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The mRNA endonuclease PMR1 initiates mRNA decay by forming a selective complex with its translating substrate mRNA. Previous work showed that the ability of PMR1 to target to polysomes and activate decay depends on the phosphorylation of a tyrosine residue at position 650. The current study shows that c-Src is responsible for activating this mRNA decay pathway. c-Src was recovered with immunoprecipitated PMR1 and it phosphorylates PMR1 in vitro and in vivo. The interaction with c-Src involves 2 domains of PMR1; Y650 and a series of proline-rich SH3 peptides in the N-terminus. In cells with little c-Src PMR1 targeting to polysomes is induced by constitutively active c-Src but not by inactive forms of the kinase. Similarly only active c-Src induces PMR1-mediated mRNA decay. Finally we show that EGF rapidly induces c-Src phosphorylation of PMR1, providing a direct link between tyrosine kinase-mediated signal transduction and mRNA decay.
mRNA decay is a major mechanism controlling gene expression, and over the past several years significant advances have been made in elucidating mechanisms of the decay process. For most mRNAs decay begins with the shortening of the poly(A) tail, after which there is a change in the nature of the mRNP, with proteins that facilitate mRNA decay (eg. Lsm1-7, Pat1, Dhh1/p54/Rck, Dcp1/2) replacing those associated with translation (eg. poly(A)-binding protein, eIF4G) (Tharun and Parker, 2001). For most yeast mRNAs and possibly most mammalian mRNAs the next step involves loss of the cap and 5′-3′ exonuclease-mediated decay of the mRNA body by Xrn1 (Stoecklin et al., 2005). The proteins that facilitate and catalyze decapping and 5′-3′ decay concentrate in discrete cytoplasmic foci termed ‘processing bodies’ (P bodies, also called GW bodies (Jakymiw et al., 2005)), and while there is evidence that decay can occur in these large RNP complexes it is not clear whether this process is restricted to P bodies or also occurs in submicroscopic complexes. In addition to this mechanism there are reports of unstable mRNAs undergoing 3′-5′ decay catalyzed by the cytoplasmic exosome (Chen et al., 2001; Mukherjee et al., 2002). The binding of proteins like HuR to AU-rich elements can stabilize mRNA whereas tristetraprolin, butyrate response factor 1 and others can promote 5′ decay (Fenger-Gron et al., 2005). Activation of different kinases can modulate the binding of stabilizing and destabilizing proteins, and in this manner regulate mRNA decay (Stoecklin et al., 2004; Briata et al., 2005; Chou et al., 2006).
In contrast to these exonuclease-based pathways a number of mRNAs are targeted by protein endonucleases that initiate decay by cleaving within the body of the mRNA while it is actively engaged by translating ribosomes. While endonucleases have been implicated in the decay of a number of mRNAs (Pastori et al., 1991; Binder et al., 1994; Wang and Kiledjian, 2000), only 3 mRNA endonucleases have been linked to specific decay pathways; PMR1 (Chernokalskaya et al., 1998), G3BP (Gallouzi et al., 1998) and IRE-1 (Hollien and Weissman, 2006). PMR1 was originally identified as a ribonuclease activity whose appearance on Xenopus liver polysomes after exposure to estrogen coincides with the post-transcriptional disappearance of serum protein mRNAs. Subsequent work showed that estrogen activates the decay of these mRNAs by inducing a 21-fold increase in the unit activity of the polysome-bound endonuclease (Cunningham et al., 2001). PMR1 is distinct in that its sequence and structure bear no relationship to the known ribonucleases. Instead it is non-heme containing member of the peroxidase gene family which, like most peroxidase gene products, is made as a larger precursor (PMR80) and processed to a smaller active form (PMR60) (Chernokalskaya et al., 1998).
Our early work used whole animals and primary frog hepatocyte cultures to characterize much of the biology of PMR1. However the technical difficulty of preparing primary frog hepatocyte cultures and their low transfection efficiency led us to explore selected mammalian cell lines as surrogates. Subsequent work showed that Xenopus PMR60 functions in a manner analogous to its native context when it is expressed in stably or transiently transfected cells regardless of whether these cells express (Bremer et al., 2003) or do not express (Yang and Schoenberg, 2004) the mammalian ortholog of this protein. Using deletion mutants of a catalytically-inactive form of Xenopus PMR1 (PMR60°) we identified specific polysome targeting domains, and experiments with catalytically-active PMR60 showed that both polysome targeting and translation of its substrate mRNA are required for endonuclease-mediated mRNA decay (Yang and Schoenberg, 2004). Interestingly a similar mechanism was recently described for IRE-1-mediated endonuclease decay of a specific set endoplasmic reticulum-associated mRNAs in response to the accumulation of unfolded proteins (Hollien and Weissman, 2006). Y650 in the C-terminus of PMR60 plays a critical role in targeting this protein to polysomes. Phosphorylation at this tyrosine generates a consensus SH2 ligand that is required for PMR60 targeting to polysomes (Yang et al., 2004). Changing this amino acid to phenylalanine or treating cells with the general tyrosine kinase inhibitor AG18 inactivated polysome targeting and endonuclease-mediated decay. Here we identify c-Src as the PMR60 tyrosine kinase and show that its activation by EGF binding to EGFR results in rapid tyrosine phosphorylation of the mRNA endonuclease.
PMR60 participates in two major complexes; a ~680 kDa mRNP (complex I), which is the polysome-bound form containing the endonuclease and its substrate mRNA (Yang and Schoenberg, 2004), and a ~140 kDa complex (complex II) that is the precursor to complex I (Yang et al., 2004). Tyrosine phosphorylation occurs in complex II, and a doublet product was seen when PMR60-containing complexes recovered from glycerol gradient fractions of this complex were probed with the phosphotyrosine monoclonal antibody PY20 whereas only a single band was observed when the same fractions were probed for the N-terminal myc tag on PMR60° (Yang et al., 2004). Since tyrosine kinases commonly undergo autophosphorylation we asked whether the smaller band identified with PY20 might be a tyrosine kinase bound to PMR60°. In the experiment in Fig. 1A immunoprecipitated complexes from cells expressing myc-tagged GFP or PMR60° were incubated in vitro with γ-[32P]ATP and radiolabeled proteins were separated by SDS-PAGE. Two bands were specifically labeled by a kinase recovered with PMR60° (lane 5). The upper band corresponds to PMR60° (filled circle) and the lower band (open circle) corresponds in size to a control of c-Src (lane 7) that was autophosphorylated when incubated under the same conditions.
To confirm that PMR60° is a substrate for c-Src we expressed His-PMR60° in bacteria (Fig. 1B, left panel) and incubated increasing amounts of recombinant protein with recombinant c-Src and γ-[32P]ATP. Autophosphorylation of c-Src was again evident (Fig. 1B, right panel) as was the increased labeling of His-PMR60° with increasing amounts of input protein. The ability of c-Src to specifically label Y650 was tested using a peptide containing this site. In Fig. 1C recombinant c-Src was added to the indicated peptide that was synthesized without (peptide) or with a phosphate (PY-peptide) on the tyrosine corresponding to Y650. Autophosphorylation of c-Src was again seen and only the unphosphorylated form of the peptide was labeled, thus confirming that PMR60° is a substrate for c-Src.
Reciprocal immunoprecipitation was used to examine the in vivo interaction of c-Src with PMR60° (Fig. 1D). U2OS osteosarcoma cells were transfected with plasmids expressing myc-tagged PMR60° or GFP, and protein complexes recovered by immunoprecipitation with monoclonal antibody to the myc tag (left panels) or a polyclonal antibody to c-Src (right panels) were analyzed by Western blot. c-Src was recovered with PMR60° but not GFP (lanes 3 and 4). Conversely, PMR60° was recovered with anti-Src but GFP was not (lanes 7 and 8), thus confirming c-Src as the kinase activity recovered in Fig. 1A. The functional impact of this interaction is shown in Fig. 1E, where treating U2OS cells for 60 min with the selective c-Src inhibitor PP2 blocked tyrosine phosphorylation of PMR60° whereas its inactive congener (PP3) had no effect.
c-Src is elevated in many cancers (Irby and Yeatman, 2000) and cancer-derived cell lines. To examine further the role of activated c-Src in catalyzing the phosphorylation of PMR60° in vivo we used a line of Cos-1 cells that has 5-times less c-Src than U2OS cells (Fig. 2A). These were transfected with plasmids expressing PMR60° plus empty vector, a dominant negative form of c-Src (DN-Src) or a constitutively active form of c-Src (CA-Src), and the immunoprecipitated protein was examined as in Fig. 1 to determine the impact of each form of c-Src on tyrosine phosphorylation (Fig. 2B). As one might anticipate from the reduced amount of c-Src there was no evidence for tyrosine phosphorylation in vector transfected cells. Similarly tyrosine phosphorylation was not observed in cells expressing DN-Src, but PMR60° was phosphorylated in cells expressing CA-Src. In the right panel the recovery of each of these forms of c-Src with PMR60° was determined by Western blot. While somewhat more DN-Src was present in the input sample in 3 independent experiments significantly more of this protein was recovered with PMR60° than CA-Src. These data suggest that the functional interaction of c-Src with PMR60° is transient and the complex dissociates after PMR60° is phosphorylated.
Among the numerous pathways in which c-Src participates the best known is its activation by EGF binding to the EGF receptor (EGFR). c-Src interacts directly with EGFR, and EGF binding activates the reciprocal phosphorylation of both proteins to initiate downstream events. To determine if this upstream signaling process induces c-Src phosphorylation of PMR60° Cos-1 cells were first cultured in serum free medium to reverse the down-regulation of EGFR caused by prolonged growth in serum-containing medium (Wong et al., 2002). Changes in tyrosine phosphorylation of immunoprecipitated PMR60° were then followed as a function of time after EGF addition (Fig. 2C). Prior to adding EGF there is little evidence for tyrosine phosphorylation of PMR60° (lane 1). Tyrosine phosphorylation is induced within 10 min and is maintained over the 60 min time course examined here, thus indicating that EGF binding to its receptor activates the phosphorylation of PMR60°. To confirm that c-Src is the responsible kinase this was repeated with cells that were treated for 30 min with PP2 or PP3 prior to adding EGF (Fig. 2D). PP3 had no impact on tyrosine phosphorylation of PMR60° but this was inhibited by PP2, thus confirming that EGF activates c-Src to phosphorylate PMR60°.
To identify features of PMR60° that define its interaction with c-Src we employed a battery of deletion constructs (diagrammed in Fig. 3A) that were used previously to map functional domains of PMR60° (Yang and Schoenberg, 2004; Yang et al., 2006). In the experiment in Fig. 3B Cos-1 cells were co-transfected with GFP or each of the indicated PMR60° deletion constructs and wild-type c-Src (WT-Src), and Western blots of the recovered proteins were probed with antibodies to the myc tag on PMR60° and GFP, and c-Src. WT-Src was used here because more of this form of the enzyme is recovered with PMR60° than CA-Src (eg. Fig. 2B and data not shown). Removal of the N-terminal 50 amino acids had no impact on the recovery of c-Src (lane 3), binding was reduced after deletion of the N-terminal 100 amino acids (lane 4), and lost after deletion of the N-terminal 150 amino acids (lane 5). The C-terminal 50 amino acids lie just downstream of Y650 and deleting this portion of the protein had no impact on the recovery of c-Src (lane 6). However, recovery of c-Src was lost when the C-terminal 100 amino acids that include Y650 were deleted (lane 7). To determine if Y650 is required for the binding of c-Src to PMR60° the experiment in B was repeated with cells transfected with wild-type PMR60° or the Y650F mutation (Yang et al., 2004), and results in Fig. 3C show that indeed Y650 is required for the binding of c-Src to PMR60°.
The SH3 domain of c-Src binds to proline-rich sequences and the N-terminal deletion that results in loss of c-Src binding includes a pair of PPXXP motifs spanning amino acids 261–265 and 269–273. To determine if either or both of these is involved in the interaction with c-Src two prolines in each repeat (P262 and P265; P270 and P273) were changed to alanine, and the resulting constructs were co-transfected with WT-Src. Results in Fig. 3D show that neither of these mutations affects PMR60° expression or its recovery with myc antibody, and both of these mutations prevent the recovery of c-Src with PMR60°. Western blotting of recovered PMR60° with PY20 confirmed that tyrosine phosphorylation was lost with each mutation (Fig. 3E). Together these data demonstrate that the interaction of c-Src with PMR60° involves both of the proline-rich SH3 binding domains in the N-terminal 100 amino acids and the tyrosine phosphorylation site at Y650.
The signature feature of PMR60-mediated mRNA decay is the formation of a polysome-bound complex of the tyrosine-phosphorylated protein with its substrate mRNA. If c-Src is the responsible kinase the active form of this enzyme should activate polysome targeting in cells with little c-Src whereas inactive forms should not. The Cos-1 cells used in the previous experiments were co-transfected with PMR60° and GFP plus empty vector or plasmids expressing CA-Src, DN-Src, or one lacking kinase activity (kinase inactive, KI-Src). The expression of each of these is shown in Fig. 4A. Cytoplasmic extracts of these cells were applied to discontinuous sucrose density gradients and the top, mRNP (interface) and polysome (pellet) fractions were analyzed as before (Yang and Schoenberg, 2004) for PMR60°, GFP and ribosomal protein S6 (Fig. 4B). In cells that are co-transfected with empty vector the majority of PMR60° remained at the top of the gradient (lanes 1–3), and PMR60° behaves similarly in cells expressing DN-Src and KI-Src. The converse is seen in cells expressing CA-Src, with the majority of PMR60° now sedimenting with polysomes. These data confirm that phosphorylation by c-Src activates polysome targeting by PMR60°.
To confirm that c-Src activated polysome targeting activates PMR60-mediated mRNA decay Cos-1 cells were transfected as above with the addition of plasmids expressing target (albumin) and non-target (luciferase) mRNAs (Yang and Schoenberg, 2004). The expression of PMR60, c-Src and luciferase is shown in Fig. 4C, and Fig. 4D shows the impact of these treatments on albumin and luciferase mRNA assayed by RNase protection. In cells without added c-Src (lanes 4–5) or expressing DN-Src (lanes 8–9) there was no evidence for PMR60-mediated decay of albumin mRNA. In contrast CA-Src activated PMR60-mediated decay of albumin mRNA without affecting luciferase mRNA (lanes 6–7).
The general decay process involving decapping and exonuclease degradation involves mRNAs that are no longer engaged by translating ribosomes (Coller and Parker, 2005; Parker and Song, 2004). This contrasts with endonuclease-mediated mRNA decay, which involves targeting of a specific endonuclease to translating substrate mRNA (Yang and Schoenberg, 2004; Hollien and Weissman, 2006). Since decapping and exonuclease decay is the default processes there must be a fundamental code that distinguishes mRNAs that are targeted by endonuclease decay. This code likely has sequence and/or structure determinants in the mRNA itself that engage a specific cohort of RNA binding proteins which in turn recruit the mRNA endonuclease. Previous work from our lab showed that in Xenopus liver PMR60 initiates endonuclease-mediated decay when bound to polysomes (Cunningham et al., 2001), and decay requires active translation of its substrate mRNA (Yang and Schoenberg, 2004). The first major insight into the selectivity of this process was the discovery that endonuclease-mediated decay required the phosphorylation of PMR60 on a specific tyrosine residue (Y650) within a domain that is required for targeting to its translating substrate mRNA (Yang et al., 2004). The resulting phosphopeptide sequence is similar but not an exact match to any of the mapped SH2 ligands (Liu et al., 2006), indicating that a yet-to-be-identified SH2 domain protein is involved in recruiting PMR60 to polysomes.
A number of serine/threonine kinases have been linked to mRNA decay (eg. Schmidlin et al., 2004; Rutault et al., 2001; Chen et al., 1998), but this has not been the case for tyrosine kinases. Since tyrosine phosphorylation is required to ‘license’ PMR60 targeting to polysomes the goal of this study was to identify the responsible kinase. The key to finding c-Src was the observation that PMR60° recovered from glycerol gradient fractions yielded a single band when probed with antibody to its epitope tag and two bands when probed with PY20. Since tyrosine kinases commonly form a complex with their substrate and undergo autophosphorylation, we reasoned that the second (smaller) band might be the PMR60° kinase. This was confirmed in Fig. 1, where the size of this kinase matched that of c-Src, and in subsequent experiments that showed c-Src binds to and phosphorylates PMR60° in vitro and in vivo. The in vivo phosphorylation of PMR60° can be blocked with the c-Src inhibitor PP2 but not its inactive congener (Fig. 1E, Fig. 2D). These data plus those in Fig. 4, which show that active c-Src is required for both targeting of PMR60 to polysomes and PMR60-mediated mRNA decay, confirm that c-Src plays and essential role in endonuclease-mediated mRNA decay.
The use of various forms of c-Src and a line of Cos-1 cells with relatively little c-Src facilitated the mapping of interacting sites on PMR60. The SH2 and SH3 domains of c-Src bind phosphotyrosine and proline-rich ligands, respectively, and results in Fig. 3 show that these features of PMR60° are required for its interaction with the kinase. PMR60° has two repeating PPXXP motifs within an N-terminal portion of the protein, each of which is required for its interaction with c-Src. The recovery of c-Src with PMR60° was lost when either of these was changed to PAXXA, indicating that the extended sequence is likely involved in c-Src binding. As noted above, c-Src was not recovered with Y650F PMR60°.
c-Src is commonly elevated in cancer and enhanced activity of Src-mediated signal transduction is a recurring theme in malignancy (Ishizawar and Parsons, 2004). EGF binding to EGFR is one of the best characterized processes that activates c-Src and results in Fig. 2C and D show that EGF rapidly activates c-Src-mediated phosphorylation of PMR60°. The finding that ‘licensing’ of PMR60 to target to polysomes is an early event in signaling through c-Src raises the interesting and important possibility that elevation of c-Src might enhance the growth of malignant cells by increasing the selective degradation of mRNAs that are targeted by endonuclease-mediated decay. This will be addressed by work in progress aimed at identifying the mRNAs present in polysome-bound complexes with PMR60°.
The preparation of plasmids expressing catalytically active PMR60, catalytically inactive PMR60°, Y650F mutant, and PMR60° deletions were described previously (Yang et al., 2004; Yang and Schoenberg, 2004). The mutations P262,265A and P270,273A were generated by site-directed mutagenesis of the proline residues at positions 262, 265, 270, and 273 using the QuikChange site-directed mutagenesis kit (Stratagene). Sense primer 5′- GTGCCAAAGAGCCTGCTTGCTTTGCCCTGAAGATTCCAC-3′ and antisense primer 5′-GTGGAATCTTCAGGGCAAAGCAAGCAGGCTCTTTGGCAC-3′ were used to generate P262,265A. Sense primer 5′-CCCTGAAGATTCCAGCAAATGACGCTCGAATTAGTAATCAGAG-3′ and antisense primer 5′-CTCTGATTACTAATTCGAGCGTCATTTGCTGGAATCTTCAGGG-3′ were used to generate P270,273A. The empty vector pUSEamp and plasmids encoding wild-type (WT-Src), kinase-inactive (KI-Src), dominant-negative (DN-Src), or constitutively-active (CA-Src) forms were from Upstate Biotechnology. The plasmid pET-32b-PMR was constructed by inserting the entire PMR60 sequence into pET32b between NcoI and NotI sites.
Monoclonal antibody to the c-Myc epitope tag (9E10), myc antibody-coupled beads, antibodies to GFP (B2), c-Src (SRC2 and B-12) and ribosomal protein S6 (E13) were purchased from Santa Cruz Biotechnology. PY20 was purchased from BD Biosciences, and horseradish peroxidase-coupled rabbit anti-mouse IgG and goat anti-rabbit IgG were purchased from Santa Cruz.
Cos-1 cells were cultured in Dulbecco’s modified Eagle’s medium plus 10% fetal bovine serum and 2 mM glutamine. U2OS cells were cultured in McCoy’s 5A medium plus 10% fetal bovine serum. For most experiments, 2.5 × 106 cells in log phase growth were transfected with 10 μg (total) of plasmid DNA by using LipofectAMINE (Invitrogen) following the manufacturer’s protocol. Unless otherwise indicated cells were collected 40 hr after transfection. To study EGF activation of c-Src Cos-1 cells were serum starved for 16 hr prior to adding 100 ng/ml EGF. To inhibit c-Src in these cells 10 μM PP3 or PP2 was added to the culture 30 min before addition of EGF.
Cytoplasmic extracts were prepared as described previously (Yang and Schoenberg, 2004). Briefly, cells were washed twice with ice-cold phosphate-buffered saline, then suspended in cell lysis buffer (10 mM HEPES-KOH, pH 7.5, 10 mM KCl, 5 mM MgCl2, 50 mM NaF, 0.5% Nonidet P-40 (v/v), 2 mM dithiothreitol, 0.5 mM phenylmethylsulfonyl fluoride, 25 μl/ml protease inhibitor mixture (Sigma), 10 μl/ml phosphatase inhibitors (Sigma), and 10 μl/ml RNaseOUT (Invitrogen)). After incubation for 15 min on ice the cells were homogenized with 30 strokes of a Dounce homogenizer (A pestle), and the homogenate was centrifuged for 15 min at 2,000 × g. Polysomes and mRNP complexes were separated on discontinuous sucrose gradients containing 1 ml each of 10% and 35% (w/v) sucrose in the buffers described above (Schoenberg and Cunningham, 1999). Postmitochondrial extract was fractionated by sedimentation for 1 hr at 147,000 × g in a Sorvall Discovery M120 SE ultracentrifuge using a S55S-1123 rotor. Polysomes were collected from the bottom of the tube and mRNP complexes were collected at the interface between the 10% and 35% sucrose layers.
The plasmid pET32b-PMR was transformed into E. coli BL21(DE3). Cells were grown to an optical density (O.D.)600 of ~1.2 and induced with 0.5 mM IPTG for 4 hr at 37°C. Cells were lyzed by sonication in the buffer containing 1 X PBS, 5 mM EDTA, 5 mM DTT, 1 mg/ml lysozyme and 1 mM PMSF. After centrifugation, the pellets were extensively washed with 1 X PBS, 5 mM EDTA, 5 mM DTT, 2 M Urea, 2% Triton X-100, then dissolved in 20 mM NaH2PO4, 500 mM NaCl, 6 M guanidine-HCl, 2 mM beta-mercaptoethanol, pH8.0. The clarified lysate was loaded onto HiTrap chelating HP column (Amersham Bioscience). After extensive washing with 20 mM NaH2PO4, 500 mM NaCl, 10 mM imidazole, pH7.5, bound proteins were eluted with a linear gradient of 10 mM to 500 mM imidazole. The purified proteins were renatured in 20 mM Tris-HCl, pH 8.5, 1 mM reduced glutathione, 0.1 mM oxidized glutathione, 9.6 mM NaCl, 0.4 mM KCl, and further purified by ion exchange chromatography on a Fast-flow DEAE-Sepharose column.
For immunoprecipitation with myc antibody cell lysates were incubated with monoclonal antibody-coupled beads on a rocking platform for 3 hr at 4°C. For immunoprecipitation with rabbit anti-Src antibody cell lysates were incubated with antibody for 3 hr at 4°C followed by 20 μl of protein A-agarose (Santa Cruz Biotechnolgy) and incubation for 1 hr on a rocking platform at 4°C. The beads were washed four times with IPP150 buffer (10 mM Tris-HCl, pH7.5, 150 mM NaCl, 0.1% NP-40) and suspended in SDS-PAGE sample buffer. For Western blot analysis, the immunoprecipitates were separated on a 10% SDS-PAGE and electroblotted onto Immobilon-P membrane (Millipore). The membrane was blocked for 1 hr at room temperature in 5% nonfat dry milk in TBST buffer (20 mM Tris-HCl, pH7.5, 150 mM NaCl, 0.1% NP-40), then incubated with the primary antibody for 4 hr at room temperature, washed and incubated with horseradish peroxidase-conjugated secondary antibody for 1 hr. Blots were developed with SuperSignal west pico chemiluminescent substrate (Pierce).
HPLC purified N-terminal biotin-labeled peptides containing the tyrosine phosphorylation site of PMR60 (ARDGDRFFYEQP) without or with phosphotyrosine at position 9 (corresponding to Y650) were obtained from American Peptide Company, Inc. (Sunnydale, CA). 6 units of purified recombinant c-Src (Upstate Biotechnology, Inc) was incubated with 250 μM of each synthetic peptide or different amounts of recombinant PMR60 dissolved in 50 μl of 25 mM Tris-HCl, pH 7.5, 30 mM MgCl2, 12.5 mM MnCl2, 1 mM EGTA, 60 μM sodium orthovanadate, 1 mM dithiothreitol). Reactions were initiated by adding of 1 μl of γ-[32P]ATP (3,000 Ci/mmol, PerkinElmer) and incubated for 30 min at 30ºC. The reaction was stopped by the addition of SDS sample buffer, and phosphorylated samples separated by SDS-PAGE were visualized by PhosphorImager.
8 × 105 cells were transiently transfected with plasmids expressing albumin and luciferase mRNA and co-transfected with empty vector (pcDNA3), or plasmid expressing catalytically active myc-PMR60, plus vector pUSEamp or plamids encoding CA- or DN -Src. Total RNA was isolated with TRIzol reagent (Invitrogen). The antisense albumin riboprobe was prepared with the MAXIscipt in vitro Transcription Kit (Ambion) using T7 promoter from a pcRII-Topo plasmid containing exons 14 and 15 of albumin cDNA. The antisense firefly luciferase riboprobe was synthesized with T3 promoter from a pBluescript(SK) plasmid containing the first 153 nucleotides of firefly luciferase cDNA. Ribonuclease protection assay was done as described previously (Yang and Schoenberg, 2004) with 5 μg of total RNA hybridized to 600 pg of each riboprobe using the Ribonuclease Protection Assay III Kit (Ambion). Protected probe was separated on a denaturing 6% polyacrylamide-urea gel and quantified by PhosphorImager analysis.
This work was supported by NIH grant GM38277 to D.R.S. Support for core facilities was provided by center grant P30 CA16058 to the OSU Comprehensive Cancer Center. We wish to thank Hidetada Matsuoka and Catherine van Vliet, and members of the Schoenberg lab for their helpful suggestions and comments on this work.
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