The Ribosomal Protein rpL11 Associates with and Inhibits the Transcrip
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《毒物学科学杂志》
Department of Veterinary Sciences and Center for Molecular Toxicology and Carcinogenesis, Pennsylvania State University, University Park, Pennsylvania 16802
3 To whom correspondence should be addressed at Center for Molecular Toxicology and Carcinogenesis, 201 Life Sciences Building, Pennsylvania State University, University Park, PA 16802. Fax: (814) 863-1696. E-mail: jpv2@psu.edu.
ABSTRACT
Peroxisome proliferator-activated receptor alpha (PPAR) is a member of the nuclear receptor superfamily whose ligands, the peroxisome proliferators (PPs), are liver tumor promoters in rodents. Interaction cloning was performed using bacterially expressed PPAR to identify proteins involved in PP signaling. The ribosomal protein L11 (rpL11), a component of the large 60S subunit, was identified as a PPAR-associated protein. Since rpL11 is a regulator of p53 and the cell cycle, the association between this protein and PPAR was examined in detail. PPAR-rpL11 interaction was confirmed using yeast and mammalian two-hybrid systems as well as in vitro pull-down assays. The association with rpL11 occurs within the D-domain (hinge-region) of PPAR. Unlike PPAR, the two closely related isoforms PPAR and do not interact with rpL11. Cotransfection of mammalian cells with rpL11 resulted in ligand-dependent inhibition of transcriptional activity of PPAR. Ribosomal protein L11–mediated inhibition of gene expression is associated with decreased binding to the PPAR-response element (PPRE) DNA sequence. Release of rpL11 from the ribosome by serum deprivation or low-dose actinomycin D did not dramatically affect PPRE-driven luciferase activity when PPAR was overexpressed by cotransfection. However, when endogenous levels of PPAR are examined and rpL11 concentration is manipulated by expression by small interference RNA, the ability of peroxisome proliferator to induce PPRE-driven reporter activity and target gene mRNA is affected. These studies show that rpL11 inhibits PPAR activity and adds further evidence that ribosomal proteins play roles in the control of transcriptional regulation.
Key Words: peroxisome proliferator-activated receptor; transcription regulation; protein–protein interaction; ribosomal protein rpL11.
INTRODUCTION
Peroxisome proliferator-activated receptor alpha (PPAR) is a member of the nuclear receptor (NR) superfamily. NRs are ligand-activated, intracellular receptors that include the progesterone (PR) and glucocorticoid receptors (GR), as well as the thyroid hormone (TR) and retinoic acid and retinoid X receptors (RAR and RXR) (Tsai and O'Malley, 1994). Multiple isoforms of the receptor (, , and ) have been cloned from various species including humans (Schmidt et al., 1992), rodents (Gottlicher et al., 1992; Issemann and Green, 1990) and amphibians (Dreyer et al., 1992). PPAR mediates the activity of peroxisome proliferators (PPs) by altering the expression of a wide variety of genes, including those involved in the peroxisomal -oxidation of fatty acids (Zhang et al., 1992). Despite the coexpression of the PPAR subtypes in various tissues (Lemberger et al., 1996), studies with a PPAR-null mouse have shown that this subtype alone is critical in the tumor-promoting activity of PPs (Peters et al., 1997, 1998). It is generally believed that PP-induced tumorigenesis is the result of altered gene expression, in particular the growth regulatory genes such as c-Myc (Belury et al., 1998; Vanden Heuvel, 1999; Vanden Heuvel et al., 1998) or oxidative enzymes such as acyl-CoA oxidase (ACO (Okamoto et al., 1997)), in target cells.
PPAR regulates gene expression in a PP-dependent manner by recognizing and binding to peroxisome proliferator-responsive elements (PPRE) in target genes which are composed of TGACCT-related direct repeats separated by one nucleotide (DR1) (Tugwood et al., 1992; Zhang et al., 1992). PPAR binds to PPREs as a heterodimer with RXR, the receptor for 9-cis-retinoic acid (Kliewer et al., 1992; Marcus et al., 1993). Genes transcriptionally regulated by PPs in a PPRE-dependent manner include ACO, liver fatty acid-binding protein (L-FABP), cytochrome P450 IVA1 (CYP4A1) (Auwerx et al., 1996; Schoonjans et al., 1996), and angiopoietin like protein-4 (Angplt4) (Kersten et al., 2000).
The transcriptional activity of this heterodimer is regulated in part by the association with coactivators, a family of proteins that are known to positively regulate NR activation (Dowell et al., 1997; Krey et al., 1997; Zhu et al., 1996). Zhu et al. cloned a novel coactivator, PPAR-binding protein (PBP) (Zhu et al., 1997), using a yeast two-hybrid system. PBP also associated with TR1, RAR, and RXR. In addition, PBP contains two LXXLL motifs, an element common to several NR coactivators (Heery et al., 1997), suggesting that PBP is a general NR coactivator. Recently, we have demonstrated that PPAR and PPAR associate with the coactivator CBP/p300 interacting transactivator with ED-rich tail 2 (CITED2, also called p35srj/mrg-1/msg-1) (Tien et al., 2004). Zamir et al. (1997) have demonstrated that PPAR associated with two corepressors, N-CoR and SMRT, which belong to a family of proteins that inhibit NR activity.
Although much is known about the basic mechanism by which PPARs regulate gene expression at the transcriptional level, upstream signaling events are less well understood. The PP signal transduction pathway includes protein-PPAR interactions that occur prior to DNA binding. For example, rat deoxyuridine triphosphatase (dUTPase) was identified as a repressor of PPAR transcriptional activity (Chu et al., 1996). When associated with dUTPase, PPAR is unable to form a heterodimer with RXR. In addition, PPAR inhibits dUTPase activity, an enzyme which plays a role in maintaining DNA fidelity (Pyles and Thompson, 1994). Our lab has shown that PPAR associates with heat shock protein 90 (hsp90) and the hepatitis X associated protein 2 (XAP2) (Sumanasekera et al., 2003a,b). PPAR, but not PPAR or , is associated with hsp90, and this interaction inhibits PPAR activity (Sumanasekera et al., 2003a). In contrast, XAP2 is an inhibitor of all three PPAR subtypes (Sumanasekera et al., 2003b). Thus, it is evident that multiple protein–protein interactions may play an important role in regulating PPAR activity.
The goal of the present study was to identify novel PPAR-associated proteins (PAPs) that may affect PP signal transduction and PPAR activity. A rat hepatoma cDNA library screened with bacterially expressed PPAR identified several interacting proteins including the rat ribosomal protein L11 (rpL11). Protein expression of L11 is increased in cold-treated plants, and this protein is inactive in strains of bacteria resistant to thiostreptone (Handa et al., 2001; Zhang et al., 2001). A PPAR–L11 association adds to a growing list of transcription factor–ribosomal protein interactions which include TR/rpL7, PR/rpL7, and c-Jun/rpL18a (Wool, 1996; Wool et al., 1995). In addition, there is increasing evidence that ribosomal proteins play a significant role in cell cycle control. In addition to rpL11 (Bhat et al., 2004; Lohrum et al., 2003; Zhang et al., 2003), rpL5 (Dai and Lu, 2004) and rpL23 (Dai and Lu, 2004; Dai et al., 2004; Jin et al., 2004) affect the turnover of p53 via their ability to bind to and suppress the E3 ligase function of HDM2. In this study, the interaction of L11 with PPAR was investigated.
MATERIALS AND METHODS
Materials.
-32P-CTP, 35S-methionine (1000 Ci/mmol), 32P-ATP, and Hy-Bond ECL nitrocellulose were purchased from Amersham (Arlington Heights, IL). Wy-14,643 ([4-chloro-6-(2,3-xylindino)-2-pyrimidinylthio]acetic acid, CAS No. 50892–23–4, >98% pure) was purchased from Chemsyn Science Laboratories (Lenexa, KS). Bezafibrate, clofibric acid, and ciglitazone were purchased from Biomol (Plymouth Meeting, PA). DH5 bacterial cells were purchased from PGC Scientific (Frederick, MD). Media components were from Invitrogen (Carlsbad, CA). Fetal bovine serum (FBS) was purchased from HyClone Laboratories (Logan, UT). Restriction endonucleases, nucleotides, and TNT in vitro transcription/translation kits were obtained from Promega (Madison, WI). Plasmid purification kits were purchased from Qiagen (Chatsworth, CA). Primers for PCR were designed using Lasergene (DNASTAR, Madison, WI) and purchased from Genosys (The Woodlands, TX) or Operon (Alameda, CA). NuSieve 3:1 agarose was obtained from FMC Bioproducts (Rockland, ME). Other chemicals and reagents were of the highest grade readily available.
Plasmids.
pFR-luciferase (UAS luciferase, Gal4 response element) was purchased from BD Biosciences Clontech (Palo Alto, CA). pRL/TK Renilla (phRL-TK), pRL/CMV Renilla (phRL/CMV), pSV/-Gal, and pGEM-T Easy were purchased from Promega (Madison, WI). The construction of pBK-CMV/rPPAR-FLAG and the mammalian two-hybrid pM/PPAR constructs was described previously (Sumanasekera et al., 2003a). The PPAR response element reporter, pACO(-581/-471)G.Luc was supplied by Dr. Jonathon Tugwood (Zeneca Pharmaceuticals, Central Toxicology Laboratories, Maccelsfield, UK) and has been previously described (Tugwood et al., 1998). Maltose binding protein- PPAR (MBP/PPAR) and MBP/rpL11 were generated by subcloning full-length rat cDNA PPAR (a gift from Dr. Frank Gonzalez, National Cancer Institute, Bethesda, MD) and rat ribosomal protein rpL11 (a gift from Dr. Ira Wool, University of Chicago, IL) into pMAL-c2X (New England Biolabs, Beverly, MA). pcDNA3.1/V5-His-rPPAR was generated by cloning into pcDNA3.1/V5-His-A (Invitrogen). VP16/rpL11 was generated by cloning rat rpL11 into pVP16 (Clontech).
cDNA library construction and screening.
FaO rat hepatoma cells (a generous gift from M.C. Weiss, Pasteur Institute, Paris, France), were grown in DMEM (Sigma) supplemented with 5% FBS, 100 units/ml of penicillin and 10 μg/ml of streptomycin, as previously described (Sterchele et al., 1996). Total RNA was isolated from FaO cells at 75% confluency using Tri Reagent (Molecular Research Center, Cincinnati, OH) and poly(A) mRNA extracted using oligo(dT)-magnetic beads (Promega). A directional cDNA library was constructed using the SuperscriptTM Lambda System (Gibco BRL) using the manufacturer's protocols. The yield and purity of the library construction was monitored at each step by incorporating -32P-CTP into the cDNA and removing small aliquots. The average transcript size of the library is 3000 nucleotides (range 1–9 kb).
PPAR-associated proteins (PAPs) were identified by screening the cDNA expression library using an overlay assay described previously (Chapline et al., 1993). Approximately 5 x 104 plaque forming units (pfus) were plated on each of 20 agar (150 mm) plates. The plates were incubated for 3 h at 37°C, after which isopropyl--D-thiogalactopyranoside (IPTG)-soaked nitrocellulose filters were overlaid, and incubation continued for an additional 4 h at 37°C. The orientation of the filters was marked, and they were removed from the plates and rinsed briefly with Tris-buffered saline (TBS), pH 7.4, containing 0.1% Tween 20 (TBS-tween). The membranes were blocked with 5% BSA (diluted in TBS-tween) for 1 h, followed by three 10-min washes with TBS-tween. The membranes were then probed with a bacterially-expressed PPAR-maltose binding protein (PPAR-MBP, see details below) for 2 h (diluted in TBS-tween plus 0.1% BSA). After three washes, the blots were incubated with an antibody to MBP (New England BioLabs, Beverly, MA) at the recommended dilutions. Detection of PPAR-MBP was performed by incubating the filters with an anti-rabbit IgG secondary antibody conjugated to alkaline phosphatase (Promega) and visualized with NBT (nitro blue tetrazolium)/BCIP (5-bromo-4-chloro-3-indolyl-phosphate). Positive plaques were eluted from agar plugs and were rescreened as described above. Positive clones from secondary and tertiary screens were excised out of Ziplox, and the inserts were sequenced in both directions by the Penn State Nucleic Acid Facility.
Transfection and reporter assays.
To confirm the interaction between PPAR and the isolated clones, Clontech's MatchMaker yeast (Y2H) and mammalian two-hybrid (M2H) systems were employed (Palo Alto, CA). Yeast SFY526 cells were transformed as per supplied protocols, and positive interactions were assessed by a -galactosidase (-gal) filter assay. Lipofectamine (Invitrogen) was used to cotransfect COS-1 cells (cultured in -MEM supplemented with 8% FBS and 100 units each of penicillin and streptomycin) with the M2H vectors and pFR/luciferase. Included in all transfections were pSV/-gal, pRL/CMV, or pRL/TK (Promega) to control for transfection efficiency. Luciferase enzymatic activity was measured with the single or dual luciferase enzyme assay system (Promega). -gal activity was measured using the -galactosidase enzyme assay system (Promega). For transient transfection experiments examining PPRE activity, pPPAR and pACO(-581/-471)G.Luc were cotransfected into COS-1 cells with or without either pCMX/RXR or pcD/rat rpL11 using Lipofectamine. Following treatment, cells were harvested, and renilla and firefly luciferase activities were examined using the Dual Luciferase Assay (Promega).
MBP pull-down assay.
PPAR or rpL11 were expressed as a maltose-binding protein (MBP) fusion protein and extracted from DH5 bacteria using amylose resin (New England Biolabs) as previously described (Riggs, 2000). Proteins were stored in "everything buffer" (20 mM Tris (pH 7.4), 200 mM NaCl, 1 mM EDTA, 10 mM -mercaptoethanol, and 1 mM sodium azide). In vitro translated PPAR or rpL11 were prepared using the TnT coupled reticulocyte lysate system (Promega) in the presence of 35S-labeled methionine (Amersham). Proteins were incubated in TBS containing 0.5% BSA and amylose resin on a rocker at 4C for 1 h. Afterward, the resin was washed five times with either 4 mM EDTA, 15 mM NaMOPS, 20 mM MOPs, 5 mM sodium azide, and 10% glycerol (precipitations A and B) or 150 mM NaCl, 1% CHAPs, and 5mg/ml BSA in everything buffer (precipitation C). The final pellet was resuspended in 2x SDS-tris/glycine sample buffer, boiled, and subjected to SDS/PAGE. Autoradiography was performed on the dried gels.
Gel shift assays.
Both strands of binding PPAR response element (CN) (5' CAAAACTAGGTCAAAGGTCA 3') and a non-binding PPAR response element (MeD) (5'-GTGTTAGAGGGCACAGGTCC) were synthesized and annealed as previously described (Juge-Aubry et al., 1997). For the first gel shift, in vitro translations of PPAR (2 μl), RXR (2 μl), and combined rpL11 and nonprogrammed lysate (6 μl) were added to each reaction. For the second assay, 0.25 μl of RXR, 2.5 μg of either MBP-empty or MBP-PPAR, and 0–3 μg of MBP-L11 were added to each reaction (0, 0.5, 1, 1.5, 2, and 3 μg in lanes 1–6 and 7–12, respectively). All reactions were performed in 4% glycerol, 1 mM MgCl2, 0.5 mM EDTA, 0.5 mM DTT, 50 mM NaCl, 10 mM Tris/HCl (pH 7.5), 0.5% CHAPS, and either 125 ng (assay one) or 64 ng (assay two) poly-deoxy-inosinic-deoxycytidylic acid (poly dI:dC) in a volume of 10 μl (assay one) or 15 μl (assay two). All reactions were incubated for 15 min before and after adding 1 μl 32P-labeled response element (>50,000 cpm). Where indicated, 1 μl (25 μM) of unlabeled CN or MeD were added. The incubations were resolved on 6% DNA Retardation Gels (Invitrogen) using 0.5x TBE (5.4g Tris base, 2.75g boric acid, and 1mM EDTA).
Manipulation of rpL11 activity.
Hepa-1 cells were maintained in Dulbecco's modified Eagle's medium containing 8% FBS in a 37°C incubator with 5% CO2. For PPRE-dependent reporter assays, cells were grown to 60% confluence and transfected with HD-Luciferase (kind gift from D. J. Waxman, Boston University) which contains a functional PPRE from the peroxisomal enoyl-CoA hydratase/3-hydroxyacyl-CoA dehydrogenase gene. Subsequently, Hepa-1 cells were treated with DMSO (0.1%) or Wy-14,643 (50 μM) concurrently with two means to modulate rpL11 activity, actinomycin D (AD) or serum deprivation (SD) (Bhat et al., 2004; Perry and Kelley, 1970). AD was dissolved in DMSO and added to cells at a final concentration of 5 nM. Serum deprivation was achieved by washing the cells twice with PBS and adding DMEM media with 0.1% serum. For endogenous gene expression studies, 60% confluent cells were treated with DMSO or Wy14,643 and subjected to AD or SD treatment as above. In all cases, AD and SD were performed concomitantly with chemical treatments for 24 h. A double-stranded RNA inhibitor for rpL11 was designed using the sequence from GenBankTM NM_025919. The sequence chosen was as follows: 5'-AAGCATTGGGATCTACGGCCT-3'. The sequence was synthesized and cloned into pSuper.neo plasmid (Oligoengine, Seattle, WA). Transfection of the RNAi plasmid into Hepa-1 cells was performed using Lipofectamine (Invitrogen) transfection reagent according to the manufacturer's protocol for 12 h, followed by serum starvation or Actinomycin D and Wy-14,643 treatment as described above. A rabbit polyclonal antibody against a C-terminal L11 peptide was a kind gift of K. H. Vousden (Beatson Institute for Cancer Research, UK).
Real-time PCR.
Total RNA was isolated by TriReagent (Sigma) according the manufacturer's instructions. The total RNA was reverse transcribed using the ABI High Capacity cDNA archive kit (Applied Biosystems, Foster City, CA). Standard curves were made using serial dilutions from pooled cDNA samples. Real Time PCR was performed using the SYBR Greeen PCR Master Mix (Applied Biosystems) according to the manufacturer's protocol and amplified on the ABI Prism 7000 Sequence Detection System.
Statistical analysis.
Where indicated, MiniTab (State College, PA) was used to evaluate data for statistical significance using One-way ANOVA and Student's t-test with significance at p < 0.05.
RESULTS
Identification of PPAR-Associated Proteins
To identify cDNAs which encode PPAR-associated proteins (PAPs) we screened a FaO Ziplox expression library, using as a probe PPAR-MBP fusion protein expressed and purified from bacterial extracts. Visualization of positive plaques was accomplished immunologically by use of a monoclonal antibody to the MBP domain. On three separate experiments, 106 phage plaques were screened. From this primary screen, 15 positives were eluted from the agar, plated, and rescreened. Of those initial positives, 7 were confirmed with subsequent secondary and tertiary screens and were excised from the phage and sequenced (Table 1). DNA sequencing revealed high homology of several clones with published sequences. PAP2 and PAP4, determined to be homologous clones isolated from primary screens during two separate experiments, had inserts of 539 and 575 base pairs (bp), respectively. PAP2/4 is identical to the rat ribosomal protein rpL11 (Chan et al., 1992), which is part of the large 60S ribosomal subunit (Tsurugi et al., 1976). While PAP2/4 did not contain the complete open reading frame of rpL11, they did contain a continual stretch that corresponds to nucleotide number 22 until 552 (of the total length of 800 bp (Chan et al., 1992). PAP1 and PAP3 demonstrated significant homology with the rat DCT1 (a member of the Nramp family of genes (Gunshin et al., 1997) and CITED2 (homologous to human coactivator CITED2/p35srj (Bhattacharya et al., 1999), respectively. PAP5 matched well with ST7/RAY1/HELG protein (accession AJ277291), while another clone (PAP6) contains less than 200 bp and was not examined further. PAP7 contained the largest insert, and matched with the rat cysteine proteinase, cathepsin L (Ishidoh et al., 1987).
Confirmation of PPAR:PAP Interaction
In order to confirm that identified PAPs interact with PPAR in vivo, a yeast-based system was utilized. The PAPs were subcloned into the yeast expression vector, pGAD424, and the rat PPAR was inserted into pGBT9. When used to transform yeast, these vectors express a chimera of the activation domain and the DNA binding domain of the GAL4 transcription factor, respectively. Interactions were assessed by a -gal filter assay in which blue plaques would indicate expression of lacZ, and therefore an interaction between the two chimeras. -gal activity was qualitatively assessed and summarized in Table 1. When full-length PPAR is expressed by itself, there is substantial amount of reporter activity (see Table 1 legend). This is not surprising, since the amino-terminus of the receptor contains transcriptional activation functions (Tsai and O'Malley, 1994). To circumvent this background activity when using this system, other researchers have deleted the transactivation domain of other nuclear receptors (Burris et al., 1995). However, we screened the library with intact transcriptional activation domains and, therefore, performed the yeast two-hybrid assay with full-length PPAR. As indicated in Table 1, when yeast were cotransfected with various PAPs and PPAR, there was a decrease in reporter activity. As a positive control, human RXR, subcloned into pGAD424, was used with pGBT9.rPPAR to cotransfect yeast cells, and a decrease in reporter activity was also observed. Based on the reduction in -gal activity in the presence of PPAR and the PAPs (as compared to PPAR alone), all PAPs were confirmed to interact with PPAR. CITED2 (PAP3) is a bona fide PPAR coactivator and has been examined elsewhere (Tien et al., 2004). Due to the fact that other ribosomal proteins are known to functionally interact with nuclear receptors, and that rpL11 affects the cell cycle, the present studies focused on the PAP2/4 (rpL11) and PPAR association.
Full-Length rpL11 Inhibits PPAR Transactivation
The effect of PAP2/4 as well as full-length rpL11 on PPAR's ability to influence the expression of a reporter gene under the control of a natural PPRE was examined (Fig. 1). Vectors containing PPAR and rpL11 were cotransfected into COS-1 cells along with pACO(-581/-471)G.Luc. As indicated in Figure 1A, cotransfecting cells with increasing amounts of the rpL11 plasmid inhibited the transcriptional activity of PPAR in a dose-dependent manner. These results were dependent on PPAR, since PAP2/4 by itself had no effect on luciferase activity (data not shown). Similar results were obtained with the partial clones of rpL11 (PAP2/4, Fig. 1B) as well as the full-length rpL11 (Figure 1C) on pM/PPAR activity using the GAL4 reporter, although the amount of reduction in PPRE-driven luciferase activity was more pronounced.
L11 Interacts with the D-Domain of PPAR
We utilized the mammalian two-hybrid assay to map the interaction between PPAR and rpL11. Various domains of PPAR were subcloned in the pM vector and examined for their ability to interact with VP16/L11 (Fig. 2A). All data is compared to the amount of reporter activity in the presence of VP16/Empty under the same treatment conditions. The D-domain (amino acids 168–273) had the highest amount of positive interaction with rpL11. Full-length rPPAR in the presence of Wy14,643 and the E/F domain in the unliganded state showed a decrease in reporter activity in the presence of VP16/rpL11. The interaction of ribosomal protein L7a enhances the effects of partial agonists but not full agonists of the estrogen receptor (Jackson et al., 1997). To test whether this holds true for the PPAR/L11 association, several known PPAR activators were examined in the M2H system (Fig. 2B). Cotransfecting with rpL11 decreased the activation of PPAR by Wy14,643. Less efficacious ligands such as bezafibrate, ETYA, and clofibric acid (not shown) activated PPAR in the presence or absence of cotransfected rpL11. The specific inhibitor of PPAR, MK886 (Kehrer et al., 2001), was also unaffected by cotransfection with rpL11 (not shown).
PPAR Subtype Preference for rpL11 Interaction
The ability of rpL11 to affect the activity of PPAR, , and was examined using M2H assays (Fig. 3). PPAR was activated with ligands specific for each subtype in the presence or absence of VP16/rpL11. Wy14,643 significantly increased PPAR activity, while carbacyclin and PGJ2 significantly activated PPAR and , respectively. The ligand-dependent activation of PPAR was decreased by cotransfection with rpL11-containing plasmid. Neither basal nor ligand-induced PPAR or PPAR activity was affected by rpL11 transfection. Similar results were obtained when the partial sequence of rpL11 was used (PAP2/4) instead of the full-length construct (not shown).
PPAR Interacts with rpL11 in Vitro
PPAR was expressed in bacteria as a maltose binding protein (MBP) fusion protein and partially purified as described under Materials and Methods. PPAR-MBP was mixed with in vitro translated full-length rpL11, and the resultant complex was affinity purified using amylose resin. The protein complex was separated on SDS/PAGE, transferred to nitrocellulose, and rpL11 was detected by autoradiography. Full-length ribosomal rpL11 copurified with PPAR-MBP (Fig. 4A lane 2) but not MBP alone (lane 3). Lane 1 indicates the amount of in vitro translated rpL11 used in the pull-down experiments. The reverse incubation was also performed where rpL11 was expressed as an MBP fusion and PPAR was in vitro translated (Fig. 4B). Increasing amounts of PPAR were copurified in the presence of increasing rpL11-MBP. Finally, various domains of PPAR fused to MBP were examined for their ability to associate with in vitro translated rpL11 (Fig. 4C). As expected from the M2H assays, the full-length (FL) and D-domains of PPAR contained the predominant rpL11 binding capability.
rpL11 Inhibits PPAR Binding to the PPRE
The effect of rpL11 on DNA binding of the PPAR/RXR heterodimer was examined using in vitro translated or bacterially expressed proteins. In Figure 5A, PPAR, RXR, and rpL11 were in vitro translated and mixed with each other prior to the addition of 32P-labeled PPRE. PPAR/RXR binding to the 32P-labeled PPRE could be competed with cold CNPPRE but not with the MeD response element (lanes 2 versus 3 and 4). In lanes 5 through 9, increasing amounts of rpL11 in vitro translation was added to a constant amount of PPAR. To this complex, RXR was added, followed by 32P-labeled PPRE. With increasing amounts of rpL11, there was a decrease in the binding to the response element. Interestingly, if in vitro translated PPAR and RXR are mixed prior to the addition of rpL11, there was no decrease in DNA binding, indicating that rpL11 could not affect the preformed heterodimer's interaction with DNA. In the experiments depicted in Figure 5B, MBP (lanes 1–6) or MBP-PPAR (lanes 7–12) was incubated with increasing concentrations of MBP-L11 (lanes 2–6, 8–12). All reactions contained an equal amount of in vitro translated RXR and equal amounts of radiolabeled PPRE probe as described in Materials and Methods. The PPAR/RXR specific band is shown as band (B) and was decreased with increasing amounts of rpL11. At higher rpL11 concentrations, two shifted bands appear (A and C), which are not dependent upon PPAR for DNA binding and are possibly direct interactions of rpL11 with DNA. Even at lower rpL11 concentrations, this ribosomal protein can inhibit PPAR from binding to DNA, whether rpL11 is in vitro translated or bacterially expressed. This effect was not due to a change in the localization of PPAR, which was unaffected by rpL11 (data not shown).
PPAR Activity under Conditions of rpL11 Manipulation
The activity of endogenous PPAR or both endogenous and transfected PPAR in Hepa-1 cells was examined using a PPRE-reporter as well as real-time PCR. As shown Figure 6 (panels A and B), in the absence of exogenously supplied PPAR there is a modest induction of HD-luciferase activity (approximately 50%) by Wy14,643. The increase due to Wy14,643 addition is approximately two-fold in the case where PPAR cDNA is supplied. Actinomycin D and serum starvation have been shown to destabilize the ribosome, allowing for potential interactions between ribosomal proteins and other cellular components (Bhat et al., 2004; Perry and Kelley, 1970). Actinomycin D treatment in cells which did not receive exogenous PPAR increased PPRE-dependent activity roughly three-fold; this effect was absent when PPAR was cotransfected. In the presence of either AD or SD, the Wy14,643-mediated induction of HD-luciferase is no longer observed, but only when endogenous PPAR is examined. When exogenous PPAR is supplied, the Wy14,643-mediated response is still observed in the presence of AD or SD. Transfection with the rpL11 RNAi plasmid resulted in an increase in HD-luciferase activity regardless of treatment (Fig. 6, panels C and D). These experiments were performed in the absence of exogenously supplied PPAR, and Wy14,643 had marginal effects on reporter activity in this case.
Since AD and SD treatment affected PPRE-driven reporters in the absence of overexpressed PPAR, these conditions were used to examine effects on known target genes of this nuclear receptor. Similar results to the reporter assay were obtained when two PPAR responsive genes were examined, Angptl4 and ACO (Fig. 7). Treatment with Wy14,643 resulted in a slight increase in both of these mRNAs in the absence of AD (Panel A) or SD (Panel B) with either no observed increase or a reversal of effect in the presence of these treatments. In general, transfection of an RNAi plasmid targeted to rpL11 increased PPAR target gene expression relative to that observed in cells which received control plasmid (Panels C–F). The amount of rpL11 inhibition by the RNAi plasmid in transient transfection of these cultures was approximately 30%.
DISCUSSION
The mechanism by which NRs are able to alter the expression of target genes has been extensively studied. The simplest model was one in which, once activated by ligand, NRs bind as dimers (homo- or hetero-) to specific sequences in the 5' regulatory region of target genes (Tsai and O'Malley, 1994). Recent work, however, suggests this scheme is overly simplified. Transcriptional activation involves interplay between many cellular processes. NRs are regulated by phosphorylation (Weigel, 1996) and heat shock proteins (Pratt and Toft, 1997), as well as two important groups of transcriptional regulators known as coactivators and corepressors (Torchia et al., 1998).
Due to the increasing body of evidence that protein–protein interactions are key regulatory steps in the NR signal transduction pathway, we intended to identify novel PPAR-associated proteins. In this report we describe the identification of the ribosomal protein L11 as such a protein. Originally detected by screening a cDNA expression library constructed from FaO mRNA, this interaction was confirmed in yeast and mammalian two-hybrid systems (Table 1 and Fig. 1, respectively). Additionally, we were able to demonstrate biological relevance when it was observed that cotransfection of PPAR with the full-length rpL11 resulted in a decrease in the transcriptional activity of the receptor and a decrease PPAR/RXR heterodimerization. Furthermore, rpL11 directly interacted with PPAR in MBP pull-down assays.
Ribosomal protein rpL11 is part of the large 60S ribosomal subunit (Tsurugi et al., 1976), and homologues have been identified in prokaryotes as well as eukaryotes. This is not the first time that a ribosomal protein has been observed to interact with a transcription factor. Indeed, two ribosomal proteins, L10 and L18a, have been identified as interaction partners for c-Jun (Chan et al., 1996; Gramatikoff et al., 1995). Furthermore, Burris and colleagues (1995) reported that the thyroid hormone (TR) and retinoic acid receptors (RAR) associated with the human ribosomal protein L7a. These interactions resulted in an inhibition of transcriptional activity of the receptors. An interesting observation made by Jackson et al. (1997) was that the association of ER, GR, or PR with L7 increases in the presence of partial, but not full, agonists. These authors also mapped the site of protein–protein interaction to the D-domain of the receptors. In the present studies we showed clear preference for full PPAR agonists such as Wy14,643 relative to partial agonists such as bezafibrate or ETYA in their ability to stimulate PPAR/L11 interactions. Although the ligand preference for the PPAR/L11 association is dissimilar to that of ER/L7a, the NR domain utilized is the same, the D- or hinge-domain. This region is variable among the PPAR subtypes, which may help explain why rpL11 affected PPAR activity but not PPAR or .
The observations that L7 interacts with RAR, PR, ER, and GR, L10 with c-jun as well as those presented herein, point to the possibility of ribosomal proteins having extraribosomal functions (Wool, 1996). In fact, it is still unclear as to whether ribosomal proteins evolved specifically for the ribosome, or whether they were co-opted from existing proteins with defined functions. As of now there is only circumstantial evidence to suggest that these proteins existed initially for other purposes. First, many ribosomal proteins contain structural motifs, such as zinc fingers, leucine zippers, and helix-turn-helix motifs, which suggest that they have the capacity to bind to DNA (Rice and Steitz, 1989; Wool et al., 1995). Bacterial expressed rpL11 caused a DNA gel shift when present at high concentrations (Fig. 5B); whether this protein binds to DNA in vivo has not yet been determined. Second, ribosomal proteins interact with transcription factors such as c-Jun, TR, and RAR, as well as our own observation that rpL11 associates with and inhibits PPAR's ability to regulate PPRE-driven reporters (Figs. 6 and 7). Last, TR and PPAR are NRs, and this superfamily is believed to have evolved more that 500 million years ago in primitive organisms (Laudet, 1997; Sumida, 1995). Since many ribosomal proteins have prokaryotic homologues (Wool et al., 1995), one could imagine primitive organisms evolving to express ribosomal proteins as the initial negative regulators of ancestral transcription factors.
The biological importance of the interaction between PPAR and rpL11, as well as other NR-ribosomal protein interactions, remains unclear. It appears that rpL11 inhibits PPAR from forming a heterodimer with RXR, thereby preventing an active complex from binding to PPREs. Similarly, Burris et al. (1995) demonstrated that ribosomal protein L7a prevented the TR/RXR heterodimer from binding to TREs, but had no effect on RXR's ability to bind to its response element. On the other hand, these researchers did not indicate whether L7a was able to interact with TR or RXR in vitro. It is possible that ribosomal proteins are able to prevent the formation of certain NR/RXR heterodimers by competing for RXR. The fact that we were able to inhibit the activity of pM/PPAR, a construct that contains a heterologous DNA binding motif and does not require RXR interaction for activity, suggests that this inhibition of PPAR activity may have multiple mechanisms. In addition to inhibiting RXR heterodimerization, rpL11 may affect coactivator recruitment, for example. Sequestering of PPAR in the cytoplasm does not appear to occur, as we were unable to detect differences in the cytosol/nucleus ratio when rpL11 is overexpressed (data not shown); alteration of suborganelle distribution, such as nucleolar colocalization of rpL11 and PPAR has not yet been examined.
The potential significance of the PPAR/rpL11 interaction has increased as a result of observations of an rpL11/HDM2/p53 complex (Lohrum et al., 2003; Zhang et al., 2003). In fact, overexpressing the ribosomal protein is able to induce cell cycle arrest (Lohrum et al., 2003). The mechanism of the rpL11 control of the cell cycle is beginning to be understood. Ribosomal protein L11 can associate with HDM2, the ubiquitin ligase of p53, and inhibit the degradation of p53 (Lohrum et al., 2003). The HDM2/L11 association is enhanced by disruption of the ribosome by actinomycin-D (Lohrum et al., 2003; Zhang et al., 2003). It has been postulated that a signal stimulating cell growth would promote rpL11 assembly into the ribosome, allowing HDM2 to inhibit p53 activity. A signal for inhibition of cell growth, such as perturbation in ribosome biogenesis, would enhance HDM2 activity and, hence, increase p53 and growth arrest (Zhang et al., 2003). The modulation of p53 activity by a ribosomal protein is not exclusive to rpL11 (Bhat et al., 2004; Lohrum et al., 2003; Zhang et al., 2003) as rpL5 (Dai and Lu, 2004) and rpL23 (Dai and Lu, 2004; Dai et al., 2004; Jin et al., 2004) affect the turnover of p53 via their ability to bind to and suppress the E3 ligase function of HDM2. It is intriguing to speculate that a PPAR/L11 association may affect the formation of the rpL11/HDM2/p53 complex. We have recently reported that PP-induced PPAR activation was able to increase entry of hepatocytes into the cell cycle (Tien et al., 2003). Thus, we would hypothesize that mitogenic agents such as Wy14,643 would decrease p53 activity. Although speculative at this point, it is possible that unliganded PPAR may enter into the p53 complex through its association with rpL11.
In summary, the ribosomal protein rpL11 interacts with the D-domain of PPAR and inhibits PPRE-driven reporter activity. Manipulation of rpL11 activity by a variety of means affects PPAR activity, whether addressed using reporter assays or endogenous gene expression. Ribsomal protein L11 joins a growing list of ribosomal proteins that can affect gene transcription. Recent interest in rpL11 has resulted from observations of an apparent rpL11/HDM2/p53 complex. Thus, the interaction of rpL11 with PPAR, a nuclear receptor implicated in cell cycle control and carcinogenesis, has increasing importance.
NOTES
1 Current address: Rutgers University, Pharmacology & Toxicology, 170 Frelinghuysen Rd., Rm. 441, EOHSI Piscataway, NJ 08854.
2 Current address: Pfizer Global Research & Development, World Wide Safety Sciences, 800 N Linderberg Boulevard, Creve Coeur, MO 63167.
ACKNOWLEDGMENTS
The authors thank the technical assistance of Kristine Walker, Tecla Brabazon, and Nicole Agostino. The authors also thank the editorial assistance of Jerry Thompson. This work was supported by NIH ES007799 (JVH).
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3 To whom correspondence should be addressed at Center for Molecular Toxicology and Carcinogenesis, 201 Life Sciences Building, Pennsylvania State University, University Park, PA 16802. Fax: (814) 863-1696. E-mail: jpv2@psu.edu.
ABSTRACT
Peroxisome proliferator-activated receptor alpha (PPAR) is a member of the nuclear receptor superfamily whose ligands, the peroxisome proliferators (PPs), are liver tumor promoters in rodents. Interaction cloning was performed using bacterially expressed PPAR to identify proteins involved in PP signaling. The ribosomal protein L11 (rpL11), a component of the large 60S subunit, was identified as a PPAR-associated protein. Since rpL11 is a regulator of p53 and the cell cycle, the association between this protein and PPAR was examined in detail. PPAR-rpL11 interaction was confirmed using yeast and mammalian two-hybrid systems as well as in vitro pull-down assays. The association with rpL11 occurs within the D-domain (hinge-region) of PPAR. Unlike PPAR, the two closely related isoforms PPAR and do not interact with rpL11. Cotransfection of mammalian cells with rpL11 resulted in ligand-dependent inhibition of transcriptional activity of PPAR. Ribosomal protein L11–mediated inhibition of gene expression is associated with decreased binding to the PPAR-response element (PPRE) DNA sequence. Release of rpL11 from the ribosome by serum deprivation or low-dose actinomycin D did not dramatically affect PPRE-driven luciferase activity when PPAR was overexpressed by cotransfection. However, when endogenous levels of PPAR are examined and rpL11 concentration is manipulated by expression by small interference RNA, the ability of peroxisome proliferator to induce PPRE-driven reporter activity and target gene mRNA is affected. These studies show that rpL11 inhibits PPAR activity and adds further evidence that ribosomal proteins play roles in the control of transcriptional regulation.
Key Words: peroxisome proliferator-activated receptor; transcription regulation; protein–protein interaction; ribosomal protein rpL11.
INTRODUCTION
Peroxisome proliferator-activated receptor alpha (PPAR) is a member of the nuclear receptor (NR) superfamily. NRs are ligand-activated, intracellular receptors that include the progesterone (PR) and glucocorticoid receptors (GR), as well as the thyroid hormone (TR) and retinoic acid and retinoid X receptors (RAR and RXR) (Tsai and O'Malley, 1994). Multiple isoforms of the receptor (, , and ) have been cloned from various species including humans (Schmidt et al., 1992), rodents (Gottlicher et al., 1992; Issemann and Green, 1990) and amphibians (Dreyer et al., 1992). PPAR mediates the activity of peroxisome proliferators (PPs) by altering the expression of a wide variety of genes, including those involved in the peroxisomal -oxidation of fatty acids (Zhang et al., 1992). Despite the coexpression of the PPAR subtypes in various tissues (Lemberger et al., 1996), studies with a PPAR-null mouse have shown that this subtype alone is critical in the tumor-promoting activity of PPs (Peters et al., 1997, 1998). It is generally believed that PP-induced tumorigenesis is the result of altered gene expression, in particular the growth regulatory genes such as c-Myc (Belury et al., 1998; Vanden Heuvel, 1999; Vanden Heuvel et al., 1998) or oxidative enzymes such as acyl-CoA oxidase (ACO (Okamoto et al., 1997)), in target cells.
PPAR regulates gene expression in a PP-dependent manner by recognizing and binding to peroxisome proliferator-responsive elements (PPRE) in target genes which are composed of TGACCT-related direct repeats separated by one nucleotide (DR1) (Tugwood et al., 1992; Zhang et al., 1992). PPAR binds to PPREs as a heterodimer with RXR, the receptor for 9-cis-retinoic acid (Kliewer et al., 1992; Marcus et al., 1993). Genes transcriptionally regulated by PPs in a PPRE-dependent manner include ACO, liver fatty acid-binding protein (L-FABP), cytochrome P450 IVA1 (CYP4A1) (Auwerx et al., 1996; Schoonjans et al., 1996), and angiopoietin like protein-4 (Angplt4) (Kersten et al., 2000).
The transcriptional activity of this heterodimer is regulated in part by the association with coactivators, a family of proteins that are known to positively regulate NR activation (Dowell et al., 1997; Krey et al., 1997; Zhu et al., 1996). Zhu et al. cloned a novel coactivator, PPAR-binding protein (PBP) (Zhu et al., 1997), using a yeast two-hybrid system. PBP also associated with TR1, RAR, and RXR. In addition, PBP contains two LXXLL motifs, an element common to several NR coactivators (Heery et al., 1997), suggesting that PBP is a general NR coactivator. Recently, we have demonstrated that PPAR and PPAR associate with the coactivator CBP/p300 interacting transactivator with ED-rich tail 2 (CITED2, also called p35srj/mrg-1/msg-1) (Tien et al., 2004). Zamir et al. (1997) have demonstrated that PPAR associated with two corepressors, N-CoR and SMRT, which belong to a family of proteins that inhibit NR activity.
Although much is known about the basic mechanism by which PPARs regulate gene expression at the transcriptional level, upstream signaling events are less well understood. The PP signal transduction pathway includes protein-PPAR interactions that occur prior to DNA binding. For example, rat deoxyuridine triphosphatase (dUTPase) was identified as a repressor of PPAR transcriptional activity (Chu et al., 1996). When associated with dUTPase, PPAR is unable to form a heterodimer with RXR. In addition, PPAR inhibits dUTPase activity, an enzyme which plays a role in maintaining DNA fidelity (Pyles and Thompson, 1994). Our lab has shown that PPAR associates with heat shock protein 90 (hsp90) and the hepatitis X associated protein 2 (XAP2) (Sumanasekera et al., 2003a,b). PPAR, but not PPAR or , is associated with hsp90, and this interaction inhibits PPAR activity (Sumanasekera et al., 2003a). In contrast, XAP2 is an inhibitor of all three PPAR subtypes (Sumanasekera et al., 2003b). Thus, it is evident that multiple protein–protein interactions may play an important role in regulating PPAR activity.
The goal of the present study was to identify novel PPAR-associated proteins (PAPs) that may affect PP signal transduction and PPAR activity. A rat hepatoma cDNA library screened with bacterially expressed PPAR identified several interacting proteins including the rat ribosomal protein L11 (rpL11). Protein expression of L11 is increased in cold-treated plants, and this protein is inactive in strains of bacteria resistant to thiostreptone (Handa et al., 2001; Zhang et al., 2001). A PPAR–L11 association adds to a growing list of transcription factor–ribosomal protein interactions which include TR/rpL7, PR/rpL7, and c-Jun/rpL18a (Wool, 1996; Wool et al., 1995). In addition, there is increasing evidence that ribosomal proteins play a significant role in cell cycle control. In addition to rpL11 (Bhat et al., 2004; Lohrum et al., 2003; Zhang et al., 2003), rpL5 (Dai and Lu, 2004) and rpL23 (Dai and Lu, 2004; Dai et al., 2004; Jin et al., 2004) affect the turnover of p53 via their ability to bind to and suppress the E3 ligase function of HDM2. In this study, the interaction of L11 with PPAR was investigated.
MATERIALS AND METHODS
Materials.
-32P-CTP, 35S-methionine (1000 Ci/mmol), 32P-ATP, and Hy-Bond ECL nitrocellulose were purchased from Amersham (Arlington Heights, IL). Wy-14,643 ([4-chloro-6-(2,3-xylindino)-2-pyrimidinylthio]acetic acid, CAS No. 50892–23–4, >98% pure) was purchased from Chemsyn Science Laboratories (Lenexa, KS). Bezafibrate, clofibric acid, and ciglitazone were purchased from Biomol (Plymouth Meeting, PA). DH5 bacterial cells were purchased from PGC Scientific (Frederick, MD). Media components were from Invitrogen (Carlsbad, CA). Fetal bovine serum (FBS) was purchased from HyClone Laboratories (Logan, UT). Restriction endonucleases, nucleotides, and TNT in vitro transcription/translation kits were obtained from Promega (Madison, WI). Plasmid purification kits were purchased from Qiagen (Chatsworth, CA). Primers for PCR were designed using Lasergene (DNASTAR, Madison, WI) and purchased from Genosys (The Woodlands, TX) or Operon (Alameda, CA). NuSieve 3:1 agarose was obtained from FMC Bioproducts (Rockland, ME). Other chemicals and reagents were of the highest grade readily available.
Plasmids.
pFR-luciferase (UAS luciferase, Gal4 response element) was purchased from BD Biosciences Clontech (Palo Alto, CA). pRL/TK Renilla (phRL-TK), pRL/CMV Renilla (phRL/CMV), pSV/-Gal, and pGEM-T Easy were purchased from Promega (Madison, WI). The construction of pBK-CMV/rPPAR-FLAG and the mammalian two-hybrid pM/PPAR constructs was described previously (Sumanasekera et al., 2003a). The PPAR response element reporter, pACO(-581/-471)G.Luc was supplied by Dr. Jonathon Tugwood (Zeneca Pharmaceuticals, Central Toxicology Laboratories, Maccelsfield, UK) and has been previously described (Tugwood et al., 1998). Maltose binding protein- PPAR (MBP/PPAR) and MBP/rpL11 were generated by subcloning full-length rat cDNA PPAR (a gift from Dr. Frank Gonzalez, National Cancer Institute, Bethesda, MD) and rat ribosomal protein rpL11 (a gift from Dr. Ira Wool, University of Chicago, IL) into pMAL-c2X (New England Biolabs, Beverly, MA). pcDNA3.1/V5-His-rPPAR was generated by cloning into pcDNA3.1/V5-His-A (Invitrogen). VP16/rpL11 was generated by cloning rat rpL11 into pVP16 (Clontech).
cDNA library construction and screening.
FaO rat hepatoma cells (a generous gift from M.C. Weiss, Pasteur Institute, Paris, France), were grown in DMEM (Sigma) supplemented with 5% FBS, 100 units/ml of penicillin and 10 μg/ml of streptomycin, as previously described (Sterchele et al., 1996). Total RNA was isolated from FaO cells at 75% confluency using Tri Reagent (Molecular Research Center, Cincinnati, OH) and poly(A) mRNA extracted using oligo(dT)-magnetic beads (Promega). A directional cDNA library was constructed using the SuperscriptTM Lambda System (Gibco BRL) using the manufacturer's protocols. The yield and purity of the library construction was monitored at each step by incorporating -32P-CTP into the cDNA and removing small aliquots. The average transcript size of the library is 3000 nucleotides (range 1–9 kb).
PPAR-associated proteins (PAPs) were identified by screening the cDNA expression library using an overlay assay described previously (Chapline et al., 1993). Approximately 5 x 104 plaque forming units (pfus) were plated on each of 20 agar (150 mm) plates. The plates were incubated for 3 h at 37°C, after which isopropyl--D-thiogalactopyranoside (IPTG)-soaked nitrocellulose filters were overlaid, and incubation continued for an additional 4 h at 37°C. The orientation of the filters was marked, and they were removed from the plates and rinsed briefly with Tris-buffered saline (TBS), pH 7.4, containing 0.1% Tween 20 (TBS-tween). The membranes were blocked with 5% BSA (diluted in TBS-tween) for 1 h, followed by three 10-min washes with TBS-tween. The membranes were then probed with a bacterially-expressed PPAR-maltose binding protein (PPAR-MBP, see details below) for 2 h (diluted in TBS-tween plus 0.1% BSA). After three washes, the blots were incubated with an antibody to MBP (New England BioLabs, Beverly, MA) at the recommended dilutions. Detection of PPAR-MBP was performed by incubating the filters with an anti-rabbit IgG secondary antibody conjugated to alkaline phosphatase (Promega) and visualized with NBT (nitro blue tetrazolium)/BCIP (5-bromo-4-chloro-3-indolyl-phosphate). Positive plaques were eluted from agar plugs and were rescreened as described above. Positive clones from secondary and tertiary screens were excised out of Ziplox, and the inserts were sequenced in both directions by the Penn State Nucleic Acid Facility.
Transfection and reporter assays.
To confirm the interaction between PPAR and the isolated clones, Clontech's MatchMaker yeast (Y2H) and mammalian two-hybrid (M2H) systems were employed (Palo Alto, CA). Yeast SFY526 cells were transformed as per supplied protocols, and positive interactions were assessed by a -galactosidase (-gal) filter assay. Lipofectamine (Invitrogen) was used to cotransfect COS-1 cells (cultured in -MEM supplemented with 8% FBS and 100 units each of penicillin and streptomycin) with the M2H vectors and pFR/luciferase. Included in all transfections were pSV/-gal, pRL/CMV, or pRL/TK (Promega) to control for transfection efficiency. Luciferase enzymatic activity was measured with the single or dual luciferase enzyme assay system (Promega). -gal activity was measured using the -galactosidase enzyme assay system (Promega). For transient transfection experiments examining PPRE activity, pPPAR and pACO(-581/-471)G.Luc were cotransfected into COS-1 cells with or without either pCMX/RXR or pcD/rat rpL11 using Lipofectamine. Following treatment, cells were harvested, and renilla and firefly luciferase activities were examined using the Dual Luciferase Assay (Promega).
MBP pull-down assay.
PPAR or rpL11 were expressed as a maltose-binding protein (MBP) fusion protein and extracted from DH5 bacteria using amylose resin (New England Biolabs) as previously described (Riggs, 2000). Proteins were stored in "everything buffer" (20 mM Tris (pH 7.4), 200 mM NaCl, 1 mM EDTA, 10 mM -mercaptoethanol, and 1 mM sodium azide). In vitro translated PPAR or rpL11 were prepared using the TnT coupled reticulocyte lysate system (Promega) in the presence of 35S-labeled methionine (Amersham). Proteins were incubated in TBS containing 0.5% BSA and amylose resin on a rocker at 4C for 1 h. Afterward, the resin was washed five times with either 4 mM EDTA, 15 mM NaMOPS, 20 mM MOPs, 5 mM sodium azide, and 10% glycerol (precipitations A and B) or 150 mM NaCl, 1% CHAPs, and 5mg/ml BSA in everything buffer (precipitation C). The final pellet was resuspended in 2x SDS-tris/glycine sample buffer, boiled, and subjected to SDS/PAGE. Autoradiography was performed on the dried gels.
Gel shift assays.
Both strands of binding PPAR response element (CN) (5' CAAAACTAGGTCAAAGGTCA 3') and a non-binding PPAR response element (MeD) (5'-GTGTTAGAGGGCACAGGTCC) were synthesized and annealed as previously described (Juge-Aubry et al., 1997). For the first gel shift, in vitro translations of PPAR (2 μl), RXR (2 μl), and combined rpL11 and nonprogrammed lysate (6 μl) were added to each reaction. For the second assay, 0.25 μl of RXR, 2.5 μg of either MBP-empty or MBP-PPAR, and 0–3 μg of MBP-L11 were added to each reaction (0, 0.5, 1, 1.5, 2, and 3 μg in lanes 1–6 and 7–12, respectively). All reactions were performed in 4% glycerol, 1 mM MgCl2, 0.5 mM EDTA, 0.5 mM DTT, 50 mM NaCl, 10 mM Tris/HCl (pH 7.5), 0.5% CHAPS, and either 125 ng (assay one) or 64 ng (assay two) poly-deoxy-inosinic-deoxycytidylic acid (poly dI:dC) in a volume of 10 μl (assay one) or 15 μl (assay two). All reactions were incubated for 15 min before and after adding 1 μl 32P-labeled response element (>50,000 cpm). Where indicated, 1 μl (25 μM) of unlabeled CN or MeD were added. The incubations were resolved on 6% DNA Retardation Gels (Invitrogen) using 0.5x TBE (5.4g Tris base, 2.75g boric acid, and 1mM EDTA).
Manipulation of rpL11 activity.
Hepa-1 cells were maintained in Dulbecco's modified Eagle's medium containing 8% FBS in a 37°C incubator with 5% CO2. For PPRE-dependent reporter assays, cells were grown to 60% confluence and transfected with HD-Luciferase (kind gift from D. J. Waxman, Boston University) which contains a functional PPRE from the peroxisomal enoyl-CoA hydratase/3-hydroxyacyl-CoA dehydrogenase gene. Subsequently, Hepa-1 cells were treated with DMSO (0.1%) or Wy-14,643 (50 μM) concurrently with two means to modulate rpL11 activity, actinomycin D (AD) or serum deprivation (SD) (Bhat et al., 2004; Perry and Kelley, 1970). AD was dissolved in DMSO and added to cells at a final concentration of 5 nM. Serum deprivation was achieved by washing the cells twice with PBS and adding DMEM media with 0.1% serum. For endogenous gene expression studies, 60% confluent cells were treated with DMSO or Wy14,643 and subjected to AD or SD treatment as above. In all cases, AD and SD were performed concomitantly with chemical treatments for 24 h. A double-stranded RNA inhibitor for rpL11 was designed using the sequence from GenBankTM NM_025919. The sequence chosen was as follows: 5'-AAGCATTGGGATCTACGGCCT-3'. The sequence was synthesized and cloned into pSuper.neo plasmid (Oligoengine, Seattle, WA). Transfection of the RNAi plasmid into Hepa-1 cells was performed using Lipofectamine (Invitrogen) transfection reagent according to the manufacturer's protocol for 12 h, followed by serum starvation or Actinomycin D and Wy-14,643 treatment as described above. A rabbit polyclonal antibody against a C-terminal L11 peptide was a kind gift of K. H. Vousden (Beatson Institute for Cancer Research, UK).
Real-time PCR.
Total RNA was isolated by TriReagent (Sigma) according the manufacturer's instructions. The total RNA was reverse transcribed using the ABI High Capacity cDNA archive kit (Applied Biosystems, Foster City, CA). Standard curves were made using serial dilutions from pooled cDNA samples. Real Time PCR was performed using the SYBR Greeen PCR Master Mix (Applied Biosystems) according to the manufacturer's protocol and amplified on the ABI Prism 7000 Sequence Detection System.
Statistical analysis.
Where indicated, MiniTab (State College, PA) was used to evaluate data for statistical significance using One-way ANOVA and Student's t-test with significance at p < 0.05.
RESULTS
Identification of PPAR-Associated Proteins
To identify cDNAs which encode PPAR-associated proteins (PAPs) we screened a FaO Ziplox expression library, using as a probe PPAR-MBP fusion protein expressed and purified from bacterial extracts. Visualization of positive plaques was accomplished immunologically by use of a monoclonal antibody to the MBP domain. On three separate experiments, 106 phage plaques were screened. From this primary screen, 15 positives were eluted from the agar, plated, and rescreened. Of those initial positives, 7 were confirmed with subsequent secondary and tertiary screens and were excised from the phage and sequenced (Table 1). DNA sequencing revealed high homology of several clones with published sequences. PAP2 and PAP4, determined to be homologous clones isolated from primary screens during two separate experiments, had inserts of 539 and 575 base pairs (bp), respectively. PAP2/4 is identical to the rat ribosomal protein rpL11 (Chan et al., 1992), which is part of the large 60S ribosomal subunit (Tsurugi et al., 1976). While PAP2/4 did not contain the complete open reading frame of rpL11, they did contain a continual stretch that corresponds to nucleotide number 22 until 552 (of the total length of 800 bp (Chan et al., 1992). PAP1 and PAP3 demonstrated significant homology with the rat DCT1 (a member of the Nramp family of genes (Gunshin et al., 1997) and CITED2 (homologous to human coactivator CITED2/p35srj (Bhattacharya et al., 1999), respectively. PAP5 matched well with ST7/RAY1/HELG protein (accession AJ277291), while another clone (PAP6) contains less than 200 bp and was not examined further. PAP7 contained the largest insert, and matched with the rat cysteine proteinase, cathepsin L (Ishidoh et al., 1987).
Confirmation of PPAR:PAP Interaction
In order to confirm that identified PAPs interact with PPAR in vivo, a yeast-based system was utilized. The PAPs were subcloned into the yeast expression vector, pGAD424, and the rat PPAR was inserted into pGBT9. When used to transform yeast, these vectors express a chimera of the activation domain and the DNA binding domain of the GAL4 transcription factor, respectively. Interactions were assessed by a -gal filter assay in which blue plaques would indicate expression of lacZ, and therefore an interaction between the two chimeras. -gal activity was qualitatively assessed and summarized in Table 1. When full-length PPAR is expressed by itself, there is substantial amount of reporter activity (see Table 1 legend). This is not surprising, since the amino-terminus of the receptor contains transcriptional activation functions (Tsai and O'Malley, 1994). To circumvent this background activity when using this system, other researchers have deleted the transactivation domain of other nuclear receptors (Burris et al., 1995). However, we screened the library with intact transcriptional activation domains and, therefore, performed the yeast two-hybrid assay with full-length PPAR. As indicated in Table 1, when yeast were cotransfected with various PAPs and PPAR, there was a decrease in reporter activity. As a positive control, human RXR, subcloned into pGAD424, was used with pGBT9.rPPAR to cotransfect yeast cells, and a decrease in reporter activity was also observed. Based on the reduction in -gal activity in the presence of PPAR and the PAPs (as compared to PPAR alone), all PAPs were confirmed to interact with PPAR. CITED2 (PAP3) is a bona fide PPAR coactivator and has been examined elsewhere (Tien et al., 2004). Due to the fact that other ribosomal proteins are known to functionally interact with nuclear receptors, and that rpL11 affects the cell cycle, the present studies focused on the PAP2/4 (rpL11) and PPAR association.
Full-Length rpL11 Inhibits PPAR Transactivation
The effect of PAP2/4 as well as full-length rpL11 on PPAR's ability to influence the expression of a reporter gene under the control of a natural PPRE was examined (Fig. 1). Vectors containing PPAR and rpL11 were cotransfected into COS-1 cells along with pACO(-581/-471)G.Luc. As indicated in Figure 1A, cotransfecting cells with increasing amounts of the rpL11 plasmid inhibited the transcriptional activity of PPAR in a dose-dependent manner. These results were dependent on PPAR, since PAP2/4 by itself had no effect on luciferase activity (data not shown). Similar results were obtained with the partial clones of rpL11 (PAP2/4, Fig. 1B) as well as the full-length rpL11 (Figure 1C) on pM/PPAR activity using the GAL4 reporter, although the amount of reduction in PPRE-driven luciferase activity was more pronounced.
L11 Interacts with the D-Domain of PPAR
We utilized the mammalian two-hybrid assay to map the interaction between PPAR and rpL11. Various domains of PPAR were subcloned in the pM vector and examined for their ability to interact with VP16/L11 (Fig. 2A). All data is compared to the amount of reporter activity in the presence of VP16/Empty under the same treatment conditions. The D-domain (amino acids 168–273) had the highest amount of positive interaction with rpL11. Full-length rPPAR in the presence of Wy14,643 and the E/F domain in the unliganded state showed a decrease in reporter activity in the presence of VP16/rpL11. The interaction of ribosomal protein L7a enhances the effects of partial agonists but not full agonists of the estrogen receptor (Jackson et al., 1997). To test whether this holds true for the PPAR/L11 association, several known PPAR activators were examined in the M2H system (Fig. 2B). Cotransfecting with rpL11 decreased the activation of PPAR by Wy14,643. Less efficacious ligands such as bezafibrate, ETYA, and clofibric acid (not shown) activated PPAR in the presence or absence of cotransfected rpL11. The specific inhibitor of PPAR, MK886 (Kehrer et al., 2001), was also unaffected by cotransfection with rpL11 (not shown).
PPAR Subtype Preference for rpL11 Interaction
The ability of rpL11 to affect the activity of PPAR, , and was examined using M2H assays (Fig. 3). PPAR was activated with ligands specific for each subtype in the presence or absence of VP16/rpL11. Wy14,643 significantly increased PPAR activity, while carbacyclin and PGJ2 significantly activated PPAR and , respectively. The ligand-dependent activation of PPAR was decreased by cotransfection with rpL11-containing plasmid. Neither basal nor ligand-induced PPAR or PPAR activity was affected by rpL11 transfection. Similar results were obtained when the partial sequence of rpL11 was used (PAP2/4) instead of the full-length construct (not shown).
PPAR Interacts with rpL11 in Vitro
PPAR was expressed in bacteria as a maltose binding protein (MBP) fusion protein and partially purified as described under Materials and Methods. PPAR-MBP was mixed with in vitro translated full-length rpL11, and the resultant complex was affinity purified using amylose resin. The protein complex was separated on SDS/PAGE, transferred to nitrocellulose, and rpL11 was detected by autoradiography. Full-length ribosomal rpL11 copurified with PPAR-MBP (Fig. 4A lane 2) but not MBP alone (lane 3). Lane 1 indicates the amount of in vitro translated rpL11 used in the pull-down experiments. The reverse incubation was also performed where rpL11 was expressed as an MBP fusion and PPAR was in vitro translated (Fig. 4B). Increasing amounts of PPAR were copurified in the presence of increasing rpL11-MBP. Finally, various domains of PPAR fused to MBP were examined for their ability to associate with in vitro translated rpL11 (Fig. 4C). As expected from the M2H assays, the full-length (FL) and D-domains of PPAR contained the predominant rpL11 binding capability.
rpL11 Inhibits PPAR Binding to the PPRE
The effect of rpL11 on DNA binding of the PPAR/RXR heterodimer was examined using in vitro translated or bacterially expressed proteins. In Figure 5A, PPAR, RXR, and rpL11 were in vitro translated and mixed with each other prior to the addition of 32P-labeled PPRE. PPAR/RXR binding to the 32P-labeled PPRE could be competed with cold CNPPRE but not with the MeD response element (lanes 2 versus 3 and 4). In lanes 5 through 9, increasing amounts of rpL11 in vitro translation was added to a constant amount of PPAR. To this complex, RXR was added, followed by 32P-labeled PPRE. With increasing amounts of rpL11, there was a decrease in the binding to the response element. Interestingly, if in vitro translated PPAR and RXR are mixed prior to the addition of rpL11, there was no decrease in DNA binding, indicating that rpL11 could not affect the preformed heterodimer's interaction with DNA. In the experiments depicted in Figure 5B, MBP (lanes 1–6) or MBP-PPAR (lanes 7–12) was incubated with increasing concentrations of MBP-L11 (lanes 2–6, 8–12). All reactions contained an equal amount of in vitro translated RXR and equal amounts of radiolabeled PPRE probe as described in Materials and Methods. The PPAR/RXR specific band is shown as band (B) and was decreased with increasing amounts of rpL11. At higher rpL11 concentrations, two shifted bands appear (A and C), which are not dependent upon PPAR for DNA binding and are possibly direct interactions of rpL11 with DNA. Even at lower rpL11 concentrations, this ribosomal protein can inhibit PPAR from binding to DNA, whether rpL11 is in vitro translated or bacterially expressed. This effect was not due to a change in the localization of PPAR, which was unaffected by rpL11 (data not shown).
PPAR Activity under Conditions of rpL11 Manipulation
The activity of endogenous PPAR or both endogenous and transfected PPAR in Hepa-1 cells was examined using a PPRE-reporter as well as real-time PCR. As shown Figure 6 (panels A and B), in the absence of exogenously supplied PPAR there is a modest induction of HD-luciferase activity (approximately 50%) by Wy14,643. The increase due to Wy14,643 addition is approximately two-fold in the case where PPAR cDNA is supplied. Actinomycin D and serum starvation have been shown to destabilize the ribosome, allowing for potential interactions between ribosomal proteins and other cellular components (Bhat et al., 2004; Perry and Kelley, 1970). Actinomycin D treatment in cells which did not receive exogenous PPAR increased PPRE-dependent activity roughly three-fold; this effect was absent when PPAR was cotransfected. In the presence of either AD or SD, the Wy14,643-mediated induction of HD-luciferase is no longer observed, but only when endogenous PPAR is examined. When exogenous PPAR is supplied, the Wy14,643-mediated response is still observed in the presence of AD or SD. Transfection with the rpL11 RNAi plasmid resulted in an increase in HD-luciferase activity regardless of treatment (Fig. 6, panels C and D). These experiments were performed in the absence of exogenously supplied PPAR, and Wy14,643 had marginal effects on reporter activity in this case.
Since AD and SD treatment affected PPRE-driven reporters in the absence of overexpressed PPAR, these conditions were used to examine effects on known target genes of this nuclear receptor. Similar results to the reporter assay were obtained when two PPAR responsive genes were examined, Angptl4 and ACO (Fig. 7). Treatment with Wy14,643 resulted in a slight increase in both of these mRNAs in the absence of AD (Panel A) or SD (Panel B) with either no observed increase or a reversal of effect in the presence of these treatments. In general, transfection of an RNAi plasmid targeted to rpL11 increased PPAR target gene expression relative to that observed in cells which received control plasmid (Panels C–F). The amount of rpL11 inhibition by the RNAi plasmid in transient transfection of these cultures was approximately 30%.
DISCUSSION
The mechanism by which NRs are able to alter the expression of target genes has been extensively studied. The simplest model was one in which, once activated by ligand, NRs bind as dimers (homo- or hetero-) to specific sequences in the 5' regulatory region of target genes (Tsai and O'Malley, 1994). Recent work, however, suggests this scheme is overly simplified. Transcriptional activation involves interplay between many cellular processes. NRs are regulated by phosphorylation (Weigel, 1996) and heat shock proteins (Pratt and Toft, 1997), as well as two important groups of transcriptional regulators known as coactivators and corepressors (Torchia et al., 1998).
Due to the increasing body of evidence that protein–protein interactions are key regulatory steps in the NR signal transduction pathway, we intended to identify novel PPAR-associated proteins. In this report we describe the identification of the ribosomal protein L11 as such a protein. Originally detected by screening a cDNA expression library constructed from FaO mRNA, this interaction was confirmed in yeast and mammalian two-hybrid systems (Table 1 and Fig. 1, respectively). Additionally, we were able to demonstrate biological relevance when it was observed that cotransfection of PPAR with the full-length rpL11 resulted in a decrease in the transcriptional activity of the receptor and a decrease PPAR/RXR heterodimerization. Furthermore, rpL11 directly interacted with PPAR in MBP pull-down assays.
Ribosomal protein rpL11 is part of the large 60S ribosomal subunit (Tsurugi et al., 1976), and homologues have been identified in prokaryotes as well as eukaryotes. This is not the first time that a ribosomal protein has been observed to interact with a transcription factor. Indeed, two ribosomal proteins, L10 and L18a, have been identified as interaction partners for c-Jun (Chan et al., 1996; Gramatikoff et al., 1995). Furthermore, Burris and colleagues (1995) reported that the thyroid hormone (TR) and retinoic acid receptors (RAR) associated with the human ribosomal protein L7a. These interactions resulted in an inhibition of transcriptional activity of the receptors. An interesting observation made by Jackson et al. (1997) was that the association of ER, GR, or PR with L7 increases in the presence of partial, but not full, agonists. These authors also mapped the site of protein–protein interaction to the D-domain of the receptors. In the present studies we showed clear preference for full PPAR agonists such as Wy14,643 relative to partial agonists such as bezafibrate or ETYA in their ability to stimulate PPAR/L11 interactions. Although the ligand preference for the PPAR/L11 association is dissimilar to that of ER/L7a, the NR domain utilized is the same, the D- or hinge-domain. This region is variable among the PPAR subtypes, which may help explain why rpL11 affected PPAR activity but not PPAR or .
The observations that L7 interacts with RAR, PR, ER, and GR, L10 with c-jun as well as those presented herein, point to the possibility of ribosomal proteins having extraribosomal functions (Wool, 1996). In fact, it is still unclear as to whether ribosomal proteins evolved specifically for the ribosome, or whether they were co-opted from existing proteins with defined functions. As of now there is only circumstantial evidence to suggest that these proteins existed initially for other purposes. First, many ribosomal proteins contain structural motifs, such as zinc fingers, leucine zippers, and helix-turn-helix motifs, which suggest that they have the capacity to bind to DNA (Rice and Steitz, 1989; Wool et al., 1995). Bacterial expressed rpL11 caused a DNA gel shift when present at high concentrations (Fig. 5B); whether this protein binds to DNA in vivo has not yet been determined. Second, ribosomal proteins interact with transcription factors such as c-Jun, TR, and RAR, as well as our own observation that rpL11 associates with and inhibits PPAR's ability to regulate PPRE-driven reporters (Figs. 6 and 7). Last, TR and PPAR are NRs, and this superfamily is believed to have evolved more that 500 million years ago in primitive organisms (Laudet, 1997; Sumida, 1995). Since many ribosomal proteins have prokaryotic homologues (Wool et al., 1995), one could imagine primitive organisms evolving to express ribosomal proteins as the initial negative regulators of ancestral transcription factors.
The biological importance of the interaction between PPAR and rpL11, as well as other NR-ribosomal protein interactions, remains unclear. It appears that rpL11 inhibits PPAR from forming a heterodimer with RXR, thereby preventing an active complex from binding to PPREs. Similarly, Burris et al. (1995) demonstrated that ribosomal protein L7a prevented the TR/RXR heterodimer from binding to TREs, but had no effect on RXR's ability to bind to its response element. On the other hand, these researchers did not indicate whether L7a was able to interact with TR or RXR in vitro. It is possible that ribosomal proteins are able to prevent the formation of certain NR/RXR heterodimers by competing for RXR. The fact that we were able to inhibit the activity of pM/PPAR, a construct that contains a heterologous DNA binding motif and does not require RXR interaction for activity, suggests that this inhibition of PPAR activity may have multiple mechanisms. In addition to inhibiting RXR heterodimerization, rpL11 may affect coactivator recruitment, for example. Sequestering of PPAR in the cytoplasm does not appear to occur, as we were unable to detect differences in the cytosol/nucleus ratio when rpL11 is overexpressed (data not shown); alteration of suborganelle distribution, such as nucleolar colocalization of rpL11 and PPAR has not yet been examined.
The potential significance of the PPAR/rpL11 interaction has increased as a result of observations of an rpL11/HDM2/p53 complex (Lohrum et al., 2003; Zhang et al., 2003). In fact, overexpressing the ribosomal protein is able to induce cell cycle arrest (Lohrum et al., 2003). The mechanism of the rpL11 control of the cell cycle is beginning to be understood. Ribosomal protein L11 can associate with HDM2, the ubiquitin ligase of p53, and inhibit the degradation of p53 (Lohrum et al., 2003). The HDM2/L11 association is enhanced by disruption of the ribosome by actinomycin-D (Lohrum et al., 2003; Zhang et al., 2003). It has been postulated that a signal stimulating cell growth would promote rpL11 assembly into the ribosome, allowing HDM2 to inhibit p53 activity. A signal for inhibition of cell growth, such as perturbation in ribosome biogenesis, would enhance HDM2 activity and, hence, increase p53 and growth arrest (Zhang et al., 2003). The modulation of p53 activity by a ribosomal protein is not exclusive to rpL11 (Bhat et al., 2004; Lohrum et al., 2003; Zhang et al., 2003) as rpL5 (Dai and Lu, 2004) and rpL23 (Dai and Lu, 2004; Dai et al., 2004; Jin et al., 2004) affect the turnover of p53 via their ability to bind to and suppress the E3 ligase function of HDM2. It is intriguing to speculate that a PPAR/L11 association may affect the formation of the rpL11/HDM2/p53 complex. We have recently reported that PP-induced PPAR activation was able to increase entry of hepatocytes into the cell cycle (Tien et al., 2003). Thus, we would hypothesize that mitogenic agents such as Wy14,643 would decrease p53 activity. Although speculative at this point, it is possible that unliganded PPAR may enter into the p53 complex through its association with rpL11.
In summary, the ribosomal protein rpL11 interacts with the D-domain of PPAR and inhibits PPRE-driven reporter activity. Manipulation of rpL11 activity by a variety of means affects PPAR activity, whether addressed using reporter assays or endogenous gene expression. Ribsomal protein L11 joins a growing list of ribosomal proteins that can affect gene transcription. Recent interest in rpL11 has resulted from observations of an apparent rpL11/HDM2/p53 complex. Thus, the interaction of rpL11 with PPAR, a nuclear receptor implicated in cell cycle control and carcinogenesis, has increasing importance.
NOTES
1 Current address: Rutgers University, Pharmacology & Toxicology, 170 Frelinghuysen Rd., Rm. 441, EOHSI Piscataway, NJ 08854.
2 Current address: Pfizer Global Research & Development, World Wide Safety Sciences, 800 N Linderberg Boulevard, Creve Coeur, MO 63167.
ACKNOWLEDGMENTS
The authors thank the technical assistance of Kristine Walker, Tecla Brabazon, and Nicole Agostino. The authors also thank the editorial assistance of Jerry Thompson. This work was supported by NIH ES007799 (JVH).
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