Cluster of differentiation 44 (CD44) is a transmembrane glycoprotein that has been identified as a cancer stem cell marker in various cancer cells. Although many studies have focused on CD44 as a cancer stem cell marker, its effect on cancer cell metabolism remains unclear. To investigate the role of CD44 on cancer cell metabolism, we established CD44 knock-down cells via retroviral delivery of shRNA against CD44 in human breast cancer cells. Silencing of CD44 decreased the glycolytic phenotype of cancer cells, affecting glucose uptake, ATP production, and lactate production. We also found that ablation of the CD44-induced lactate dehydrogenase (LDH) isoenzyme results in a shift to LDH1 due to LDHA down-regulation and LDHB up-regulation, implying the importance of LDH isoenzyme modulation on cancer metabolism. The expression of glycolysis-related proteins including hypoxia inducible factor-1α (HIF-1α) and LDHA was decreased by CD44 silencing. These effects were due to the up-regulation of liver kinase B1 (LKB1)/AMP-activated protein kinase (AMPK)α activity by reduction in c-Src and Akt activity in CD44 knock-down cells. Finally, induction of LKB1/AMPKα activity blocked the expression of HIF-1α and its target gene, LDHA. Inversely, LDHB expression was repressed by HIF-1α. Collectively, these results indicate that the CD44 silencing-induced metabolic shift is mediated by the regulation of c-Src/Akt/LKB1/AMPKα/HIF-1α signaling in human breast cancer cells.
Most normal cells produce energy through mitochondrial oxidative phosphorylation (OXPHOS). Since OXPHOS is more efficient in the generation of adenosine triphosphate (ATP) than in glycolysis, OXPHOS is activated and glycolysis is inhibited in the presence of oxygen. This mode of metabolism is called the ‘Pasteur effect’ . However, most cancer cells depend on aerobic glycolysis for ATP generation. This unique metabolism, known as the ‘Warburg effect’, can produce ATP more rapidly than OXPHOS. Therefore, aerobic glycolysis supports rapid cell proliferation, migration, and invasion of tumor cells [2–5].
Cancer cells constantly uptake the major substrate of metabolism such as glucose to increase their ATP output by forcing the aerobic glycolysis. Upon uptake, glucose is converted into lactate and ATP is produced during the process, which can be used as an energy source for sustained growth. As aerobic glycolysis is maintained, oncogenes are activated and tumor suppressor genes are inhibited. Previous studies showed that altered expression of oncogenes, such as hypoxia inducible factor-1α (HIF-1α), c-Myc, and Akt, and tumor suppressor genes, such as PTEN and p53, allows stable maintenance of aerobic glycolysis [6,7].
Cluster of differentiation 44 (CD44) is a major adhesion protein that is involved in various biological processes, including cell proliferation, cell migration, cell invasion, and cancer cell metabolism [8,9]. Many studies have focused on CD44 as a cancer stem cell marker, whereas its role in the mechanisms related to aerobic glycolysis in cancer cells remains unknown. Previously, it has been shown that the intracellular domain of CD44 interacts with pyruvate kinase M2 (PKM2), which is involved in the Warburg effect [10–12]. Moreover, the expression of CD44 enhanced glycolytic phenotypes via the regulation of PKM2 in p53-deficient or hypoxic cancer cells .
The Src family kinases, which have been reported to be oncogenic proteins, are related to cell proliferation, cell migration, and cell invasion [13–15]. The Src family kinases have direct or indirect interactions with the membrane receptor proteins and can modulate intracellular signaling. CD44-mediated signaling was shown to involve Src family kinases. In particular, c-Src binds to the cytoplasmic tail of CD44 with high affinity [16–18]. We recently showed that silencing of CD44 inhibits cell proliferation, migration, and invasion in breast cancer cells via down-regulation of c-Src transcription .
The AMP-activated protein kinase (AMPK) is a highly conserved serine/threonine protein kinase complex with an αβγ heterotrimeric structure. AMPK has important roles as a metabolic sensor in the maintenance of cellular energy homeostasis. When an increase in AMP or ADP:ATP ratio is sensed, AMPK immediately responds to the declining fuel supply and activates cellular energy processes to induce ATP production [20,21]. Previously, at least two upstream kinases, liver kinase B1 (LKB1) and calcium/calmodulin-dependent kinase kinase, have been shown to phosphorylate Thr172 of the catalytic α-subunit of AMPK in order to increase ATP production [22–26]. Ultimately, AMPK promotes the maintenance of ATP level under conditions of metabolic stress by inhibiting anabolic processes, such as lipid metabolism via the regulation of upstream kinases  and mTORC1-dependent downstream protein synthesis [28,29].
Lactate dehydrogenase (LDH) catalyzes the bidirectional conversion from pyruvate to lactate and vice versa. In mammals, there are five tetrameric LDH isoenzymes, which are composed of various combinations of LDHA (muscle type, M) and LDHB (heart type, H) . LDH4 (A3B1) and LDH5 (A4), which contain higher ratios of LDHA than LDHB, promote the shift from pyruvate to lactate production. In contrast, LDH1 (B4) and LDH2 (A1B3) favor the conversion from lactate to pyruvate [31–33]. Therefore, LDH1 and LDH2 are predominantly involved in the conversion of pyruvate from lactate, which is used in OXPHOS, whereas LDH4 and LDH5 are preferentially associated with the metabolic transition toward glycolysis to produce ATP in cancer cells . The expression of LDHA is associated with tumor cell progression. In fact, c-Myc and HIF-1α bind to specific sites on the LDHA promoter in various cancer cells and regulate its transcription toward a glycolytic phenotype [34–36]. In this report, our data indicate that CD44 drives the metabolic switch via modulation of LDH isoenzyme expression by the regulation of the c-Src–Akt–LKB1–AMPKα signaling pathway.
Materials and methods
Cell culture and transfection
MDA-MB-231, Hs578T, MCF7, and 293T cells were maintained in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 100 U/ml penicillin, and 100 mg/ml streptomycin. Cells were incubated at 37°C in a 5% CO2 in a humidified atmosphere. Glucose withdrawal conditions were accomplished by washing with glucose-free medium followed by DMEM with free glucose (contained 10% fetal bovine serum, 100 U/ml penicillin, and 100 mg/ml streptomycin). Calcium phosphate transfection was performed in 293T cells as described previously .
Cell viability analysis
Cells were seeded at 8 × 103 cells per well in 96-well plates and exposed to indicated concentrations of rotenone, CoCl2. Cell viability was measured by adding 20 μl of 10 mg/ml MTT (3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide) (Sigma, St. Louis, MO, USA) to 100 μl of culture medium and incubating for 3 h at 37°C. After 3 h, the medium was removed and formazan was dissolved in dimethyl sulfoxide (Sigma). The optical density was assessed at 590 nm using a Multiscan EX (Thermo, Vantaa, Finland).
Glucose uptake cell-based assay
The glucose uptake levels were assessed using a Glucose Uptake Cell-Based Assay kit (Cayman CHEMICAL, Ann Arbor, MI, USA). Cells were seeded at 1 × 104 cells per well in a 96-well Nunc™ F96 MicroWell™ Black Polystyrene Plate (Thermo, Vantaa, Finland). After 24 h, the levels of glucose uptake were measured according to the manufacturer's instruction. Relative fluorescence units were examined at 485–535 nm using a VARIOSKAN FLASH (Thermo, Vantaa, Finland).
The lactate production levels were determined using a Lactate Assay kit (Biovision, Milpitas, CA, USA). Cells were seeded at 1 × 106 cells in 100-mm cell culture dishes. After 24 h, the culture medium was changed with DMEM without FBS, followed by 8 h incubation. Culture media were collected from each sample and centrifuged at 2000 rpm for 5 min to obtain the supernatant without cell debris. Lactate assays were performed according to the manufacturer's protocol, and the colorimetric density was assessed at 570 nm using a Multiscan EX.
Quantification of ATP levels
The cellular ATP levels were measured using a CellTiter-Glo® Luminescent Cell Viability Assay kit (Promega, San Luis Obispo, CA, USA). Cells were seeded at a density of 2 × 104 cells per well in 96-well plates. After 24 h, the cellular ATP levels were examined according to the manufacturer's instruction. The luminescence was measured using a VARIOSKAN FLASH.
LDH isoenzyme zymography
LDH isoenzyme patterns were performed based on the native gel electrophoresis method. Cells were seeded at 1 × 106 cells in 100-mm cell culture dishes. After 24 h, cells were washed once with PBS and lysed in a lysis buffer without SDS for 1 h (10 mM Tris–HCl, pH 7.4, 10 mM NaCl, 3 mM MgCl2, 0.1% Triton X-100). After protein quantification, 30 μg of protein per sample was loaded onto a 7% non-denaturing polyacrylamide gel (running gel: 40% acrylamide 1.75 ml, Tris–HCl, pH 8.8, 2.5 ml, 10% ammonium persulfate 50 μl, dH2O 5.75 ml, TEMED 10 μl; stacking gel: 40% acrylamide 0.4 ml, Tris–HCl, pH 6.8, 1 ml, 10% ammonium persulfate 20 μl, dH2O 2.6 ml, TEMED 5 μl) and electrophoresed at 4°C for 90 min at 200 V. Following electrophoresis, the gels were placed in 10 ml of staining solution for 10 min at 37°C with 0.1 M lithium lactate, 1.5 mM NAD+, 0.1 M Tris–HCl, pH 8.6, 10 mM NaCl, 5 mM MgCl2, 0.3 mg phenazinmethosulphate (PMS), and 2.5 mg nitrobluetetrazolium (NBT). The assay was based on the conversion from lactate to pyruvate, with production of NADH and H+. NADH reduces PMS followed by a reduction in NBT to produce insoluble diformazan. Protein extracted from mouse heart served as the positive control.
Endogenous cellular oxygen consumption rate
The endogenous cellular oxygen consumption rate was measured using a Clark type O2 probe (Hansatech Instruments Ltd., Norfolk, UK) according to the manufacturer's instructions. Briefly, the culture media containing 1 × 106 cells were placed in a 37°C chamber. The measurement was carried out until the oxygen concentration reached extremely low level. The experiments were carried out in triplicate and repeated at least three times.
Western blot analysis
Cells were washed once with PBS and lysed in a lysis buffer [20 mM Tris–HCl, pH 7.4, 0.1 mM EDTA, 150 mM NaCl, 1% NP-40, 0.1% Triton X-100, 0.1% SDS, 20 mM NaF, 1 mM Na3VO4, 1× protease inhibitor (Roche, Basel, Switzerland)]. Protein samples were boiled for 10 min in SDS sample buffer, separated on SDS–PAGE gels, and transferred to nitrocellulose membrane (Whatman, Dassel, Germany). After blocking with 5% skim milk in TBS-T for 1 h, the membranes were incubated with the appropriate primary antibodies overnight. Membranes were washed once with TBS-T and incubated with horseradish peroxidase-conjugated secondary antibodies for 2 h. Protein bands were visualized with the WEST-ZOL-plus Western Blot Detection System (iNtRON Biotechnology, Seoul, Korea). The quantified values were located under each blot, which were normalized to actin or β-tubulin protein.
Quantitative real-time polymerase chain reaction
RNA was isolated using Trizol reagent (MRC, Cincinnati, OH, USA), and RT-PCR was carried out using an RNA PCR core kit (Roche). Quantitative real-time PCR (qPCR) was performed with an SYBR FAST qPCR kit (KAPA) in a Thermal Cycler Dice (Takara, Otsu, Shiga, Japan) according to the manufacturer's instructions. The C(t) value was normalized using GAPDH.
Cell fractionation assays
For cell fractionation assays, cells were seeded at a density of 1 × 106 cells per 100-mm dish. Cells were harvested in cytoplasmic extraction buffer [10 mM HEPES, pH 7.9, 10 mM KCl, 0.1 mM EDTA, 0.1 mM EGTA, 1 mM dithiothreitol (DTT), 0.5 mM PMSF] and incubated for 15 min on ice. The cells were then agitated for 10 min at 4°C, and NP-40 was added to produce a final concentration of 0.5%. The samples were centrifuged at 13 000 rpm for 5 min, and the supernatant was collected as the cytosolic fraction. The nuclear pellets were washed three times with cold PBS, resuspended in a nuclear extraction buffer (20 mM HEPES, pH 7.9, 400 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1 mM DTT, 1 mM PMSF), and the homogenates were incubated for 15 min on ice. The nuclear extracts were agitated for 10 min at 4°C and then centrifuged at 13 000 rpm and 4°C. The resulting supernatants were collected as the nuclear fraction.
Dual luciferase assays
Cells grown in 12-well plates were transfected with commercial plasmids (12 µg/well) and pCMV-RL (0.02 µg/well) as an internal control. Dual luciferase assays were performed according to the manufacturer's protocol (Promega). The experiments were carried out in triplicate and repeated at least three times.
Chromatin immunoprecipitation assays
For chromatin cross-linking, MDA-MB-231, Hs578T control shRNA, and CD44 shRNA cell lines (3 × 107 cells) were harvested, washed once with PBS, and incubated in 1.42% formaldehyde for 15 min at room temperature. The cells were then lysed in a lysis buffer (10 mM Tris, pH 8.0, 10 mM NaCl, 0.2% NP-40, 1× protease inhibitor), and the resulting lysates were centrifuged for 1 min at 12 000 rpm. Precipitated nuclei were treated with 200 U Micrococcal Nuclease (New England Biolabs, Hitchin, Hertfordshire, UK), resuspended in nuclei lysis buffer (50 mM Tris, pH 9.0, 10 mM EDTA, 1% SDS, 1× protease inhibitor), and diluted in IP dilution buffer (20 mM Tris, pH 8.0, 150 mM NaCl, 2 mM EDTA, 0.01% SDS, 1% Triton X-100, 1× protease inhibitor). The lysates were then disrupted with a sonicator (2 pulses, 30 s on/60 s off at 50% amplitude) to obtain DNA fragments of 200–500 bp in length (Dr. Hielscher, GmbH, Germany). Digested chromatin was immunoprecipitated with HIF-1α antibody overnight at 4°C with rotation. The immunocomplexes were collected with Protein G Sepharose 4 Fast Flow 50% slurry (w/v) for 3 h at 4°C with rotation and washed sequentially as follows: once with washing buffer 1 (20 mM Tris, pH 8.0, 150 mM NaCl, 2 mM EDTA, 0.1% SDS, 1% Triton X-100), once with washing buffer 2 (10 mM Tris, pH 8.0, 0.25 M LiCl, 1 mM EDTA, 1% NP-40, 1% deoxycholate), once with 0.1× TE buffer (1 mM Tris, pH 7.6, 0.1 mM EDTA), and twice with elution buffer (0.1 M NaHCO3, 1% SDS) for 5 min each. Eluted chromatin was reverse cross-linked overnight in 250 mM NaCl at 65°C, and DNA extraction was performed using phenol/chloroform and ethanol precipitation. The LDHA and LDHB promoter regions were PCR amplified.
All experiments were performed in triplicate. The results of cell viability analyses, glucose uptake cell-based assays, lactate assays, quantification of ATP levels, luciferase assays, and real-time PCR assays are expressed as mean ± standard deviation. Standard deviations for all measured biological parameters are displayed in the appropriate figures.
Ablation of CD44 suppresses the protein and mRNA expression of glycolytic genes
To investigate the effects of CD44 ablation on metabolism-related genes, we examined the phosphorylation status of LKB1 and AMPKα and the expression of HIF-1α, GLUT1, LDHA, and LDHB. The results shown in Figure 1A,C indicate that the expression of both the mRNA and protein of glycolytic genes, such as HIF-1α and LDHA, were inhibited in CD44 shRNA cells compared with control shRNA cells (Figure 1A,C). Interestingly, the level of LDHA, a well-known target gene of HIF-1α , was decreased in CD44 knock-down cells relative to control shRNA cells, but the levels of GLUT1, another target gene of HIF-1α , were not significantly different between the cell lines. To further confirm the effect of CD44 on metabolism-related genes, 293T and MCF7 cells were transfected with CD44 (Figure 1B,C). We found that CD44-overexpressing 293T and MCF7 cells exhibited increased expression of HIF-1α and LDHA and decreased LDHB expression, LKB1 and AMPK phosphorylation compared with 293T and MCF7 cells transfected with vector alone. LDH catalyzes the bidirectional reaction from pyruvate to lactate and from lactate to pyruvate depending on LDH isoenzyme ratio. LDH4 (A4) and LDH3 (A3B1) preferentially reduce pyruvate to lactate, and LDH1 (B4) and LDH2 (B3A1) oxidize lactate to pyruvate . Contrary to the expression of LDHA, the total protein and mRNA levels of LDHB were increased in CD44 shRNA cells (Figure 1A,C). These results suggest that CD44 ablation preferentially induces the reaction from lactate to pyruvate by increasing the LDHB expression ratio.
Effects of CD44 silencing on the protein and mRNA levels of glycolytic genes in human breast cancer cells.
Silencing of CD44 attenuates glucose uptake, ATP production, and lactate production and induces an LDH isoenzyme shift
Having shown that CD44 ablation regulates the expression of metabolism-related genes, we investigated the effects of CD44 knock-down on glucose uptake, cellular ATP production, lactate production, the LDH isoenzyme expression pattern, and endogenous cellular oxygen consumption rate. The glucose uptake capacity, cellular ATP level, and lactate production of CD44 knock-down cells were all down-regulated compared with the control shRNA cells (Figure 2A–C).
To further confirm the effect of CD44 ablation on the expression pattern of LDH isoenzymes, we performed native in-gel assays. The results indicate that control shRNA cells mainly contained the LDHA-rich isoenzymes LDH5 and LDH4. However, the knock-down of CD44 effectively shifted the LDH isoenzyme toward the LDHB-rich isoenzyme LDH2 (Figure 2D). We also observed enhancement of endogenous cellular oxygen consumption rate in CD44 knock-down cells compared with the control shRNA cells (Figure 2E). These results suggest that the ablation of CD44 undergoes metabolic shift toward mitochondrial respiration.
CD44 ablation suppresses glucose uptake, ATP production, and lactate production and induces an LDH isoenzyme shift in human breast cancer cells.
Silencing of CD44 induces a metabolic shift toward mitochondrial respiration
Previous studies have reported that most cancer cells rely on glycolysis for ATP generation regardless of the presence of oxygen . To test whether the ablation of CD44 affects the characteristics of cancer cell metabolism, we withdrew glucose from the medium or treated the cells with the mitochondrial respiratory chain complex 1 inhibitor, rotenone. Interestingly, 24 h of glucose withdrawal resulted in more cell death in the control shRNA cells than in the CD44 shRNA cells (Figure 3A). In contrast, treatment with rotenone for 48 h resulted in more cell death in CD44 shRNA cells than in control shRNA cells (Figure 3B). These results suggest that, in contrast with control shRNA cells, CD44 knock-down cells are more dependent on mitochondrial ATP production than on glycolysis. The shift toward reduction in glycolytic lactate production in CD44 knock-down cells might be associated with LDH isoenzyme changes (Figure 2D).
Silencing of CD44 induces a metabolic shift toward mitochondrial respiration.
CD44 ablation decreases HIF-1α binding to the LDHA promoter via suppression of HIF-1α expression
The data described above showed that CD44 regulates a metabolic transition via modulation of LDH isoenzyme expression (Figures 2 and 3), with the expression of glycolytic proteins, such as HIF-1α and LDHA down-regulated in CD44 shRNA cells compared with control shRNA cells (Figure 1A,C). Since previous publications have reported that HIF-1α is physically associated with the HIF-1α response element (HRE) site in the LDHA promoter region , we tested whether changes in LDHA level by CD44 ablation might be mediated by HIF-1α. The results shown in Figure 1C indicated that the HIF-1α mRNA level was decreased in CD44 knock-down cells, and that, as HIF-1α transcription was suppressed by CD44 knock-down, HIF-1α-dependent transcriptional activity was also down-regulated in CD44 shRNA cells compared with control shRNA cells (Supplementary Figure S1A). In general, transcription factors translocate to the nucleus and regulate target gene transcription by binding directly to the target promoter. To verify this in our tested cells, we performed cell fractionation assays. As expected, translocation of HIF-1α to the nucleus was decreased in CD44 shRNA cells relative to control shRNA cells as a result of reduced HIF-1α expression in CD44 knock-down cells (Supplementary Figure S1B).
To confirm that LDHA expression was regulated by HIF-1α, we treated the cells with cobalt chloride (CoCl2), a chemical inducer of HIF-1α [39,40]. CoCl2 induced the translocation of HIF-1α into the nucleus and promoted the expression of LDHA mRNA and protein (Figure 4A,B). When the cells were transfected with HIF-1α, the mRNA and protein levels of LDHA were increased in cells that overexpressed HIF-1α (Supplementary Figure S1C,D,G). We also observed reductions in LDHA promoter luciferase activity in CD44 knock-down cells relative to control shRNA cells (Supplementary Figure S1F).
In addition, the association of HIF-1α with the LDHA promoter region was also decreased in CD44 shRNA cells compared with control shRNA cells (Figure 4C), as determined by chromatin immunoprecipitation (ChIP) assays. These results indicate that silencing of CD44 suppresses HIF-1α binding to LDHA promoter via reduction in HIF-1α expression.
Silencing of CD44 decreases the expression of LDHA, but increases the LDHB expression via suppression of HIF-1α expression.
CD44 suppresses the expression of LDHB via HIF-1α-dependent transcriptional repression
We demonstrated that silencing of CD44 down-regulates LDHA expression, but up-regulates LDHB expression (Figure 1A,C). Previous studies have also reported that HIF-1α negatively regulates the expression of the human cad gene as a transcriptional repressor . Therefore, we suggest that HIF-1α is causally associated with enhancement of LDHB expression in CD44 shRNA cells, in contrast with its effects on LDHA expression.
We used CoCl2 treatment to assess if HIF-1α negatively regulates LDHB expression. Interestingly, as HIF-1α nuclear translocation or expression was increased by CoCl2, the mRNA and protein expression of LDHB was suppressed (Figure 4D,E and Supplementary Figure S2A,B). To exclude the possibility that the decrease in LDHB mRNA expression by HIF-1α is due to CoCl2-induced cell death, cell viability was measured 24 h after CoCl2 treatment. No significant decrease in cell viability was observed in any of the cell lines (Supplementary Figure S2C). In addition, as a result of HIF-1α overexpression, there was a reduction in both mRNA and protein levels of LDHB (Figure 4F,G and Supplementary Figure S2D,F). To further confirm the regulation of LDHA expression by HIF-1α, we inhibited HIF-1α expression by siRNA transfection. The result showed that silencing of HIF-1α down-regulated LDHA expression, while up-regulating LDHB expression (Supplementary Figure S1E). Based on these data, we investigated whether down-regulation of LDHB was regulated by HIF-1α at the transcriptional level. The LDHB promoter sequence, available from the National Center for Biotechnology Information (NCBI, NG_017038.1), was retrieved and analyzed for putative promoter sequence containing HRE consensus region using Promoter 2.0 Prediction Server (www.cbs.dtu.dk/services/Promoter/) and PROMO (http://alggen.lsi.upc.es/cgi-bin/promo_v3/promo/promoinit.cgi?dirDB=TF_8.3). The two HRE sequences (5′-RCGTG-3′) were identified at the -2927/-2923 and -8467/-8463 of the LDHB promoter (Supplementary Figure S2E). To demonstrate that this HRE site in the LDHB promoter was functional, we cloned LDHB promoters [-8661/-7682, LDHB promoter (1); -3534/-2521, LDHB promoter (2)] into a pGL3-basic luciferase reporter plasmid. Cells were then transfected with LDHB promoter luciferase plasmids and subjected to HIF-1α overexpression. The results shown in Figure 4H and Supplementary Figure S2H indicate that the repression of LDHB expression is regulated by HIF-1α at the transcriptional level on LDHB promoter region (2) despite the fact that one of the HRE sites, LDHB promoter region (1) was not a functional site in our cell lines (Supplementary Figure S2G). In contrast, HIF-1α overexpression did not inhibit LDHB promoter activity in our cell lines transfected with plasmids containing a mutated HRE site (Figure 4I).
To demonstrate that HIF-1α physically binds to the LDHB promoter HRE site and represses LDHB promoter activity, we performed ChIP assay and analyzed the binding of HIF-1α to the LDHB promoter. The binding of HIF-1α to the LDHB promoter was significantly increased by HIF-1α transfection (Figure 4J and Supplementary Figure S2I). These results indicate that, as a transcriptional repressor, HIF-1α directly binds to the LDHB promoter HRE site and negatively regulates the expression of LDHB.
CD44 ablation attenuates HIF-1α expression via the LKB1/AMPKα/mTOR signaling pathway
So far, we have shown that HIF-1α inversely regulates LDHA and LDHB expression. A recent study reported that mTORC1 regulates HIF-1α mRNA transcription . To identify the missing link between CD44 and HIF-1α, we treated cells with rapamycin, a specific inhibitor of mTOR signaling, and analyzed the mRNA and protein expressions of HIF-1α, LDHA, and LDHB. The results showed that rapamycin reduced the Ser2448 phosphorylation of mTORC1, a marker of mTORC1 sensitivity to rapamycin , and the levels of HIF-1α and LDHA, while inducing LDHB level. However, we did not see a dramatic induction of LDHB in CD44 shRNA cells due to their higher basal expression level of LDHB compared with control shRNA cells (Figure 5A,B).
It has been observed that AMPK negatively regulates the Warburg effect via the down-regulation of mTORC1 signaling. Also, it has been reported that the AICAR-induced AMPK activation resulted in a decrease in Ser2448 phosphorylation of mTORC1 . To verify this, we treated our samples with metformin, an AMPK activator, or transfected with a full-length AMPKα1 and α2 constructs.
Treatment with metformin for 24 h resulted in suppression of mTOR phosphorylation, HIF-1α, and LDHA levels and enhancement of the LDHB level (Figure 5E). In addition, the cells overexpressing full-length AMPKα1 and α2 exhibited the same results (Figure 5C,D). To further confirm that AMPK mediates the effect of CD44 on the expression of LDHA and LDHB, we transfected our samples with AMPKα1/2 siRNA. The results shown in Figure 5E indicate that the expression of LDHA and LDHB are regulated by AMPK/mTOR/HIF-1α signaling axis (Figure 5F).
CD44 ablation attenuates HIF-1α expression via the LKB1/AMPKα/mTOR signaling pathway.
It is known that LKB1 phosphorylates the Thr172 of the catalytic α-subunit of AMPK to increase ATP production. To investigate the role of LKB1 in the CD44-mediated regulation of HIF-1α, we examined the effects of CD44 ablation on LKB1. We found that the phosphorylation of Ser428 by LKB1 was increased in CD44 shRNA cells relative to control shRNA cells (Figure 1A), suggesting that CD44 ablation leads to activation of LKB1.
Taken together, these observations suggest that CD44-dependent modulation of LDH isoenzyme expression is regulated by HIF-1α via serial phosphorylation of LKB1, AMPKα, and mTORC1 in our cell lines.
CD44 induces the differential expression of LDH isoenzymes via modulation of the c-Src/Akt/LKB1/AMPKα/mTOR/HIF-1α pathway
We recently showed that CD44 ablation inhibits cell proliferation, migration, and invasion via down-regulation of the c-Src–Akt pathway . Recent data have also indicated that Akt phosphorylates Ser487 in the ST loop of AMPKα and down-regulates its activation by inhibiting the phosphorylation of Thr172 . To test whether inversely expressed LDH levels are regulated by the c-Src–Akt–LKB1–AMPKα–mTOR–HIF-1α pathway in our cell lines, we treated the cells with PP2, a specific inhibitor of c-Src, or Akt#8, an Akt inhibitor, and transfected the cells with HA-tagged c-Src or myristoylated Akt1 and examined the levels of downstream signaling molecules.
Treatment with inhibitors resulted in increased phosphorylation of LKB1 and AMPKα, decreased phosphorylation of mTOR, mRNA, and protein expression of HIF-1α and LDHA, and increased expression of LDHB in our cell lines (Figure 6C and Supplementary Figure S3). However, overexpression of HA-tagged c-Src or myristoylated Akt1 resulted in inhibition of LKB1 and AMPKα phosphorylation, increased phosphorylation of mTOR, up-regulation of HIF-1α at both mRNA and protein levels, increased expression of LDHA, and decreased expression of LDHB in our cell lines (Figure 6A,B and Supplementary Figure S3). These results suggest that silencing of CD44 down-regulates LDHA expression and up-regulates LDHB level via modulation of the c-Src–Akt–LKB1–AMPKα–mTOR–HIF-1α pathway.
CD44 regulates LDH expression via modulation of the c-Src/Akt/LKB1/AMPKα/mTOR/HIF-1α pathway.
The CD44-mediated metabolic switch is mediated by the c-Src/Akt/LKB1/AMPKα/mTOR/HIF-1α pathway
We previously demonstrated that CD44 regulates LDH expression via modulation of the c-Src–Akt–LKB1–AMPKα–mTOR–HIF-1α pathway. To further confirm that the CD44-mediated metabolic transition is mediated by the c-Src–Akt–LKB1–AMPKα–mTOR–HIF-1α pathway, we treated cells with PP2, Akt#8, CoCl2, and HA-tagged c-Src and analyzed the metabolic phenotypes. As a result of treatment with PP2 and Akt#8, glucose uptake capacity and lactate production were suppressed. However, treatment with CoCl2 promoted the glycolytic phenotypes. Also, overexpression of c-Src induced the glucose uptake capacity (Supplementary Figure S4). Interestingly, when the cells were subjected to co-treatment with PP2 and CoCl2, the increases in glucose uptake capacity and lactate production by CoCl2 were decreased by PP2 (Figure 7A,B), suggesting that the glycolytic phenotypes promoted by HIF-1α can be overridden by inhibition of c-Src downstream signaling.
Based on these results, we assumed that CD44 was involved in the metabolic switch by the regulation of LDH expression. The activation of c-Src by CD44 could lead to decreased LKB1 and AMPKα activation via Akt signaling, which might lead to increased HIF-1α expression via mTORC1 signaling, resulting in up-regulation of LDHA and down-regulation of LDHB level. As a result of enhanced glycolysis, glucose uptake ability, cellular ATP production, and lactate production are promoted in CD44-expressing breast cancer cells (schematically illustrated in Figure 7C).
Schematic representation of the CD44-mediated metabolic switch.
Various cancer cells depend on glycolytic phenotypes characterized by high rates of glucose uptake, lactate production, and cellular activity to produce ATP regardless of oxygen concentration, a phenomenon referred to as the ‘Warburg effect’ . Recently, targeting metabolism-related signaling pathway in cancer cells exhibiting Warburg effect emerges as a novel approach to find new therapeutics.
Cancer cells have many metabolism-related signaling pathways, including the PI3K, HIF-1α, Myc, AMPK, p53, and OCT1 pathways [47–51]. Among these pathways, the mechanism for CD44-dependent regulation of aerobic glycolysis in cancer cells remains largely unknown. CD44 is commonly reported to be a central upstream signaling molecule that regulates cell proliferation, migration, and invasion [52,53]. To date, CD44 has been reported to be a metastatic tumor marker in various cancer cells . Interestingly, it has previously been reported that CD44 directly associates with PKM2, which regulates glycolysis and the pentose phosphate pathway , and that CD44 takes center stage in the regulation of cancer cell metabolism. Because many previous studies identified that cancer stem-like cells have more glycolytic phenotype, and CD44 was known as breast cancer stem cell marker, we established CD44 knock-down cell line using endogenous CD44-positive breast cancer cell lines, MDA-MB-231 and Hs578T, and performed various experiments to identify the effect of CD44 ablation on cancer cell metabolism.
In our study, the silencing of CD44 resulted in suppression of glycolytic gene expression, such as HIF-1α and LDHA, leading to inhibition of glucose uptake, ATP production, lactate production, a change in LDH isoenzyme expression pattern, and increase in endogenous cellular oxygen consumption rate. Under hypoxic conditions, a normal cell shifts its energy metabolism from OXPHOS to glycolysis to generate ATP. However, cancer cells produce cellular ATP mainly by glycolysis, regardless of oxygen concentration , and a reduction in glycolytic phenotypes by CD44 silencing suggests that cancer cell metabolism shifts toward OXPHOS after CD44 ablation. Therefore, in the present study, we identified CD44 as a key molecule determining the metabolic shift from glycolysis to OXPHOS.
HIF-1α is an important transcription factor that is expressed when cells are exposed to hypoxic conditions [36,55]. Previous studies have reported that HIF-1α regulates the expression of ALDA, enolase 1, and LDHA genes as a transcription factor, with the promoter sequences of these genes activated in hypoxic cells [56–58]. In contrast with prior reports, a novel role of HIF-1α as a transcriptional repressor has also been reported . In our study, we observed a decrease in the binding of HIF-1α to the LDHA promoter as a result of suppression of HIF-1α expression in CD44 knock-down cells. In contrast, we observed increased expression of LDHB as a result of CD44 silencing via transcriptional repression by HIF-1α. Therefore, we suggest that the metabolic shift toward OXPHOS in CD44-silenced cells is due to inversely expressed LDH isoform levels and HIF-1α plays a pivotal role in determining metabolic shift between glycolysis and OXPHOS.
AMPK has an important role in maintaining cellular energy homeostasis . AMPK inhibits cellular processes, such as anabolism to maintain the cellular ATP level in conditions of metabolic stress. We showed that silencing of CD44 induces AMPK activity, which is reported to be a downstream molecule of LKB1. AMPK is a kinase that negatively regulates the Warburg effect and suppresses mTORC1 signaling and mTORC1-dependent protein synthesis [28,29]. In addition, mTORC1 is reported to be a direct regulator of HIF-1α transcription . In the present study, we also identified that CD44 ablation suppresses both mRNA and protein expression of HIF-1α, because of the decrease in mTOR1 signaling.
We recently showed that CD44 regulates cell proliferation, migration, and invasion in breast cancer cells via modulation of c-Src transcription . c-Src has been known to bind the cytoplasmic tail of CD44 [16–18], and Akt has been identified as the downstream molecule of c-Src . Also, it has been known that LKB1 is an upstream molecule of AMPK. Here, we demonstrated that suppression of c-Src and Akt activity by CD44 ablation results in LKB1 activation by Ser428 phosphorylation of LKB1, culminating in the regulation of AMPK and mTOR activity, and expression of HIF-1α, LDHA, and LDHB. Currently, we do not have an explanation for the inhibition of Ser428 phosphorylation in LKB1 by enhanced Akt activity, although we have evidence that induction of c-Src or Akt expression by genetic constructs or inhibition of c-Src or Akt by chemical inhibitors leads to a decrease or increase in Ser428 phosphorylation in LKB1. Further studies are needed to elucidate direct linkage between LKB1 Ser428 phosphorylation and Akt kinase activity.
In conclusion, the results of the present study indicate that the breast cancer stem cell marker CD44 plays a pivotal role in the regulation of cancer cell metabolism by modulation of LDH levels. We suggest that ablation of CD44 results in inverse regulation of LDH isoenzyme expression through decreased c-Src and Akt activity and subsequent LKB1 and AMPK activation, which in turn suppresses mTOR phosphorylation and HIF-1α expression. CD44 was shown to play a role in the regulatory pathways that affect LDH isoenzyme expression, resulting in the regulation of glucose uptake, cellular ATP production, and lactate production in human breast cancer cells (schematically illustrated in Figure 7).
AMPK, AMP-activated protein kinase; ATP, adenosine triphosphate; CD44, cluster of differentiation 44; ChIP, chromatin immunoprecipitation; CoCl2, cobalt chloride; DMEM, Dulbecco's modified Eagle's medium; DTT, dithiothreitol; FBS, fetal bovine serum; HIF-1α, hypoxia inducible factor-1α; HRE, HIF-1α response element; LDH, lactate dehydrogenase; LKB1, liver kinase B1; MTT, 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide; NBT, nitrobluetetrazolium; OXPHOS, oxidative phosphorylation; PKM2, pyruvate kinase M2; PMS, phenazinmethosulphate.
K.N. contributed to the design and performance of the experiments, analyzed the data and wrote the manuscript. S.O. contributed to the analysis and interpretation of data. I.S. conceived and led the study. All authors have discussed with the results and approved the submission of the manuscript.
This work was supported by an NRF grant [2013R1A1A2059143] from the Korea Research Foundation; the Converging Research Center Program funded by the Ministry of Science, ICT & Future Planning [Project No. 2015054348]; and the Civil research projects for solving social problems through the National Research Foundation of Korea funded by the Ministry of Science, ICT & Future Planning (grant no. [2015M3C8A6A06012226]).
The Authors declare that there are no competing interests associated with the manuscript.