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Available online at www.sciencedirect.com
www.elsevier.com/locate/yexcr
Research Article
PHD1 interacts with ATF4 and negatively regulates its transcriptional activity without prolyl hydroxylation Yusuke Hiwatashi, Kohei Kanno, Chikahisa Takasaki, Kenji Goryo, Takuya Sato, Satoru Torii, Kazuhiro Sogawa, Ken-ichi Yasumoto⁎ Department of Biomolecular Sciences, Graduate School of Life Sciences, Tohoku University, Aoba-ku Sendai 980-8578, Japan
A R T I C L E I N F O R M A T I O N
A B S T R A C T
Article Chronology:
Cellular response to hypoxia plays an important role in both circulatory and pulmonary diseases and
Received 3 October 2010
cancer. Hypoxia-inducible factors (HIFs) are major transcription factors regulating the response to
Revised version received
hypoxia. The α-subunits of HIFs are hydroxylated by members of the prolyl-4-hydroxylase domain
4 September 2011
(PHD) family, PHD1, PHD2, and PHD3, in an oxygen-dependent manner. Here, we report on the iden-
Accepted 9 September 2011
tification of ATF4 as a protein interacting with PHD1 as well as PHD3, but not with PHD2. The central
Available online 17 September 2011
region of ATF4 including the Zipper II domain, ODD domain and β-TrCP recognition motif were involved in the interaction with PHD1. Coexistence of PHD1 stabilized ATF4, as opposed to the desta-
Keywords:
bilization of ATF4 by PHD3. Moreover, coexpression of ATF4 destabilized PHD3, whereas PHD1
Hypoxia
stability was not affected by the presence of ATF4. Mutations to alanine of proline residues in ATF4
HIF prolyl hydroxylation
that satisfied hydroxylation consensus by PHDs did not affect binding activity of ATF4 to PHD1 and
ATF4
PHD3. Furthermore, in vitro prolyl hydroxylation assay clearly indicated that ATF4 did not serve as
PHD1
a substrate of both PHD1 and PHD3. Coexpression of PHD1 or PHD3 with ATF4 repressed the tran-
PHD3
scriptional activity of ATF4. These results suggest that PHD1 and PHD3 control the transactivation activity of ATF4. © 2011 Elsevier Inc. All rights reserved.
Introduction Adaptation to hypoxia is essential for the survival of normal tissues and tumors. The most important aspect of how cells respond to hypoxia is the regulation of the transcriptional activity of the hypoxiainducible factors (HIFs) [1,2]. An unstable alpha subunit (such as HIF-1α) and a stable beta subunit (such as HIF-1β, also termed ARNT) constitute an active transcription factor, HIF (such as HIF-1), that specifically binds to hypoxia response element (HRE) located in the promoter region of the hypoxia induced genes [3,4]. There
are three alpha subunits in the mammalian cells, HIF-1α, HIF-2α and HIF-3α. HIF-α protein abundance is regulated with three prolyl hydroxylase domain-containing proteins named PHD1, PHD2, and PHD3 in mammalian cells [5,6]. These PHDs belong to the superfamily of oxygenases that require O2, 2-oxoglutarate (2-OG), and Fe2+ for their enzymatic activity [7]. These PHDs catalyze the hydroxylation of specific proline residues within the oxygen-dependent degradation (ODD) domains of HIF-α subunits [3]. Subsequently, hydroxylated HIF-α subunits are bound to the von Hippel–Lindau tumor suppressor gene product (pVHL), which is a recognition component of an
⁎ Corresponding author. Fax: + 81 22 795 6594. E-mail address:
[email protected] (K. Yasumoto). Abbreviations: ATF4, activating transcription factor 4; β-TrCP, β-transducin repeat-containing protein; FRET, fluorescence resonance energy transfer; GLUT-1, glucose transporter type 1; HIF, hypoxia-inducible factor; NAD, N-terminal activation domain; 2-OG, 2-oxoglutarate; PHD, prolyl hydroxylase domain-containing protein; ODD, oxygen-dependent degradation. 0014-4827/$ – see front matter © 2011 Elsevier Inc. All rights reserved. doi:10.1016/j.yexcr.2011.09.005
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E3 ubiquitin ligase complex. Finally, hydroxylated α-subunits are subjected to rapid ubiquitination and proteasomal degradation [8,9]. Notably, PHDs function in an O2 concentration-dependent manner and are regarded as cellular O2 sensors. In response to oxygen deprivation leading to hypoxia, PHD activity decreases and HIF-α subunits are stabilized and translocated to the nucleus where it may transactivate target genes after the heterodimerization with HIF-1β [1]. Most of known target genes activated by HIFs are centrally involved in both systemic responses to hypoxia, such as angiogenesis and erythropoiesis (e.g. VEGF and Epo, respectively), and in cellular responses, such as alteration in glucose/energy metabolism (e.g. GLUT1 and glycolytic enzymes) [3]. Moreover, hypoxic response is one of the important stress responses in the cancer progression. Tumor hypoxia is the result of poorly formed tumor vasculature [10], and is a significant obstacle to therapy. Impairment of blood supply in the tumor microenvironment limits the availability not only of oxygen but also of nutrients such as glucose and amino acids. There have been several reports that expression levels of transcription factor ATF4 were significantly higher in primary human tumors compared to corresponding normal tissues [11,12]. ATF4 expression is upregulated via the integrated stress response (ISR) that is activated by several discrete stresses such as ER stress, amino acid starvation, heme deficiency, and viral infection. Activation of four specific kinases, PERK, GCN2, HRI, and PKR, by those respective stresses results in the phosphorylation of the translation initiation factor eIF2α [13]. Global translation is arrested by the phosphorylation whereas ATF4 mRNA translation is elevated, resulting in the activation of the expression of ATF4 target genes [14]. It has been revealed that target genes of ATF4 include genes for redox balance (such as heme oxygenase-1) [15], apoptosis (such as CHOP, TRB3) [14,16], and amino acid metabolism (such as ASNS, CAT-1) [17,18]. A recent study reported that the activation of ATF4 facilitates tumor cell survival under nutrient deprivation [19] suggesting that the regulation mechanism of ATF4 in the tumor microenvironmental region could be the target for cancer therapy. In this study, we report that ATF4 directly interacts with both PHD1 and PHD3, and these proteins repress the transcriptional activity of ATF4. Meanwhile, it is revealed that ATF4 does not serve as a substrate of both PHD1 and PHD3 using in vitro prolyl hydroxylation assay. These results showed a novel function of PHD1 and PHD3 that is not correlated with the prolyl hydroxylation activity.
subcloning the deletion mutants or substitution mutants of ATF4 into pBOS-NLS-VP16 vector. pCitrine-C1 was constructed by introducing one amino acid substitution, Q69M, into pEYFP-C1 (Clontech). pCitrine-PHD1 and pCitrine-PHD2 were constructed as described previously [22]. An open reading frame of human PHD3 was amplified by PCR and inserted into pCitrine-C1 termed as pCitrine-PHD3. pCerulean-C1 was constructed by introducing three amino acid substitutions, S72A, Y145A, and H148D, into pECFP-C1 (Clontech). pCerulean-ATF4 was constructed by the insertion of ATF4 open reading frame into pCerulean-C1. Synthesized oligonucleotides encoding three tandemlyrepeated Myc tag were ligated into pBOS vector (designated as pBOS-3Myc). Full-length or partial regions of ATF4 were amplified by PCR and ligated into pBOS-3Myc vector. To add Flag-tag in front of the termination codon, the open reading frame region of PHD1, PHD2, PHD3 and ATF4 was amplified by PCR using specific primers containing Flag-tag encoding sequence, and subsequently the amplified fragments were ligated into pBOS vector. Obtained plasmids were designated as pBOS-PHD1-Flag, pBOS-PHD2-Flag, pBOS-PHD3-Flag, and pBOS-ATF4-Flag, respectively. Mutant plasmids of pBOS-ATF4-Flag were constructed using QuikChange II site-directed mutagenesis kit (Stratagene) with appropriate primers. CHOP promoter-Luc plasmid was constructed by inserting CHOP (GADD153) promoter region corresponding from −772 to +51 into pGL3-basic vector. pCP2-ATF4, expressing hexahistidine (H6)tagged ATF4 in E. coli, was constructed by inserting H6-ATF4 encoding fragment into pCP2 vector [23].
Cell culture HeLa and HEK293T cells were maintained at 37 °C in minimal essential medium (MEM) and low-glucose Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 50 U/ml penicillin, and 5 μg/ml streptomycin, respectively. HepG2 cells were cultured at 37 °C in MEM containing 10% FBS, 50 U/ml penicillin, 5 μg/ml streptomycin, 1% MEM nonessential amino acid (Wako chemicals), 1 mM sodium pyruvate. For hypoxia conditions, O2 levels were decreased to 1% O2 with N2 in an oxygen-controlled incubator. In some experiments, cells were treated with 10 μM MG132 (Peptide Institute), 20 μM E-64 (Roche), 1 μg/ml Pepstatin A, 10 μg/ml cycloheximide for the indicated time periods.
Material and methods Mammalian two-hybrid assay Plasmid construction pBOS, pBOS-GAL4DBD, and pBOS-LacZ were described previously [20]. pG5ELuc was produced by inserting five copies of the GAL4 binding site with adenovirus E1b promoter region into pGL3basic vector (Promega). Each open reading frame of human PHD1, PHD2 and PHD3 was cloned by PCR amplification and inserted into the pBOS-GAL4DBD vector, and the resulting plasmid was designated as pBOS-GAL4DBD-PHD1, pBOS-GAL4DBD-PHD2 and pBOS-GAL4DBD-PHD3, respectively. pBOS-NLS-VP16 was produced as pNV vector as described previously [21]. An open reading frame of ATF4 was amplified by PCR and inserted into pBOS-NLS-VP16 vector, designated as pBOS-NLS-VP16-ATF4. Various pBOS-NLS-VP16-ATF4 derivatives were constructed by
HepG2 cells were transfected with 1 μg of luciferase reporter plasmid pG5E-LUC, 1 μg of pBOS-LacZ for the internal standardization, 1.5 μg of pBOS-GAL4DBD or its derivatives, and 1.5 μg of pBOSNLS-VP16 or its derivatives by the calcium phosphate precipitation method as described previously [24]. 16 h after transfection, cells were harvested and lysed in 100 mM potassium phosphate buffer (pH 7.7) containing 1 mM dithiothreitol (DTT) by three cycles of freezing and thawing treatment. Subsequently, luciferase activity was measured with luciferase assay system (Promega) using a Lumat LB9507 luminometer (Berthold) and normalized with each β-galactosidase activity, which was used as an internal control for transfection efficiency. All values are the means calculated from the results of at least three independent experiments.
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Immunoblot analysis HEK293T cells were transiently transfected with 5 μg of expression plasmid by the calcium phosphate precipitation method. The indicated time periods after transfection, cells were harvested and lysed in RIPA buffer (10 mM Tris–HCl (pH 7.4), 150 mM NaCl, 1% Triton X-100, 1% deoxycholic acid, 0.1% sodium dodecyl sulfate (SDS), 50 mM NaF, 1 mM Na3VO4, 1 mM DTT, 1 mM EDTA, 1% protease inhibitor cocktail (Nacalai Tesque), and 1 μM MG132). Proteins were separated by 8–12% SDS-PAGE. After electrophoresis, the proteins were blotted onto Immobilon-P (Millipore) and probed with rabbit anti-Flag polyclonal antibody (Sigma), rabbit anti-Myc polyclonal antibody (Medical & Biological Laboratories), rabbit anti-HA polyclonal antibody (Medical & Biological Laboratories), or mouse anti-Myc monoclonal antibody (clone 9E10; Santa Cruz Biotechnology). The detection was performed using ECL plus detection kit (GE Healthcare).
Immunoprecipitation assay HEK293T cells were transiently transfected 5 μg of expression plasmid by the calcium phosphate precipitation method. 24 h after transfection, cells were harvested and lysed in immunoprecipitation buffer (50 mM Tris–HCl (pH 8.0), 150 mM NaCl, 0.5% Triton X-100, 10% glycerol, 0.1% NP-40, 1 mM DTT, 1 mM EDTA, 1% protease inhibitor cocktail, and 10 μM MG132). For immunoprecipitations, cell lysates were pre-cleared with normal mouse IgG and protein G sepharose (GE Healthcare) for 1 h at 4 °C, and supernatants were incubated with mouse anti-Myc monoclonal antibody and protein G sepharose, or anti-FLAG M2 agarose affinity gel (Sigma) for more than 2 h at 4 °C. Immune complexes were washed with PBS and then eluted with 2× SDS sample buffer (0.125 M Tris–HCl (pH 6.8), 4% SDS, 20% glycerol, 10% 2mercaptoethanol (2-ME)). Proteins were separated by SDSPAGE and detected by immunoblot analysis using appropriate antibody.
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performed with an Olympus IX-71 fluorescence microscope equipped with a filter set (Olympus U-MCFPHQ and U-MYFPHQ for Cerulean and Citrine, respectively) and the Olympus DP70 CCD camera.
In vitro 1-[ 14C]-CO2 capture assay PHD1 and PHD3 were expressed and purified as described previously [27]. HIF-1α NAD was prepared as follows. The recombinant protein of glutathione S-transferase fused to HIF-1α NAD (482– 575, C520A) with Flag tag was expressed in E. coli BL21 (DE3) strain. Cell lysates were applied to Phenyl-Sepharose, Glutathione-Sepharose (GE Healthcare), and Protein Pak G-DEAE column (Waters) to obtain the purified GST-HIF-1α NAD protein. Subsequently, purified protein fraction was applied to thrombin digestion, and HIF-1α NAD fragment was isolated with μBondasphere C18 reverse-phase HPLC (Waters). H6-ATF4 was purified as follows. E. coli strain BL21 (DE3) with pCP2-ATF4 was cultured, and cell pellets were resuspended in buffer A (50 mM Tris–HCl (pH7.8), 0.5 M NaCl, 5 mM 2-ME, 1 mM PMSF (phenylmethylsulfonyl fluoride), 1 mM benzamidine, 0.2% Triton X-100) containing 20 mM imidazole. The suspension was sonicated on ice, and centrifuged to remove cell debris. The supernatant was applied to the Ni-NTA agarose column. Subsequently, the bound proteins were eluted with buffer A containing 200 mM imidazole, and dialyzed against buffer A containing 20 mM imidazole. The fraction was re-applied to the Ni-NTA agarose. Finally, the purity of purified H6-ATF4 was confirmed by SDS-PAGE followed by the Coomassie staining. Erabutoxin b was prepared as described previously [28]. The prolyl hydroxylation activity was analyzed by a modified method based on the hydroxylation-coupled decarboxylation of 1-[14C]-2-oxoglutarate [29,30]. The reaction was performed in a final volume of 50 μl as described previously [27] although the amount of the substrate was reduced from 20 nmol to 5 nmol.
FRET analysis in living cells Luciferase assay HeLa cells or HepG2 cells were transfected with 0.5–1 μg of CHOP promoter-Luc, 0.5–1 μg of pBOS-LacZ for internal standardization, and 3–4 μg of various expression plasmids by the calcium phosphate precipitation method. The total amount of DNA was kept constant at 5 μg. 4 h after transfection, cells were treated with PBS containing 20% glycerol for 2 min, and subsequently the glycerol solution was replaced with the fresh medium. Tunicamycin (Sigma) was added at this point to the final concentration of 2 μg/ml. 16 h after glycerol treatment, cells were harvested and the procedures after the harvest were the same as described in the mammalian two-hybrid assay.
Imaging of the localization of the fluorescent proteins HeLa cells grown on the cover glass were transfected with 0.5 μg of plasmids expressing PHDs fused to Citrine [25] or ATF4 fused to Cerulean [26] using FuGENE6 transfection reagent (Roche). 24 h after transfection, cells were washed with PBS and fixed for 15 min in 4% paraformaldehyde in PBS. The fixed cells were incubated in 0.2 μg/ml of DAPI (4′,6′-diamidino-2-phenylindole) labeling solution for 3 min at room temperature. Imaging was
HEK293T cells were transfected with plasmids expressing fluorescent proteins using FuGENE6 transfection reagent. 24 h after transfection, cells were trypsinized and suspended in PBS, then applied to the detection of FRET (fluorescence resonance energy transfer). Each cell was measured for both fluorescence intensity and phase shift against modulated excitation light with Flicyme, fluorescence lifetime cytometer (Mitsui Engineering & Shipbuilding) according to the manufacturer's instructions. A total of 100,000 cells were measured at one experiment, and the fluorescent intensity of both Cerulean (FL1) and Citrine (FL2) were shown as cytogram (Supplementary Fig. 1). Positive cells were gated in the polygonal region on the cytogram, and applied to the analysis of fluorescence lifetime using frequency domain method [31,32]. Briefly, excitation light is modulated in a sinusoidal fashion, and Flicyme detects the phase shift (θ [rad]) between the excitation and emission. The values of modulation frequency (ωM [rad/s]) and phase shift give the fluorescence lifetime (τ) as described previously [32]. From the comparison of the fluorescent lifetime of donor protein (Cerulean or its derivatives), significant reduction of the lifetime indicates the interaction between two fluorescent proteins in living cells.
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Results Both PHD1 and PHD3 interact with ATF4 Using the yeast two-hybrid system, a cDNA expression library derived from mouse brain was screened using PHD3 as bait. Several clones remained positive through secondary and tertiary screenings, and ATF4 (activating transcription factor 4) was selected
A
and subjected to further validation. We examined the interaction between ATF4 and three PHD isoforms in mammalian cells using a mammalian two-hybrid assay. Since ATF4 contains a transcriptional activation domain that interacts with the transcriptional co-activator p300/CBP, we applied each PHD isoform as bait in this assay. The results implied that ATF4 interacts with PHD1 as well as PHD3 in HepG2 cells (Fig. 1A). No interaction was found between ATF4 and PHD2 (Fig. 1A). To confirm these interactions, we performed an immunoprecipitation assay with tagged ATF4
B Relative luciferase activity
450 400 350
PHD1-Flag
–
+
+
3Myc-ATF4
–
–
+
300
(kDa) 50
250 200 40
WB : anti-Flag Input
30 20
WB : anti-Myc 50
10
0 NLS-VP16-ATF4
–
–
–
–
+
+
+
+
GAL4DBD-PHD1
–
+
–
–
–
+
–
–
GAL4DBD-PHD2
–
–
+
–
–
–
+
–
GAL4DBD-PHD3
–
–
–
+
–
–
–
+
IP : anti-Flag
WB : anti-Myc 50 PHD2-Flag
–
+
+
3Myc-ATF4
–
–
+
(kDa) 50
WB : anti-Flag
C
Input Citrine
Citrine
Citrine
Cerulean
-PHD1
-PHD2
-PHD3
-ATF4
WB : anti-Myc 50
IP : anti-Flag
WB : anti-Myc 50
Citrine
Cerulean
Merge
DAPI
Citrine-PHD1 +
PHD3-Flag
–
+
+
3Myc-ATF4
–
–
+
(kDa)
Cerulean-ATF4
WB : anti-Flag Citrine-PHD2 +
25
Input
Cerulean-ATF4
WB : anti-Myc 50 Citrine-PHD3 + Cerulean-ATF4
IP : anti-Flag
WB : anti-Myc 50
Fig. 1 – (A) Mammalian two-hybrid assay using various bait plasmids and pBOS-NLS-VP16-ATF4 as prey. Cotransfection of each (1 μg) of the various bait plasmids and the ATF4 prey plasmid or pBOS empty plasmid (1 μg) into HepG2 cells was carried out together with the reporter gene, pG5ELuc (2 μg) and pBOS-LacZ (1 μg) as an internal control. The expressed luciferase activities were normalized by β-galactosidase activities used as a control of transfection efficiency. The means± standard deviation of three independent experiments are shown. (B) Interaction of PHDs with ATF4. Flag-tagged PHD proteins were expressed in the presence or absence of 3Myc-ATF4 in HEK293T cells. The PHDs were then immunoprecipitated (IP) with anti-Flag antibodies. Co-immunoprecipitated proteins were subjected to SDS-PAGE and detected with immunoblot analysis using anti-Myc antibodies. (C) Intracellular localization of PHDs and ATF4. PHD1, PHD2 and PHD3 were fused to Citrine, and ATF4 was fused to Cerulean. Each chimeric protein was transiently expressed in HeLa cells. Images of Citrine, Cerulean, their merged images, and DAPI fluorescence were shown.
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and PHDs (Fig. 1B). The results showed that both PHD1 and PHD3 efficiently co-precipitated with ATF4 indicating the interaction between ATF4 and PHD1 or PHD3, but not PHD2. To examine the intracellular localization of ATF4 and PHDs, these proteins were fused to fluorescent proteins, Cerulean or Citrine, and distribution of expressed chimeric proteins was observed using fluorescent microscopy. As described previously [33,34], ATF4 was dominantly located in the nucleus. Both PHD1 and PHD3 were co-localized with ATF4 in the nucleus, whereas PHD2 was exclusively distributed in the cytoplasm where no ATF4 signals were found (Fig. 1C). Taken together, these results indicated that ATF4 interacts with both PHD1 and PHD3 but not with PHD2. To assess the direct interaction between ATF4 and PHD1/PHD3 in the living cells, we measured decrease in fluorescence lifetime caused by FRET (fluorescent resonance energy transfer) using a fluorescence lifetime cytometer, Flicyme (Mitsui Engineering & Shipbuilding). Initially, two fluorescent proteins, Cerulean (donor) [26], and Citrine (acceptor) [25], were expressed independently or simultaneously in HEK293T cells. Each lifetime of Cerulean (detected in channel Tc1) and Citrine (detected in channel Tc2) was measured (Supplementary Fig. 1) and shown in Table 1. From the comparison of the donor lifetime derived from Cerulean alone and Cerulean with Citrine, the reduction of the lifetime was not significant, suggesting no interaction between Cerulean and Citrine. On the other hand, coexpression of Citrine-PHD1 reduced the lifetime of the donor fluorescent protein, Cerulean-ATF4, by 0.25 ns, representing the direct interaction between ATF4 and PHD1 (Table 1). The lifetime of Cerulean-ATF4 was also reduced by the co-expression of Citrine-PHD3 (Table 1). These data strongly suggested that ATF4 directly bound to PHD1 and PHD3 in living cells.
Mapping the domain in ATF4 binding to PHD1 To analyze the region of ATF4 required for the interaction with PHD1, we constructed several deletion mutants based on its reported domain structure (Fig. 2A), and applied them to the immunoprecipitation assay. The result demonstrated that the central region (91–277) of ATF4 corresponds to the interaction domain with PHD1 (Fig. 2B). Both p300 interaction site (1–90) and bZIP domain (278–349) were not required for the interaction with PHD1. Interestingly, both regions, 1–180 and 181–349, were co-precipitated with PHD1 suggesting that there are multiple interaction sites in
Table 1 – Lifetime data of fluorescent proteins expressed in living HEK293T cells.
Fluorescent protein channel
Lifetime / ns Reduction / ns
Cerulean Citrine Cerulean + Citrine
Tc1 : 1477 Tc2 : 1338 Tc1 : 1447 Tc2 : 1471
Tc1 : 3.61 Tc2 : 3.27 Tc1 : 3.53 Tc2 : 3.59
Cerulean-ATF4 Citrine-PHD3 Cerulean-ATF4 + Citrine-PHD3
Tc1 : 1389 Tc2 : 1280 Tc1 : 1257 Tc2 : 1396
Tc1 : 3.39 Tc2 : 3.13 Tc1 : 3.07 Tc2 : 3.41
Cerulean-ATF4 Citrine-PHD1 Cerulean-ATF4 + Citrine-PHD1
Tc1 : 1389 Tc2 : 1308 Tc1 : 1286 Tc2 : 1415
Tc1 : 3.39 Tc2 : 3.19 Tc1 : 3.14 Tc2 : 3.45
Tc1 : 0.08
Tc1 : 0.32
Tc1 : 0.25
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ATF4. However, we could not conclude which domain (Zipper II, ODD, and β-TrCP recognition motif) in that region is responsible for the interaction with PHD1. Another immunoprecipitation experiments also indicated that the endogenous PHD1 could interact with the central region (91–277) of ATF4 (Supplementary Fig. 8).
ATF4 does not serve as a substrate of PHD1/PHD3 Interaction between ATF4 and PHD1/PHD3 prompted us to evaluate whether ATF4 could serve as a substrate for PHD1 and/or PHD3. Sequence specificity for the enzymatic activity of the prolyl hydroxylases reported that the alanine residue in front of the hydroxylated proline is necessary for substrate recognition and catalysis [35]. There are three alanine–proline dipeptide sequences in the amino acid sequence of ATF4, and we regarded these proline residues (P124, P155, P241) as candidates of the target for the hydroxylation with PHD1 or PHD3. Therefore, we substituted each proline residue to alanine and those substitution mutants were applied to the mammalian two-hybrid assay to assess the effect of the substitution on the binding activity. A previous report described that the P564A mutation in the HIF-1α oxygen-dependent degradation domain essentially abolished the interaction between PHDs and HIF-1α [35] suggesting that the alanine substitution at the target residue for the prolyl hydroxylation leads to a reduction of the luciferase activity. Substitutions of the proline residues for alanine did not impair the activity binding to PHD3 (Fig. 3A) or PHD1 (Fig. 3B) when compared with the wild type. These results suggested that those proline residues were unable to be hydroxylated. Furthermore, the luciferase activity was not reduced by the alanine mutation of P213 (Fig. 3B), whose hydroxylation may affect key phosphorylation at S218 and S223 in ATF4 and subsequent binding of β-TrCP as an E3 ubiquitin ligase [36]. To investigate the effect of those substitutions to the stability, wild type or mutant ATF4-Flag proteins were expressed in HEK293T cells with or without coexpression of PHD3-Flag for 24 h (Fig. 3C). The result indicated that there were no significant differences in the expression level between wild type and four mutants, suggesting that those proline residues are not associated with the proteasomal degradation subsequent to the prolyl hydroxylation. Furthermore, P3A (P124A, P155A, P241A) mutant was constructed and applied to the immunoblot analysis (Supplementary Fig. 2). The result indicated that the substitutions of those three proline residues did not affect to the stability of ATF4. Previous report documented that PHD3 destabilizes ATF4, however, there was no direct evidence for the prolyl hydroxylation of ATF4 with PHD3 [37]. Our results suggested that the proline residues in the alanine–proline sequences are not the targets for the prolyl hydroxylation (Figs. 3A and B). Therefore, we investigated whether PHD1 or PHD3 directly hydroxylates ATF4 using in vitro 1-[14C]-CO2 capture assay [29,30]. This assay measures the release of 1-[14C]-CO2 from 1-[14C]-2-oxoglutarate (2-OG) to evaluate the activity of 2-OG dependent oxygenases including HIF-prolyl hydroxylases. PHD1 or PHD3 was expressed in E. coli and purified as described previously [27]. Hexahistidine (H6)tagged ATF4 was expressed in E. coli and purified as nearly homogeneity to examine as a substrate (Fig. 3D). We used HIF-1α NAD (N-terminal activation domain) as a positive control substrate [27], and erabutoxin b that has 4 proline residues in 62 residues long as a negative control substrate. As shown in Fig. 3E, both purified PHD1 and PHD3 exhibited significant prolyl hydroxylation activity toward HIF-1α NAD, indicating that purified PHD1 and
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p300 interaction Zipper II site
A
β-TrCP ODD recognition motif
bZIP
WT 91 - 349 181 - 349 278 - 349 1 - 277 1 - 180 91 - 277 181 - 277
B
PHD1-Flag
–
+
+
3Myc-ATF4
–
–
WT
WB : anti-Flag
+
+
+
91-
181- 278-
349
349
349
+
+
+
+
1-
1-
91-
181-
277
180
277
277 (kDa) 50 37
Input WB : anti-Myc
75 50 37 25 20 15
IP : anti-Flag WB : anti-Myc
75 50 37 25 20 15
Fig. 2 – (A) Schematic representation of the ATF4 structure. Location of the p300 interaction site, Zipper II domain, oxygen-dependent degradation domain (ODD), β-TrCP recognition motif, and basic and leucine zipper domain (bZIP) is indicated at the top. Deletion mutants used were exhibited at the bottom. (B) Interaction of PHD1 with ATF4 derivatives. Flag-tagged PHD1 expressing plasmid was co-transfected with plasmids expressing 3Myc-tagged ATF4 deletion mutants in HEK293T cells. Cell lysates were subjected to immunoprecipitation (IP) with anti-Flag antibody. Co-precipitates were applied to SDS-PAGE and detected with immunoblot analysis using anti-Myc antibody.
PHD3 possess significant and comparable enzymatic activity. On the other hand, prolyl hydroxylation activity toward ATF4 was quite low and comparable with the activity toward erabutoxin b. These results indicated that ATF4 did not serve as a direct substrate of PHD1 or PHD3.
The effect of the interaction to the stability of PHD1 and PHD3 We noticed that the expression level of PHD3 was decreased when ATF4 was coexpressed (Fig. 3C). Therefore, we explored whether the stability of ATF4, PHD1, and PHD3 was influenced by their interactions. Three PHD isozymes were expressed in HEK293T cells with or without the expression of ATF4. The expression level of PHD3 was diminished by the coexpression of ATF4 (Fig. 4A) that is consistent with the result shown in Fig. 3C. On the other hand, PHD1 and PHD2 expression levels remained constant regardless
of the expression of ATF4 (Fig. 4A). Further, we examined whether the reduction of PHD3 by ATF4 was caused by the lysosomal or proteasomal degradation. Although E-64 and pepstatin A, inhibitors for lysosomal degradation, did not affect the expression level of PHD3, a proteasome inhibitor, MG132, recovered the expression level of PHD3 under the presence of ATF4, suggesting that ATF4 stimulated the proteasomal degradation of PHD3 (Fig. 4A). We investigated the effect of the interaction between ATF4 and PHD1 or PHD3 on their stability in the presence of cycloheximide. PHD1 was quite stable and ATF4 exhibited no effects on the stability of PHD1 (Fig. 4B). In contrast, PHD3 was degraded rapidly and the coexpression of ATF4 accelerated the degradation (Fig. 4C). A previous report described that PHD3 stimulates the degradation of ATF4 [37]. Therefore, we examined the effect of the expression of PHD1 on the stability of ATF4. Surprisingly, the coexpression with PHD1 increased the lifetime of ATF4 protein (Fig. 4D), resulting in opposite effect on ATF4 stability.
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A
B
800
60
Relative luciferase activity
Relative luciferase activity
70
50 40 30 20 10
700 600 500 400 300 200 100
0 GAL4DBD-PHD3
–
+
+
+
+
0 GAL4DBD-PHD1
–
+
NLS-VP16-ATF4
–
WT
P124A
P155A
P241A
NLS-VP16-ATF4
–
WT
C
ATF4-Flag PHD3-Flag
– –
WT +
–
+
+
+
+
+
P124A P155A P213A P241A
P124A P155A P213A P241A –
+
–
+
–
+
–
+
(kDa)
55
ATF4
43 WB : anti-Flag 34 PHD3 26 Actin
(kDa) 170 130 95 72
1
E
2
55 43 H6-ATF4 34 26 17
Prolyl hydroxylase activity (cpm)
D
43
3000 PHD1 PHD3
2500 2000 1500 1000 500 0
HIF-1α-NAD
ATF4
Erabutoxin b
Fig. 3 – (A) Mammalian two-hybrid assay for detection of the interaction between PHD3 and ATF4 containing substitution of proline for alanine. Wild type or a substitution mutant of ATF4 fused to NLS-VP16 was coexpressed with the bait GAL4DBD-PHD3 in HepG2 cells. The luciferase activity of each sample was shown as relative ratio compared to the activity using GAL4DBD and NLS-VP16 as a standard. (B) Mammalian two-hybrid assay for detection of the interaction between PHD1 and ATF4 containing substitution of proline for alanine. Wild type or a substitution mutant of ATF4 fused to NLS-VP16 was co-expressed with the bait GAL4DBD-PHD3 in HepG2 cells. The luciferase activity of each sample was shown as a relative ratio compared to the activity using GAL4DBD and NLS-VP16 as a standard. (C) Immunoblot analysis of the expression level of wild type or a substitution mutant ATF4 in the presence or absence of PHD3. Flag-tagged ATF4 variants were expressed with or without Flag-tagged PHD3 in HEK293T cells. Cell lysates were subjected to SDS-PAGE and immunoblot analysis probed with anti-Flag antibody. Endogenous actin was detected with anti-actin monoclonal antibody (clone C-2; Santa Cruz Biotechnology) for a loading control. (D) Purification of His-tagged ATF4. His-tagged ATF4 expressed in E. coli was purified using Ni-affinity column twice. Purity of His-tagged ATF4 was analyzed by SDS-PAGE and Coomassie staining. Elution at the 1st step of Ni-affinity chromatography (lane 1). Elution at the 2nd step of Ni-affinity chromatography (lane 2). (E) Comparison of the prolyl hydroxylation activity of PHD1 and PHD3 using the method of the hydroxylation-coupled decarboxylation of 1-[14C]-2-oxoglutarate. The enzymatic activity was represented as the radioactivity derived from 1-[14C] CO2 which was release from 1-[14C]-2-oxoglutarate in the prolyl hydroxylation-dependent manner. Values represent mean ± s.d. for at least three measurements.
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Both PHD1 and PHD3 repress the transcriptional activity of ATF4 We investigated the influence of PHD1-3 on the transcriptional activity of ATF4. It has been reported previously that CHOP (GADD153) is one of the genes whose expression is regulated by ATF4 [14,38]. CHOP promoter containing the amino acid response
A
element (AARE), which is the binding site of ATF4, was fused to the luciferase reporter gene to examine the transcriptional activity of ATF4. The reporter activity was upregulated by the expression of ATF4 about 2-fold higher than the basal activity in HeLa cells. Both PHD1 and PHD3 abolished the ATF4-dependent transcriptional activity, although PHD2 had no effect (Figs. 5A and B). Similar results were also obtained in HepG2 cells (Supplementary Fig. 3).
B PHD1-Fla g –
3Myc-ATF4
+
PHD2-Fla g –
PHD3-Flag
+
–
+
PHD1-Flag
+
+
+
+
+
+
+
+
+
+
3Myc-ATF4 CHX (h)
– 0
– 2
– 4
– 8
– 24
+ 0
+ 2
+ 4
+ 8
+ 24
WB : anti-Flag
WB : anti-Flag WB : anti-Myc
–
–
–
+
–
MG132
– – –
– + –
– + +
– + +
+ + +
PHD3-Flag 3Myc-ATF4
Relative PHD1 levels (%)
160 E-64 + Pepstatin A
WB : anti-Flag
WB : anti-Myc
PHD1 PHD1+ATF4
140 120 100 80 60 40 20 0 0
5
10
15
20
25
CHX treatment time (h)
C
D
PHD3-Flag
+
+
+
+
+
+
+
+
+
+
3Myc-ATF4 CHX (h)
– 0
– 2
– 4
– 8
– 24
+ 0
+ 2
+ 4
+ 8
+ 24
WB : anti-Flag
PHD1-Flag
–
–
–
–
+
+
+
+
3Myc-ATF4 CHX (h)
+ 0
+ 2
+ 4
+ 8
+ 0
+ 2
+ 4
+ 8
WB : anti-Myc
PHD3 PHD3+ATF4
100
Relative ATF4 levels (%)
Relative PHD3 levels (%)
120
80 60 40 20
120
ATF4 ATF4+PHD1
100 80 60 40 20 0 0
2
4
6
8
10
CHX treatment time (h)
0 0
5
10
15
20
25
CHX treatment time (h)
Fig. 4 – (A) Immunoblot analysis of the expression level of PHDs in the presence or absence of ATF4. (Top panel) Flag-tagged PHDs were expressed with or without 3Myc-tagged ATF4 in HEK293T cells. Cell lysates were subjected to SDS-PAGE and immunoblot analysis probed with anti-Flag or anti-Myc antibody. (Bottom panel) Cells were treated with proteasomal inhibitor MG132, or lysosomal inhibitor E-64 + Pepstatin A to detect the degradation pathway of PHD3 in the presence of ATF4. (B) The stability of PHD1 in the presence or absence of ATF4 was analyzed. HEK293T cells were transfected with PHD1-Flag and 3Myc-ATF4 expressing plasmids. At 24 h later, cells were treated with 10 μg/ml of cycloheximide to inhibit de novo protein synthesis. After the indicated time points, cells were lysed and the expression level of PHD1 was detected by immunoblot analysis using anti-Flag antibody. The top panel exhibits the results of one experiment. Graphic representation of quantitated levels of PHD1 from three independent experiments is shown at the bottom panel. (C) The stability of PHD3 in the presence or absence of ATF4 was analyzed. (D) The stability of ATF4 in the presence or absence of PHD1 was analyzed. ATF4 was detected by immunoblot analysis using anti-Myc antibody.
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Furthermore, we examined the effect of PHD1-3 on the transcriptional activity of endogenously induced ATF4 using tunicamycin, an ER stress inducer. Treatment with tunicamycin for 16 h enhanced the CHOP promoter about 3-fold, and both PHD1 and PHD3
B
Relative luciferase activity
3.0 2.5 2.0 1.5 1.0 0.5 0.0 – + – –
+ + – –
– – + –
+ – + –
+ – – +
* **
2.5 2.0 1.5 1.0 0.5 0.0 pBOS
PHD1
PHD2
3.5
5.0
Tunicamycin
4.5
3.0 2.5 2.0 1.5 1.0 0.5 – – –
– – –
+ – –
+ – –
– + –
– + –
– – +
– – +
2.5 2.0 1.5 1.0 0.5 0.0
** **
4.0 3.5 3.0 2.5 2.0 1.5 1.0 0.5 0.0 pBOS
F Normoxia Hypoxia CoCl2
PHD3
** : p<0.01
DMSO
3.0
Relative luciferase activity
– – – +
3.0
D
4.0
0.0 PHD1-Fla g PHD2-Fla g PHD3-Fla g
E
+ – – –
PHD1
PHD2
3.0
Relative luciferase activity
Relative luciferase activity
C
– – – –
Fold induction (Tu/DMSO)
ATF4-Fla g PHD1-Fla g PHD2-Fla g PHD3-Fla g
** : p<0.01, * : p<0.05
Fold induction (ATF4[+]/ATF4[–])
A
significantly repressed the reporter activity (Figs. 5C and D). On the other hand, PHD2 did not inhibit the induction by the tunicamycin treatment. This repression was also observed in HeLa cells in hypoxia (1% O2) or chemical hypoxia caused by CoCl2, suggesting
PHD3
Normoxia Hypoxia CoCl2
2.5 2.0 1.5 1.0 0.5 0.0
ATF4-Flag
–
+
–
+
ATF4-Flag
–
+
–
+
PHD1-Flag
–
–
+
+
PHD3-Flag
–
–
+
+
Fig. 5 – (A) Repression of transactivation activity of ATF4 by the expression of PHDs. CHOP promoter-Luc was cotransfected into HeLa cells with ATF4 and PHDs expression plasmids. 16 h after transfection, cell lysates were extracted and assayed for luciferase activity. The luciferase activity was normalized by β-galactosidase activity to adjust the transfection efficiency. (B) From the results of A, the transactivation ratio by the expression of ATF4 in the presence or absence of PHDs was calculated and shown. Asterisks mark statistically significant differences. (C) Repression of tunicamycin-induced activation of CHOP promoter-Luc by the expression of PHDs. CHOP promoter-Luc was cotransfected into HeLa cells with PHDs expressing plasmids. Cells were treated with 2 μg/ml of tunicamycin or DMSO as a vehicle for 16 h, followed by the preparation of cell lysates. (D) From the results of C, the induction of the luciferase activity by the administration of tunicamycin was calculated and shown. (E) The effects of hypoxia or CoCl2 treatment to the repression of ATF4 by the expression of PHD1. After transfection, HeLa cells were treated with 21% O2 (normoxia, gray bars), 1% O2 (hypoxia, white bars), and 100 μM CoCl2 (black bars) for 16 h. (F) The effects of hypoxia or CoCl2 treatment to the repression of ATF4 by the expression of PHD3.
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that prolyl hydroxylase activity of PHD1/PHD3 may not be required for the suppression (Figs. 5E and F). To clarify that the prolyl hydroxylase activity of PHD1/PHD3 is not required for the suppression of ATF4 transactivation, we constructed three substitution mutants, PHD3(R205K), PHD1(H358A), and PHD1(R367K), all of which are catalytically inactive. PHD3(R205K) cannot bind to 2-OG but it still retains the ability to interact with ATF4 in living cells using FRET analysis (Supplementary Fig. 1D and Supplementary Table 1). Reporter assay using CHOP-promoter revealed that the expression of PHD3(R205K) significantly suppressed ATF4-dependent transcriptional activity indicating that the prolyl hydroxylase activity of PHD3 is not required for the suppression (Supplementary Fig. 4). Mammalian two-hybrid assay exhibited that PHD1(H358A) completely lost the ability to interact with ATF4 and PHD1(R367K) showed reduced but significant interaction with ATF4 (Supplementary Fig. 6). Reporter assay indicated that PHD1(H358A) did not inhibit the ATF4 transactivation, but PHD1(R367K) suppressed the transcriptional activity of ATF4 (Supplementary Fig. 5). Furthermore, the effect of the suppression of each wild type or the mutant PHD1/PHD3 to the tunicamycin-induced endogenous ATF4 is similar to the effect to the expressed ATF4 (Supplementary Fig. 7). These results indicated that the prolyl hydroxylase activity of PHD1/PHD3 is not required for the suppression of ATF4 transactivation. Taken together, it is suggested that the interaction between ATF4 and PHD1/PHD3 is responsible for the repression of the transcriptional activity of ATF4.
Discussion Our results clearly indicated that both PHD1 and PHD3 interact with ATF4, whereas PHD1 and PHD3 hydroxylate no proline residues of ATF4. It has been described previously that PHD3 interacts with ATF4 and regulates the stability of ATF4 [37]. They revealed that DMOG, a PHD inhibitor, or silencing of PHD3 by siRNA stabilizes ATF4. On the other hand, they didn't detect the interaction between PHD1 and ATF4 using the yeast two-hybrid system. The inconsistency between two studies may be caused by the difference in the assay system. We have corroborated the interaction using three different types of binding experiments. It is interesting that the interaction of PHD1 or PHD3 with ATF4 showed opposite effects on the stability of ATF4. Meanwhile, in vitro 1-[14C]-CO2 capture assay revealed that both PHD1 and PHD3 have no activity to hydroxylate ATF4. These results suggest that one of the unidentified factors, which regulate the stability of ATF4, might be the target molecule with prolines hydroxylated by PHD3. It has been described previously that the stability of ATF4 is dominantly regulated by β-TrCP, an F-box protein of the E3 ubiquitin ligase family [39]. Phosphorylation of ATF4 is necessary for the specific binding of β-TrCP (transducin repeatcontaining protein), and subsequently ATF4 is ubiquitinated and rapidly degraded by proteasome [36]. However, recent report revealed that β-TrCP is not responsible for the degradation of ATF4 in normoxia [40]. Thus it is conceivable that another pathway to regulate the degradation of ATF4 may be controlled by PHD3 in an oxygen-dependent manner. Further studies should be carried out to resolve the mechanism of the oxygendependent degradation of ATF4. In addition, destabilization of PHD3 by the coexpression of ATF4 was observed indicating that ATF4 and PHD3 accelerate each other's degradation. It is likely
that the coexpression of ATF4 and PHD3 is harmful to the cellular surviving. ATF4 is induced by several stresses including severe hypoxia or anoxia followed by ER stress, oxidative stress, and amino acid deprivation [13,39]. These stresses are associated with the solid tumor microenvironment, and cancer progression is tightly depended on the response to the microenvironment in which the expression of ATF4 is induced [11,12]. ATF4 transactivates a variety of genes involved in amino acid metabolism, redox chemistry, mitochondrial function, and apoptosis [41]. Recent study has been described that ATF4 is essential for xenograft tumor growth and the suppression of ATF4 blocks tumor growth in vivo [19]. Thus, ATF4 induced in the tumor microenvironment may be the clinical target for cancer therapy. Interestingly, Erez et al. reported that the ectopic expression of mouse PHD1 inhibits tumor growth [42]. It is noteworthy that ATF4 also positively regulates VEGF expression [43,44]. Thus, it is conceivable that the suppression of ATF4 by the ectopic PHD1 via direct interaction could be regarded as another pathway to inhibit the tumor growth. Whether the inhibition of ATF4 by PHD1/PHD3 could be exploited in cancer therapy needs to be considered. Supplementary materials related to this article can be found online at doi:10.1016/j.yexcr.2011.09.005.
Acknowledgments We thank Shigeyuki Nakada and Hironori Hayashi (Mitsui Engineering & Shipbuilding, Tamano, Japan) for technical assistance.
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