Autophagy activated after DNA damage or other stresses mitigates cellular damage by removing damaged proteins, lipids, and organelles. Activation of the master metabolic kinase AMPK enhances autophagy. Here we report that cyclin D1 restrains autophagy by modulating the activation of AMPK. In cell models of human breast cancer or in a cyclin D1–deficient model, we observed a cyclin D1–mediated reduction in AMPK activation. Mechanistic investigations showed that cyclin D1 inhibited mitochondrial function, promoted glycolysis, and reduced activation of AMPK (pT172), possibly through a mechanism that involves cyclin D1-Cdk4/Cdk6 phosphorylation of LKB1. Our findings suggest how AMPK activation by cyclin D1 may couple cell proliferation to energy homeostasis. Cancer Res; 77(13); 3391–405. ©2017 AACR.

Autophagy is an evolutionarily conserved catabolic process that degrades damaged proteins and organelles, recycling their biochemical components for energy production and other biosynthetic reactions. As a protective mechanism, autophagy allows the cell to survive in response to different stressors, such as nutrient deprivation, DNA damage, and low energy conditions (1). The core pathway governing autophagy is highly conserved (1). 5′ AMP-activated protein kinase (AMPK) is a sensor of cellular energy homeostasis and is activated by increased cellular adenosine mono- and diphosphate (AMP and ADP; ref. 2). Glucose deprivation induces phosphorylation of AMPK at Threonine 172 (T172), which restrains growth and switches cellular metabolism toward both catabolic and oxidative processes. The ability of AMPK to respond to altered AMP levels is dependent upon liver kinase B1 (LKB1), which phosphorylates AMPK (3). LKB1 was originally identified as a mutated tumor suppressor in Peutz–Jeghers syndrome. The discovery that LKB1 was the major kinase required for AMPK activation in response to metabolic stress provided a direct link between AMPK and cancer (4). LKB1 is constitutively active in cells (5) and its phosphorylation in response to oncogenic B-RAF compromises the ability of LKB1 to bind and activate AMPK (6).

The mitigation of cellular damage by autophagic removal of damaged proteins and lipidated organelles is important for tumor suppression in some circumstances, but promotes proliferation and maintains tumorigenesis in other circumstances. Thus, autophagy inhibits progression and delays tumor initiation, whereas it is required for tumor maintenance (7). The promotion of tumor growth by autophagy involves provision of energy substrates required for cellular proliferation. In contrast, there is a strong correlation between mitogenic signaling and the inhibition of autophagy (8). In a genome-wide screen, a large percentage of genes that negatively regulate autophagy were also involved in cellular growth and proliferation (9). Despite strong evidence for an association between mitogenic signaling in the restraint of autophagy, the molecular mechanisms remain poorly defined. In this regard LKB1-AMPK activates p27KIP1, a cyclin-dependent kinase inhibitor that leads to cell-cycle arrest (10). Furthermore, the induction of RAF-MEK-ERK negatively regulates AMPK, as ERK and Rsk phosphorylate LKB1, reducing AMPK activity (6). The phosphorylation of LKB1 at Serine 325 (S325) occurs within a motif that is predicted to serve as a substrate for proline-directed serine kinases, including the cell-cycle regulators belonging to the cyclin-dependent kinase family (11).

The CCND1 gene encodes cyclin D1, the regulatory subunit of the holoenzyme that phosphorylates and inactivates the retinoblastoma protein (Rb). CCND1 is overexpressed in a variety of malignancies and is increased in the early phase of breast cancer malignancy, including ductal carcinoma in situ (12). In recent studies of transgenic mice, cyclin D1–mediated promotion of breast tumorigenesis was kinase-independent, but autophagy restraint was kinase-dependent raising the question of the molecular mechanisms governing autophagy restraint (13, 14). The current studies confirmed these prior findings and extended these observations to identify the molecular mechanisms. The phosphorylation of LKB1 and consequent reduction in AMPK by cyclin D1 may serve to couple cellular proliferation to energy homeostasis.

Cell culture, vectors, and reagents

Cell lines were authenticated by examination of morphology and growth profile, and were mycoplasma free. MCF10A cells transformed with the oncogene NeuT (MCF10A-NeuT), Ha-Ras (MCF10-Ha-Ras), or v-Src (MCF10A-Src; MCF10A lines were generated in 2009 at Thomas Jefferson University, Philadelphia, PA), human embryonic kidney cells (HEK293T), cyclin D1 wild-type (cyclin D1+/+), and cyclin D1 knockout (cyclin D1−/−) mouse embryonic fibroblasts (MEF) and immortalized mouse embryonic fibroblasts (3T3) were described previously (15). MEF and 3T3 cells were generated in 2011 at Thomas Jefferson University, Philadelphia, PA.

The expression vectors encoding LKB1 with an N-terminal FLAG epitope (FLAG-LKB1) and the point mutant S325A (FLAG-LKB1-S325A) were described previously (6). The pSM2 vector containing shRNA for cyclin D1 and control vector were used as described previously (16). The pCDNA3.1-3X-FLAG expression vector encoding human cyclin D1 wild-type (cyclin D1WT), human cyclin D1 kinase-dead mutant (cyclin D1KE), and control vector were used as described previously (17).

MCF10A-NeuT cells were incubated with the Cdk4/6 inhibitor PD0332991 (catalog no. PZ0199, Sigma-Aldrich) at a working concentration of 500 nmol/L for 24 hours. Autophagy was induced by treating the cells either with 10 μmol/L of rapamycin (catalog no. R0395, Sigma-Aldrich) or Hank buffered salt solution (HBSS; catalog no. 21-020, Corning) for 2 hours. To activate the AMPK signaling pathway, MEFs were treated using metabolic stressors including ultraviolet (UV) or hydrogen peroxide (H2O2, 100 μmol/L).

Transgenic mice

Conditional cyclin D1fl/fl mice were a generous gift from Dr. Sicinski (Department of Genetics, Harvard Medical School, Cambridge, MA). C57BL/6J ROSA26-Cre-ERT2 mice were a gift from Dr. Streamson C. Chua Jr. (Department of Medicine, Albert Einstein College of Medicine, Bronx, NY). The appropriate Thomas Jefferson University institutional committee–approved protocols were followed when working with these mice. ROSA26-Cre-ERT2-cyclin D1fl/fl mice were generated by crossing cyclin D1fl/fl and ROSA26-Cre-ERT2 mice. Ccnd1 knockout mice were generated by a daily intraperitoneal injection of tamoxifen (1 mg/200 μL) for 5 days, followed by a washout period of 4 weeks. Mammary gland tissues were then collected. Wild-type Cre-ERT2 mice were used as control.

Immunofluorescence

Autophagy activation was measured by immunodetection of microtubule-associated protein light-chain 3 (LC3) puncta using LC3-A/B antibody (catalog no. 4108, Cell Signaling Technology). α-Tubulin (catalog no. T6199, Sigma-Aldrich) was used as a reference marker for cellular area. The amount of LC3 puncta was measured with ImageJ and normalized by cell area. The autofluorescent agent monodansylcadaverine (Sigma; ref. 18) was used to measure autophagy activation in 96-well plates. Cells were plated at a density of 2 × 104 cells/cm2 one day before treatment and were incubated overnight at 37°C. After treatment, as described above, cells were incubated for 10 minutes with 0.05 mmol/L monodansylcadaverine at 37°C and were then observed with a fluorescence plate reader using a UV filter. Autophagic flux was assessed using LC3-GFP in the presence of chloroquine.

To assess autophagy flux, cyclin D1+/+ and cyclin D1−/− 3T3 cells (1 × 106) were transfected by electroporation (Nucleofector 2b Device, Program NIH 3T3 U-030; Lonza) with the mCherry-EGFP-LC3 dual reporter (Addgene, #22418). Analysis of the labeled LC3 puncta was conducted by confocal microscopy (Nikon Cl PLUS) and quantitated using ImageJ (19).

Western blot and immunoprecipitation Western blot analyses

Whole-cell lysates (50 μg) were separated by SDS-PAGE and the proteins transferred to a nitrocellulose membrane for Western blotting as described previously (20). Specific primary antibodies were prepared according to the manufacturer's instructions; Cyclin D1 (catalog no. sc-753, clone H295, Santa Cruz Biotechnology), phospho Rb-S780 (catalog no. 9307S, Cell Signaling Technology), LC3-A/B (catalog no. 4108, Cell Signaling Technology), AMPK (catalog no. 2532S, Cell Signaling Technology), phosphorylated AMPK T172 (catalog no. 2531S, Cell Signaling Technology), Bnip3 (catalog. no. 3485-1, Epitomics), and vinculin (catalog no. V9131, Sigma-Aldrich), which was used as an internal protein-loading control. The LKB1 phospho-serine 325 (pS325) antibody was described previously (6). Anti-rabbit (catalog no. sc-2004, Santa Cruz Biotechnology) or mouse (catalog no. sc-2005, Santa Cruz Biotechnology) secondary HRP–conjugated antibodies were diluted at 1:3,000.

Immunoprecipitation kinase assays

HEK293T cells were cotransfected with expression vectors encoding FLAG-LKB1 and either cyclin D1WT or cyclin D1KE. The lysates were prepared using cellular lysis buffer (HEPES pH = 7.0, 150 mmol/L NaCl 1% NP-40, 1 mmol/L Na3VO4, and protease inhibitor cocktail). Protein concentrations were normalized using the Bradford assay. For immunoprecipitation, cell lysates were incubated with primary antibodies of anti-FLAG M2 (catalog no. F3165, Sigma-Aldrich) affinity gel overnight at 4°C, followed by incubation with protein A/G sepharose for 1 hour at 4°C. Immunoprecipitated samples were incubated in kinase assay buffer in the presence of γ-32P dATP for 20 minutes at 37°C. After labeling, samples were run on a 10% SDS-PAGE gel. The gel was dried and exposed to an autoradiography film (21).

Statistical analysis

The statistical analysis was performed using Prism 4 software (GraphPad Software). Differences were considered statistically significant with P values <0.05.

Measurements of oxygen consumption rate and extracellular acidification rate

A Seahorse Bioscience XF24-3 Extracellular Flux Analyzer was used to measure the rate change of dissolved O2 and pH of medium immediately surrounding adherent cells cultured in a XF24 V7 cell culture microplate (Seahorse Bioscience). Measurements of oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) were performed after equilibration in assay medium for 1 hour. Measurements of O2 concentration and pH were made periodically over 4 minutes, and OCR and ECAR were obtained from the slopes of concentration change in these parameters versus time.

Lactate assay, ATP assay, and NAD+/NADH ratio

Lactate concentration in cell culture media was assayed using BioAssay Systems' EnzyChromTM lactate assay kit. Total cellular ATP was assayed on the basis of the manufacturers' guidelines (Molecular Probes, Invitrogen). The NAD+/NADH ratio was quantitated per the manufacturer's guidelines (BioVision) through a reaction that specifically detects NADH and NAD+, but not NADP nor NADPH.

Oxygen consumption assay

OCRs in cells were conducted in an Oxygen Biosensor System (BD Biosciences). Oxygen consumption rates were measured over 3 hours every 15 minutes using a fluorescent plate reader.

Quantitation of mitochondrial morphology

The morphologic analysis of mitochondria in live MEF was performed as described previously (22), by using confocal Z-stack maximal projections of TMRE-loaded mitochondria and running the analysis using ImageJ.

FACS analysis

For FACS analysis of mitochondria, MEFs were loaded with Vybrant DyeCycle Violet according to manufacturer's protocol (LifeScience) and 10 μmol/L JC-1 dye (Molecular Probes) for 30 minutes at 37°C. Green and red fluorescence components from JC-1 staining were measured and segregated based on the cell-cycle phase.

Mitochondrial FRAP assay

Live MEFs were stained with tetramethylrhodamine methyl ester (TMRM; 50 nmol/L) in DMEM for 30 minutes at 37°C and mitochondrial dynamics were analyzed by FRAP as published previously (23) using a Zeiss inverted confocal microscope equipped with a 63× Oil objective. Ethacrinic acid (50 μg/mL) and FCCP (5 μmol/L) were used as positive and negative controls for the analysis. Recovery of fluorescence was monitored for 120 seconds. Three different clones were used for each genotype, and for each of them 3 to 5 cells were analyzed. High resolution images of TMRM-loaded mitochondria were used for quantification of mitochondrial morphology using ImageJ.

Endogenous cyclin D1 restrains oncogene-induced autophagy

Cyclin D1 abundance is induced during oncogenic transduction of immortalized human breast MCF10A epithelial cells and is a direct transcriptional target of NeuT, v-Src, and Ha-Ras (24–26). To determine whether cyclin D1 regulates autophagy induced by distinct oncogenes, isogenic human breast cancer cell lines were assessed. We first determined the role of endogenous cyclin D1 in regulating basal autophagy by using short hairpin RNA (shRNA) against cyclin D1. We verified that oncogene transformation of MCF10A-ErbB2 (NeuT) and MCF10A-Ha-Ras induced autophagy compared with control (ctrl) parental MCF10A cells (Fig. 1A). Both NeuT (P = 0.012) and Ha-Ras (P = 0.002) induced autophagy based on Becn1 abundance (Fig. 1B). In MCF10A-NeuT cells, cyclin D1 shRNA reduced the abundance of cyclin D1 by >80% (Fig. 1C). We determined the basal autophagy status by detecting both isoforms of microtubule-associated protein 1 light-chain 3 (LC3) I and II, as the lipidated form (LC3-II), stably associates with the autophagosomes, serving as a marker of autophagy induction (Fig. 1C and D; P = 0.021; ref. 27). Next, we deployed MEFs and 3T3 cells derived from cyclin D1−/− and cyclin D1+/+mice. LC3-II was increased in cyclin D1/-MEF (P = 0.013) and 3T3 cells (P = 0.014) compared with cyclin D1+/+cells (Fig. 1E–H). We verified that the mechanism of cyclin D1 inhibition of autophagy involved cyclin-dependent kinase activity by treating cells with a cdk4/6 inhibitor (PD0332991 - palbociclib). PD0332991 treatment of MCF10A-NeuT (Fig. 1I and J) and MCF10A-Ha-Ras (Fig. 1K and L) significantly reduced the phosphorylation of Rb at serine 780 (pRb S780; cdk4/6 phosphorylation site) with a commensurate increase in LC3-II form (P = 0.016 and P = 0.006, respectively) and Bnip3.

MCF10A-NeuT and MCF10A-v-Src cells were treated with distinct autophagy inducers (Fig. 2A and B). Cyclin D1 shRNA enhanced basal autophagy in MCF10A-NeuT cells approximately 2-fold and in MCF10A-v-Src cells approximately 5-fold (P < 0.001) (Fig. 2C–E). Cyclin D1 shRNA enhanced autophagy in HBSS-treated MCF10A-NeuT cells approximately 1.7-fold and in MCF10A-v-Src cells approximately 3.5-fold (P < 0.001; Fig. 2C–E) and cyclin D1 shRNA enhanced autophagy in rapamycin-treated MCF10A-NeuT cells approximately 1.8-fold and in MCF10A-v-Src cells approximately 5-fold (P < 0.001; Fig. 2C and E). Collectively, these data demonstrate that endogenous cyclin D1 restrains autophagy in breast cancer cells transformed by ErbB2, Ha-Ras, or oncogenic v-Src.

Cyclin D1 restrains stress-induced AMPK activation

AMPK is a central mediator of cellular adaption to metabolic flux and promotes autophagy. AMPK is activated upon depletion of nutrients by binding AMP and by LKB1 inducing phosphorylation at threonine 172 (T172). To determine whether cyclin D1–mediated restraint of autophagy involves the inhibition of AMPK, we examined AMPK signaling in tissue culture and in vivo upon cyclin D1 depletion. We deployed MCF10A-NeuT and MCF10A-Ha-Ras cells treated with cyclin D1 shRNA. Cyclin D1 depletion correlated with a significant increase in the phosphorylation level of AMPK at T172 in both MCF10A-Ha-Ras (P < 0.001; Fig. 3A and B) and MCF10A-NeuT (P < 0.001; Fig. 3C and D).

To determine whether cyclin D1 restrains AMPK activation by cellular stressors, cyclin D1−/− and cyclin D1+/+MEFs were treated with the cellular stressors H2O2 and UV. Cellular stress activated pAMPK (T172), which was further enhanced by cyclin D1 knockdown in both treatments, UV and H2O2 (P = 0.023 and P = 0.004, respectively; Fig. 3E and F). Total AMPK kinase was unchanged by cyclin D1 in MCF10A-NeuT, MCF10A-Ha-Ras, and MEFs. Total AMPK remained unchanged upon bioenergetic stress (Fig. 3). In addition, we determined that AMPK phosphorylation at T172 was inhibited by cyclin D1 in 3T3 cells (Fig. 3G and H) commensurate with a reduction in lipidated LC3 with AMPK1α2α/-3T3 cells used as a control.

AMPK is activated following cell stress and is required under stress to promote cell viability (28). We therefore tested the role of cyclin D1 in glucose stress. Cyclin D1 sensitized MEF to glucose stress, as growth rates of cyclin D1+/+MEFs versus cyclin D1/-MEFs were compromised at glucose concentrations below 4.5 g/L, as shown in Supplementary Fig. S1A (P = 1E–05 and P = 0.05, respectively) and 1B (cyclin D1/-vs. cyclin D1+/+at 2 g/L of glucose P = 3.5E–06, at 1 g/L of glucose P = 4.2E–08 and minus glucose P = 8E–05).

Endogenous cyclin D1 restrains autophagy in the mammary gland

To determine whether cyclin D1 restrains autophagy in vivo, we examined transgenic mice that delete the Ccnd1 gene in the adult mouse (ROSA26-Cre-ERT2-Ccnd1fl/fl mice; Fig. 4A). Animals were treated with tamoxifen to induce Cre expression, and analysis was conducted 4 weeks later to ensure washout of tamoxifen. Mammary gland tissues of these mice were assessed using IHC for autophagy (Becn1 and Bnip3) and AMPK activity (pAMPK T172; Fig. 4B). Cyclin D1 was detected in the mammary epithelium and fibroblasts of the vehicle-treated bi-transgenic mice, but was not detectable in animals that had been treated with tamoxifen (Fig. 4B). pAMPK at T172 was increased in the cyclin D1/- mammary epithelial cells (P < 0.05; Fig. 4C). The autophagy markers, Becn1 and Bnip3 were increased in the cyclin D1/- mammary gland tissue (P < 0.05 and P < 0.001, respectively; Fig. 4D and E). IHC was conducted on murine mammary gland–targeted cyclin D1 transgenic mice (rtTA/CCND1WT; Fig. 4F and G). Transient expression of cyclin D1 (17 days), through doxycycline induction of a tet-regulated cyclin D1 cDNA cassette directed to the murine mammary gland, induced cyclin D1 abundance, decreased pAMPK T172 (a correlate of AMPK activity), and reduced autophagy as attested by IHC showing reduced Becn1 and Bnip3 (P < 0.001 and P < 0.01, respectively; Fig. 4H–J).

Cyclin D1 restrains chloroquine-induced autophagic cargo flux

We next conducted analysis of autophagy cargo flux using endogenous LC3. Prior to engaging in the assessment of cyclin D1 restraint of autophagic cargo flux in the model cell lines, we examined the autophagic response via treatment with chloroquine. Chloroquine inhibition of autophagy measured by lipidated LC3 in MCF10A-NeuT (Supplementary Fig. S2A and S2B), cyclin D1+/+MEFs, and cyclin D1+/+3T3 cells was induced by 5-fold (P < 0.001), 1.7-fold (P = 0.007), and 1.7-fold (P < 0.001), respectively (Fig. 5A–D). We had previously confirmed that retroviral transduction of fluorescent protein–tagged LC3 relocates to the autophagosome membrane upon induction of autophagy. The induction of autophagy by culturing cells in nutrition-free balanced salt solution HBSS (Fig. 5E) was quantitated using confocal images and the corresponding general intensity surface spots were quantitated as mean data ± SEM (Fig. 5F). A comparison was made between cyclin D1−/− and cyclin D1+/+ MEFs. The redistribution of the LC3-RFP to autophagosomes to form puncta was quantitated. In response to HBSS, the relative abundance of LC3 in autophagosomes characterized by puncta was increased, with a further increase in cyclin D1−/− compared with cyclin D1+/+ MEF (P = 0.004). An increase in the relative amount of LC3-II can be due to either an increase in autophagy or a decrease in the degradation of autophagic intermediates. Autophagic activity may therefore reflect changes in autophagic flux. Flux can also be measured by blocking lysosomal degradation of autophagosomes using chloroquine, resulting in a buildup of autophagic intermediates. After chloroquine treatment, GFP-LC3 puncta increased 7-fold, which was increased 10-fold in cyclin D1−/− cells (Fig. 5F; P = 3.7E–06). In addition, we assayed autophagic flux by analyzing the ratio of LC3-II to p62. As p62 levels increase when autophagy is inhibited, and decreased when autophagy is induced, p62 can be used as a marker to study autophagic flux. Autophagic flux in 3T3 cells derived from cyclin D1−/− (p62/LC3II ratio = 15.7) compared with cyclin D1+/+mice (p62/LC3II ratio = 0.41; P = 0.0014) showed a significantly greater LC3-II/p62 ratio (Fig. 5G and H). These, findings are consistent with cyclin D1 restraining autophagic cargo flux.

To define further the role of cyclin D1 in autophagic cargo flux, we deployed a Autophagy Tandem Sensor to visualize maturation of the autophagosome to the lysosome. By combining an acid-sensitive GFP with an acid-insensitive RFP, the change from autophagosome to autolysosome can be visualized by imaging the specific loss of the GFP fluorescence, leaving only red fluorescence allowing for dynamic visualization of the autophagic process. The autophagy flux in both 3T3 cyclin D1+/+ and cyclin D1−/− cells was assessed using a dual report mCherry-EGFP-LC3. At basal condition, the knocking down of cyclin D1 in 3T3 cells shows an increase of both signals, EGFP (P < 0.037) and mCherry (P < 0.025) compared with 3T3 cyclin D1+/+ (Fig. 5I). Moreover, in 3T3 cells′ cyclin D1−/−, there is an increase of autophagosomes labeled exclusively with mCherry signal (Fig. 5J). These data suggest that autophagy flux is restrained by cyclin D1.

Deletion of cyclin D1 leads to a gain-of-function phenotype, resulting in increased mitochondrial respiration

AMPK is a central cellular mediator of the nutrient stress response. Under conditions of nutrient flux, AMPK inhibits anabolic processes including lipogenesis, protein metabolism, and cell growth and activates catabolic processes that increase cellular ATP (β-fatty acid oxidation, glycolysis, β-fatty acid oxidation, mitochondrial biogenesis and autophagy; ref. 29). A metabolic analysis was conducted to determine whether cyclin D1 inhibition of AMPK recapitulated the metabolic profile of AMPK inhibition. OCRs were determined using an oxygen biosensor system (Fig. 6A). Endogenous cyclin D1 blunted OCRs (cyclin D1+/+versus cyclin D1/MEFs, P < 0.001). Reflecting increased mitochondrial oxidation rates, the NAD/NADH ratio was increased in cyclin D1−/− MEFs (Fig. 6B; P = 0.04). Cellular ATP levels (Fig. 6C; P = 3.7E–14) and increased lactate secretion was observed in cyclin D1−/− MEFs (Fig. 6D; P = 0.039).

Next, we deployed the Seahorse Extracellular Flux Analyzer (XF) that measures oxygen consumption (OCR) and glycolytic flux (ECAR) in cyclin D1+/+MEFs and cyclin D1/-MEFs. Multiple independent MEF preparations were analyzed using a standard bioenergetics protocol (basal, oligomycin, FCCP, and antimycin A; BOFA). Glycolytic flux measured through external acidification rates (ECAR) was decreased by cyclin D1 (Fig. 5E and F). OCRs were significantly increased in cyclin D1−/− MEFs under basal conditions, indicating a gain-of-function phenotype in cyclin D1−/− MEF (Fig. 6G and H; P = 9.1E–15).

Following oligomycin addition, the maximal respiratory rate was determined following uncoupling of mitochondrial metabolism by FCCP. Cyclin D1−/− MEF showed increased mitochondrial oxygen consumption compared with cyclin D1+/+MEF cells and significantly increased OCR following mitochondrial uncoupling with FCCP (Fig. 6I; P = 3.67E–06), indicative of greater mitochondrial reserve in the cyclin D1−/− MEF. The OCR due to nonmitochondrial sources, determined as OCR in the presence of antinomycin A, was unchanged.

We next determined the ability of cyclin D1 to oxidize exogenously added fatty acids. Substrate utilization of cyclin D1−/− versus cyclin D1+/+MEFs was determined with palmitate-BSA in a fatty acid oxidation protocol on the Seahorse Extracellular Flux (XF) Analyzer. OCRs were 2-fold higher in the cyclin D1−/− MEF compared with cyclin D1+/+MEF (Fig. 6J and K; P = 9.3E–05).

Cyclin D1 inactivation reduces mitochondrial interconnectivity and mitochondrial membrane potential

Mitochondrial interconnectivity can be measured by fluorescent recovery after photobleaching (FRAP). To determine whether the mitochondrial network of cyclin D1−/− MEFs demonstrated increased connectivity and therefore a more efficient capacity for substrate oxidation, we conducted FRAP on cyclin D1−/− versus cyclin D1+/+MEF cells alongside AMPK1α−/− fibroblasts as a control (Supplementary Fig. S3). We treated cells with FCCP, a protonophore (H+ ionophore) and uncoupler of oxidative phosphorylation in mitochondria (fragmented mitochondrion; negative control), or ethacrynic acid, a potent inducer of mitochondrial fusion (interconnected mitochondrion; positive control) as controls, and compared cyclin D1−/− versus cyclin D1+/+MEF (Supplementary Fig. S3A and S3B). Cells were loaded with the fluorescent tracer, TMRM, and then a mask was used to photobleach a small region of the mitochondrion. The fluorescent recovery rate was timed (Supplementary Fig. S3A and S3B). In cyclin D1−/− MEFs, the fluorescent recovery was significantly faster, indicating increased mitochondrial connectivity. The AMPK1α2α−/− fibroblasts demonstrated increased recovery compared with cyclin D1/-MEFs, implying that in AMPK1α2α−/− fibroblasts, the mitochondria have increased connectivity (Supplementary Fig. S3C)

To determine whether cyclin D1 expression reduces mitochondrial area and number, quantification was conducted on MEF. Endogenous cyclin D1 inhibited both mitochondrial area and number (Supplementary Fig. S4A and S4B; P = 0.015 and P = 0.003, respectively). Reduced AMPK levels correlates with a reduction in mitochondrial membrane potential and mass (30). Mitochondrial membrane potential (red component of JC1) and mitochondrial mass (green component of JC1) were inhibited by cyclin D1 at each phase of the cell cycle (Supplementary Fig. S4C; mitochondrial mass at G0–G1, P = 8.4E–04; S-phase, P = 3.4E–05; and G2–M, P = 0.003; and Supplementary Fig. S4D; mitochondrial membrane potential at G0–G1, P = 0.001; S-phase, P = 0.0003; and G2–M, P = 0.0039).

Cyclin D1–associated kinase phosphorylates LKB1 at S325A

Herein, cyclin D1 restrained autophagy and inhibited AMPK phosphorylation. LKB1 binds to phosphorylated AMPK at threonine 172 (T172). Phosphorylation of LKB1 at Serine 325 by ERK and Rsk compromises its ability to bind and activate AMPK (6). As the LKB1 phosphorylation site resembles a consensus for a proline-directed serine/threonine kinase, we considered the possibility that LKB1 served as a substrate for cyclin D1/cdk4/6. HEK293T cells were transfected with a mammalian expression vector encoding cyclin D1 with an N-terminal FLAG epitope. Immunoprecipitation of cell extracts using the anti-FLAG antibody to precipitate cyclin D1, coprecipitated cdk4 and LKB1 (Fig. 7A). To determine whether LKB1 serves as a substrate for cyclin D1/cdk4, using GST-LKB1 as a substrate, we conducted a cyclin D1-IP kinase assay. Cells were transfected with an expression vector encoding cyclin D1, or a kinase-defective cyclin D1 cDNA (cyclinD1KE; Fig. 7B). Expression of cyclin D1 enhanced LKB1 phosphorylation as assessed by incorporation of γ-32P into LKB1. Cells transduced with a kinase-defective mutant of cyclin D1 did not significantly enhance LKB1 phosphorylation (Fig. 7B and C). Western blot analyses of cells transfected with an expression vector encoding LKB1WT or a point mutant at Serine 325 (LKB1S325A) were immunoblotted with an antibody to phosphorylated LKB1 (S325A). We demonstrated the presence of the phosphorylated form of LKB1, which was reduced by the addition of the cdk4/6 inhibitor PD0322991 and abolished by point mutation of LKB1 at S325A (Fig. 7D).

To determine whether phosphorylation of LKB1 at S325 by cyclin D1 cdk4/6 contributes to autophagic restraint, we deployed a point mutation of LKB1 S325A and conducted Western blots for activated AMPK and autophagic markers. Expression of LKB1 S325A in wild-type MEF abrogated cyclin D1 restraint of autophagy compared with MEF-expressing LKB1 WT (Fig. 7E and F). To determine whether cyclin D1/cdk4/6 phosphorylation of LKB1 at S325 also contributed to mitochondrial function, we deployed the point mutation of LKB1 S325A and conducted ECAR and OCR analysis using the Seahorse Extracellular Flux Analyzer (XF). The expression of LKB1 S325A in cyclin D1+/+ MEF significantly increased OCR (2-fold; P = 0.02; Fig. 7G) and increased ECAR (2 fold; P = 0.052; Fig. 7H).

Cyclin D1 restrains autophagy

The current studies reveal that endogenous cyclin D1 restrains autophagy demonstrated by using shRNA to cyclin D1, cyclin D1 kinase inhibition, or cyclin D1 gene deletion. The restraint of autophagy was demonstrated by the reduction in LC3-II/LC3-I by Western blot analysis and the formation of autophagic puncta visualized using an LC3-GFP fusion protein. Endogenous cyclin D1 restrained the induction of autophagy induced by either ErbB2, Ha-Ras, or v-Src, or by the cellular stress of nutrient deprivation or by rapamycin treatment. The current findings that cyclin D1, which is induced by a variety of mitogenic stimuli, restrains autophagy is consistent with prior studies on mitogenic stimuli and autophagy, as there is a strong correlation between mitogenic signaling and the inhibition of autophagy (8). In prior studies, the mammary gland of MMTV-ErbB2-cyclin D1KE/KE knock-in mice showed enhanced mammary epithelial cell cellular proliferation similar to MMTV-ErbB2-cyclin D1WT/WT knock-in mice, but increased autophagy, suggesting cyclin D1 restrained basal autophagy in a kinase-dependent manner (14). The current studies extend these findings by demonstrating the mechanism; cyclin D1 phosphorylates LKB1 at S325 to inactivate AMPK and thereby restrain autophagy. The demonstration that the cell-cycle control protein cyclin D1 controls autophagy via phosphorylation of LKB1 brings together two of the important hallmarks of cancer, deregulation of cellular energetics, and sustained proliferative signaling (31).

Cyclin D1 inhibits AMPK

Cyclin D1 reduced AMPK activity in tissue culture and in the mammary gland in vivo. Increased phosphorylation of AMPK (T172) was demonstrated upon reduction of cyclin D1 in human breast cancer cell lines, MEFs, and in the tissue of mammary epithelial cell–targeted inducible cyclin D1 deletion mice. Endogenous cyclin D1 restrained AMPK induction by the cellular stresses of nutrient deprivation, reactive oxygen species (H2O2), and ultraviolet irradiation. AMPK is downregulated in tumors compared with normal epithelium through an unknown mechanism (29). Cyclin D1 is overexpressed in the majority of human breast tumors and may contribute to the downregulation of AMPK in tumors. AMPK is known to induce (2) and cyclin D1 is known to restrain mitochondrial biogenesis and thereby increase cytosolic glycolysis (21, 32).

In the current studies, endogenous cyclin D1 was shown to enhance glycolytic flux, blunt OCR, and to reduce OCR when MEFs were supplemented with palmitate. Cyclin D1 reduced mitochondrial area and number and reduced mitochondrial connectivity. Cyclin D1 increased lactate secretion and reduced oxidative phosphorylation in the mitochondria. AMPK activation increases shuttling of glycolytic pyruvate into the mitochondria. The decrease in lactate secretion in the cyclin D1−/− MEF is indicative of elevated shuttling of glycolytic pyruvate into the mitochondria. Introducing the LKB1 325A mutant abrogates the suppression of glycolysis and oxygen consumption evidenced by increased ECAR and OCR in cyclin D1+/+LKB1S325A cells compared with cyclin D1+/+LKB1WT.

AMPK is known to be involved in mitochondrial biogenesis in hepatocytes (33, 34). Relevant to the finding here, both mitochondrial respiration and basal level of ATPs were significantly lower in hepatocytes isolated from a liver-specific AMPKα1α−/− mouse compared with control mice (35). Reduced AMPK activity may be an important factor in the reduced mitochondrial function. AMPK has been reported to induce cellular glycolysis under a number of cellular stress events (36–38). These findings are consistent with our findings that cyclin D1 inhibits mitochondrial function and reduces glycolytic flux through regulating the LKB1/AMPK signaling axis.

Cyclin D1 phosphorylates LKB1 requiring S325A

To examine the mechanism by which cyclin D1 inhibited AMPK, we considered the role of the upstream kinase LKB1. LKB1 itself is constitutively active in cells (5, 6); however, phosphorylation by Raf reduced AMPK phosphorylation (6). The MAPKs, ERK and Rsk, phosphorylate Ser325 inactivating LKB1-dependent AMPK activity (6). The phosphorylation of LKB1 interferes with the ability of LKB1 to phosphorylate and activate AMPK (6). Herein, a cyclin D1 complex, including cdk4/6, associated with LKB1. Transfection of HEK293T cells with cyclin D1 enhanced LKB1 phosphorylation 3-fold; however, a kinase-dead mutant of cyclin D1 failed to induce LKB1 phosphorylation. By Scan Site analysis, LKB1 Ser325 was a candidate site for proline-dependent serine/threonine kinases and was required for phosphorylation in cyclin D1 immune kinase assays. Finally, treatment of cells with the cdk inhibitor PD0322991 reduced LKB1 phosphorylation. The binding and inactivation of the LKB1 tumor suppressor by cyclin D1 (Supplementary Fig. S5) is consistent with the known role of cyclin D1 to bind and inactivate other tumor suppressors (pRb, p130, p107, BRCA1; refs. 39, 40). Endogenous cyclin D1 phosphorylated LKB1 and repressed the induction of AMPK associated with cellular stress. It has been suggested that activation of AMPK may explain the tumor suppressor effects of LKB1, as AMPK is the only downstream target of LKB1 known to cause inhibition of cell growth (2).

What might be the clinical significance of these findings?

The finding that cyclin D1 restrains autophagy may be important in therapeutic strategies depending upon the role of autophagy in tumor progression and therapy resistance. The removal of damaged proteins by autophagy may be important for both cell survival and tumor suppression (8). Autophagy has a cellular context–dependent role in cancer and may also vary depending on the stage of tumorigenesis. Evidence exists for both a tumor suppressor and for a tumor-promoting role of autophagy. Evidence for a tumor-suppressive role includes findings that autophagy may inhibit tumorigenesis by limiting necrosis and inflammation, which are thought to be sites that generate cancer-causing mutations (41). Defects in autophagy are associated with increased ROS production (18) and tumor development and several oncogenic pathways (PI3K mutations, AKT amplification, Pten loss) decrease autophagy (42). The tumor suppressor p53 induces autophagy (43). Furthermore, Beclin 1, the mammalian homolog of yeast ATG6, functions as a tumor suppressor with haploinsufficiency (44).

In some circumstances, autophagy functions to maintain tumors, providing energy substrates required for cellular growth (45). Autophagy may promote tumor development later in the tumorigenic process by providing energy substrates during periods of nutrient limitation (46). Autophagy is required for the survival of tumor cells in hypoxic tumor regions. Oncogenic RAS transformation upregulates basal autophagy required for the maintenance and progression of tumorigenesis (46). Engineered mouse models have also demonstrated a protumorigenic role for autophagy. Deletion of Atg7 in KrasG12D or BrafV600E non–small cell lung cancer (NSCLC) in adult mice suppresses tumor cell proliferation. Autophagy promotes tumorigenesis by mitigating Trp53 tumor suppression (47, 48). Using temporally controlled genetic ablation of the autophagy gene Atg7, recent studies suggest the function of autophagy may change with time (7). In the setting of cancer, 5 weeks of acute autophagy ablation converted established lung adenocarcinomas to oncocytomas and blocked cell proliferation.

The current studies are consistent with, and extend, the repertoire of cyclin D1–mediated functions. Autophagy limits genomic instability and the inhibition of this autophagic function increases genomic instability (49). The inhibition of autophagy by cyclin D1 is therefore consistent with the known effects of cyclin D1 to induce chromosomal instability (13, 20). The current studies extend the reported cyclin D1 functions that may contribute to the collaborative oncogenic function of cyclin D1.

No potential conflicts of interest were disclosed.

The Institute specifically disclaims responsibility for any data analysis, interpretations or conclusions.

Conception and design: M.C. Casimiro, G. Di Sante, C. Wang, P.E. Scherer, R.G. Pestell

Development of methodology: M.C. Casimiro, G. Di Sante, T. Pestell, X. Jiao, C. Wang, B. Zheng, R.G. Pestell

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): M.C. Casimiro, G. Di Sante, A. Di Rocco, E. Loro, C. Pupo, T. Pestell, S. Bisetto, M.A. Velasco-Velázquez, X. Jiao, Z. Li, C.M. Kusminski, E.L. Seifert, C. Wang, D. Ly, R.G. Pestell

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): M.C. Casimiro, E. Loro, M.A. Velasco-Velázquez, E.L. Seifert, D. Ly, R.G. Pestell

Writing, review, and/or revision of the manuscript: M.C. Casimiro, G. Di Sante, P.E. Scherer, R.G. Pestell

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): C.-H. Shen

Study supervision: M.C. Casimiro, G. Di Sante, R.G. Pestell

This work was supported in part by NIH grants (1R01CA137494, R01CA132115, R01CA086072 to R.G. Pestell), the Sidney Kimmel Cancer Center NIH Cancer Center Core grant P30CA056036 to R.G. Pestell, generous grants from The Breast Cancer Research Foundation to R.G. Pestell, and the Dr. Ralph and Marian C. Falk Medical Research Trust to R.G. Pestell, a grant from the Pennsylvania Department of Health to R.G. Pestell, and an American-Italian Cancer Foundation Post-doctoral Research Fellowship to G. Di Sante.

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

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