al. 2002).

The role of other MAP kinase signalling pathways in regulating cell cycle progression through G₂ phase and mitosis may also play a role during genotoxic or other stress conditions. Activation of p38 MAP kinase was reported to mediate G₂‐phase arrest in response to genotoxic or osmotic stress (Bulavin et al. 2001; Dmitrieva et al. 2002; Hirose et al. 2003). Similarly, conditional activation of MEKK3, a kinase that mediates inflammatory responses, induces G₂‐phase arrest through a mechanism that is partially dependent on p38 MAP kinase activity (Ellinger‐Ziegelbauer et al. 1999; Garner et al. 2002). Several MAP kinases may mediate cell cycle arrest in the context of breast cancer cell lines treated with the chemotherapeutic drug paclitaxel, which causes G₂‐phase arrest and apoptosis in a manner that requires ERK and p38 MAP kinase activities but not a functional p53 protein (Bacus et al. 2001). However, in our studies, we have found no evidence that p38 MAP kinase is activated in G₂ or M‐phase cells treated with or without PMA (data not shown). This agrees with others who report that PMA is a potent activator of ERK but not p38 MAP kinase (Morton et al. 2004).

During DNA damage or microtubule disruption, the c‐Jun N‐terminal kinase (JNK) pathway may also promote G₂‐phase arrest and susceptibility to undergo apoptosis through a mechanism involving inhibition of cdc2 and the anti‐apoptotic protein Bcl‐2 (Yamamoto et al. 1999; Goss et al. 2003). Similar to ERK, a basal level of JNK activity may regulate proper progression through G₂ and M phase of the cell cycle. Recent studies suggest that inhibition of JNK activity with relatively non‐specific chemical inhibitors or by genetic knockout of MKK7, a JNK activator, causes a delay in G₂ to M‐phase progression (Mingo‐Sion et al. 2004; Wada et al. 2004). Although we have found no evidence that JNK proteins are activated in untreated or PMA‐treated synchronized HeLa cells during G₂ phase (data not shown), the three major MAP kinase family members may co‐ordinate cell cycle progression during G₂ phase in a manner that may be dependent on cell type and extracellular signal.