A Complex Signaling Pathway Regulates SRp38 Phosphorylation and Pre-mRNA Splicing in Response to Heat Shock

Molecular Cell

Article A Complex Signaling Pathway Regulates SRp38 Phosphorylation and Pre-mRNA Splicing in Response to Heat Shock Yongsheng Shi1 and James L. Manley1,* 1Department of Biological Sciences, Columbia University, New York, NY 10027, USA *Correspondence: [email protected] DOI 10.1016/j.molcel.2007.08.028

SUMMARY

Although pre-mRNA splicing is known to be regulated by cell signaling, the underlying mechanisms are poorly understood. SRp38 is a member of the SR protein family and, when dephosphorylated, functions as a general and potent splicing repressor in response to heat shock. Here we show that SRp38 is dephosphorylated by the phosphatase PP1, which is activated by dissociation of its inhibitors, including NIPP1. PP1 is targeted to SRp38 through direct interaction via its arginine/serine-rich (RS) domain. The specific dephosphorylation of SRp38 and not other SR proteins is determined largely by the low activities of SR protein kinases for it compared to other SR proteins. Finally, we show that 14-3-3 proteins associate with SRp38 and protect it from dephosphorylation under nonstress conditions, but dissociate upon heat shock. Together, our study delineates a complex mechanism involving multiple factors by which a stress signaling pathway regulates protein phosphorylation and, in turn, pre-mRNA splicing.

INTRODUCTION Pre-mRNA splicing is an essential step of gene expression for the vast majority of metazoan genes. As it is currently estimated that over two-thirds of human genes produce alternatively spliced mRNAs (Blencowe, 2006), alternative splicing has been recognized as a major mechanism for generating proteomic complexity (Maniatis and Tasic, 2002). Aberrant splicing has been linked to a broad spectrum of human diseases (Licatalosi and Darnell, 2006), underscoring the importance of splicing and its regulation. Like transcription, splicing can be regulated both on a global level and in a gene-specific manner by cell signaling, for example during the cell cycle and in response to cellular stress (Shin and Manley, 2004). However, the underlying mechanisms remain poorly understood.

Splicing takes place in the spliceosome, a macromolecular complex containing five snRNAs (U1, U2, U4, U5, and U6) and over 100 proteins (Jurica and Moore, 2003). SR proteins constitute a family of splicing factors that play critical roles in both constitutive and alternative splicing (Manley and Tacke, 1996; Graveley, 2000). All members of this family contain one or two RNA-binding domains and a domain rich in serine/arginine dipeptide repeats (RS domain). SR proteins can function by both RNA sequence dependent and sequence independent mechanisms. SR proteins play an essential role in constitutive splicing, at least in part by stabilizing the association between snRNPs and the substrate RNAs, thereby facilitating spliceosome assembly. The functions of SR proteins in alternative splicing often involve direct binding to specific sequences in the target exons, called exonic splicing enhancers (ESEs). The ESEbound SR proteins can bridge splicing factors on the splice sites at both ends of the exon and improve exon recognition. The RS domains of SR proteins are heavily phosphorylated (Manley and Tacke, 1996; Graveley, 2000). This modification has been shown to modulate interactions of SR proteins with other splicing factors, and is critical for the function of SR proteins in splicing (Xiao and Manley, 1997). As both hyper- and hypophosphorylation inhibit splicing (Prasad et al., 1999), SR protein phosphorylation must be subject to tight regulation by both kinases and phosphatases. There are two major families of SR protein kinases, the SRPK family and the Clk/Sty family (Misteli, 1999). Kinases from both families can phosphorylate SR proteins in vitro, and when overexpressed, both disrupt the localization of SR proteins to nuclear speckles in vivo. There are also important differences between these two families, in both enzymatic properties (Velazquez-Dones et al., 2005) and subcellular localizations (Gui et al., 1994; Colwill et al., 1996). Phosphatases also play important roles in regulating SR protein phosphorylation. Serine/threonine-specific phosphatases are classified into PP1, PP2A, PP2B, and the more diverged PP2C families (Cohen 1990). Although both PP1/PP2A phosphatases and PP2Cg are required for splicing (Murray et al., 1999; Shi et al., 2006), only PP1 and PP2A have been suggested to play a role in regulating SR protein phosphorylation (Misteli, 1999). Excess

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recombinant PP1 causes dephosphorylation of SR proteins and, in turn, splicing inhibition in vitro (Mermoud et al., 1994). Similarly, microinjection of recombinant PP1 into nuclei leads to dephosphorylation of SR proteins and disruption of their localization to speckles (Misteli and Spector, 1996). How PP1 specificity and activity are regulated in splicing is not well understood. PP1 is associated with many regulatory proteins, and it has been suggested that several of these proteins could modulate PP1 activity and/or target it to specific substrates or subcellular localizations (Cohen, 2002). Furthermore, it has been shown that, in certain cell types, Fas activation or heat shock induces de novo ceramide synthesis, which then leads to PP1 activation and dephosphorylation of SR proteins (Chalfant et al., 2001; Jenkins et al., 2002). These results suggest that SR proteins are regulated by multiple kinases and phosphatases. But it is unclear how the activities of these kinases and phosphatases are coordinated in controlling SR protein phosphorylation. SRp38 is a member of the SR protein family with unique properties (Cowper et al., 2001; Shin and Manley, 2002; Shin et al., 2004). Although structurally similar to canonical SR proteins, SRp38 fails to activate splicing in vitro like other SR proteins. When dephosphorylated, however, it functions as a potent and general splicing repressor, at least in part by disrupting the association of snRNPs and pre-mRNAs and preventing spliceosome assembly. SRp38 becomes dephosphorylated during mitosis and in response to heat shock and is responsible for suppressing splicing under these conditions. SRp38-deficient cells have a prolonged G2/M phase and are not able to recover well from severe heat shock (Shin et al., 2004), underscoring the critical role of SRp38-mediated splicing repression in cell-cycle progression and cell survival under stress conditions. In this study, we set out to elucidate how SRp38 is targeted for dephosphorylation during heat shock. We first show that SRp38 is dephosphorylated by the phosphatase PP1. PP1 is activated by dissociation from an inhibitor, NIPP1, during heat shock. PP1 targets SRp38 through direct interaction mediated via its RS domain. Strikingly, the specific dephosphorylation of SRp38 and not other SR proteins is determined by the low activity of SR protein kinases toward SRp38. Finally, we show that SRp38 associates with 14-3-3 proteins, and they play a significant role in protecting SRp38 from dephosphorylation in the absence of stress. Thus a surprisingly complex pathway controls SRp38 phosphorylation status.

phorylated (Shin et al., 2004). Because this ‘‘in vitro heat shock’’ assay recapitulates the heat-induced SRp38 dephosphorylation observed in vivo, we used it in our initial experiments to identify the phosphatase(s). We first wished to determine whether the major serine/threonine phosphatases PP1 and PP2A are involved. For this purpose, we added to NE increasing concentrations of okadaic acid (OA), a specific inhibitor of the PP1 and PP2A family phosphatases (Cohen, 1991), and performed the in vitro heat shock assay (Figure 1A). Heat-induced dephosphorylation of SRp38 was unaffected by OA up to 50 nM (Figure 1A, lane 6), a concentration known to completely inhibit PP2A. However, 100 nM-1 mM OA, which inhibits both PP1 and PP2A, abolished SRp38 dephosphorylation (Figure 1A, lane 7 and data not shown). This OA-sensitivity profile suggests that PP1 is the major SRp38-directed phosphatase in vitro. To obtain further evidence for this, we tested the effect of inhibitor-2 (I-2), a protein inhibitor that is highly specific for PP1 (Cohen, 1991), using the same assay. We observed that I-2 inhibited heat-induced SRp38 dephosphorylation in a dosedependent manner (Figure 1B). These results indicate that PP1 is responsible for dephosphorylating SRp38 during heat shock in vitro. We next wished to determine whether PP1 is responsible for dephosphorylating SRp38 following heat shock in vivo. To this end, we depleted PP1 in HeLa cells by RNAi and tested the effect of this on heat-induced SRp38 dephosphorylation. Because there are three PP1 isoforms (a, b, and g) encoded by different genes in HeLa cells (Trinkle-Mulcahy et al., 2001), three isoform-specific siRNAs were used in order to deplete PP1 completely. For comparison, we also depleted PP2A by RNAi. Western blotting shows that both PP1 and PP2A were specifically and efficiently depleted (Figure 1C). When these cells were subjected to heat shock, SRp38 dephosphorylation was significantly reduced in PP1-depleted cells but unaffected in PP2A-depleted cells (Figure 1D). To assess the relative contribution of each PP1 isoform in this process, we depleted each one individually by RNAi and subjected these cells to heat shock (Figures 1E and 1F). We found that depleting PP1a alone had no detectable effect, while depleting PP1b or PP1g resulted in a partial inhibition of SRp38 dephosphorylation. Each of these siRNAs individually had minimal effects on cell viability. Coupled with the in vitro results, an indirect effect is therefore unlikely. These results suggest that PP1b and PP1g are the major SRp38directed phosphatases during heat shock.

RESULTS

SRp38 Directly Associates with PP1 via Its RS Domain PP1 is either targeted to specific substrates via associated proteins (targeting factors) (Cohen 2002), or, in some cases, by directly binding to its substrates, such as Rb (Tamrakar and Ludlow, 2000) and Cdc25 (Margolis et al., 2003). To understand how PP1 is targeted to SRp38, we first examined whether PP1 can associate with SRp38 in vivo. Unfortunately, immunoprecipitations (IPs) using

SRp38 Is Dephosphorylated by PP1 during Heat Shock A prerequisite in elucidating the mechanism by which SRp38 is dephosphorylated in response to heat shock was to identify the phosphatase(s) involved. We previously showed that when HeLa nuclear extract (NE) was incubated at higher temperatures, SRp38 was rapidly dephos-

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Figure 1. SRp38 Is Dephosphorylated by PP1 in Response to Heat Shock (A) In vitro heat shock assay and the effect of okadaic acid (OA). HeLa nuclear extract (NE) was untreated, treated with CIP, or incubated at 45 C in the presence of 0, 2 nM, 10 nM, 50 nM, 1 mM, or 5 mM of OA for 30 min. SRp38 was monitored by western blotting. SRp38, dephosphorylated SRp38 (dSRp38), and the smaller isoform SRp38-2 are marked. (B) In vitro heat shock in the presence of 0, 1 nM, 10 nM, 100 nM, and 1 mM inhibitor-2 (I-2). (C) HeLa cells were transfected with control, siPP1, or siPP2A siRNAs. PP1, PP2A, and actin levels were monitored by western blotting. (D) HeLa cells transfected with control, siPP1, or siPP2A siRNAs were incubated at 37 C ( HS) or heat-shocked at 44 C for 40 min (+HS). SRp38 was monitored by western blotting. (E) HeLa cells were transfected with control, or siPP1a-, b-, g-specific siRNAs, and the three PP1 isoforms and actin were monitored by western blotting. (F) HeLa cells treated with control or siPP1a, b, g, or bg siRNAs were incubated at 37 C ( HS) or heat-shocked at 44 C for 40 min (+HS).

the SRp38 antibody were poor and our attempts to generate a stable cell line expressing epitope-tagged SRp38 were not successful. Therefore, we transfected HEK293 cells with a plasmid encoding Flag-tagged SRp38, and immunoprecipitated SRp38 using anti-Flag antibodies (Figure 2A). Western blotting showed that PP1, but not PP2A, coimmunoprecipitated with SRp38, suggesting that PP1 and SRp38 are in fact associated in vivo (Figure 2A). Next we wished to examine whether the association between SRp38 and PP1 is direct. To this end, we performed GST pull-down assays using purified GST or GST-PP1 (a, b, g isoforms) with purified 6His-tagged SRp38 (Figure 2B) or in vitro-translated SRp38 (data not shown). The results showed that SRp38 was specifically precipitated by all three PP1 isoforms, suggesting that all PP1 isoforms directly interact with SRp38 in vitro. This conclusion was confirmed by reciprocal pull-downs showing that purified 6His-tagged PP1 was specifically precipitated by GSTSRp38 (Figure 2C). In the same assay, however, no PP1 was precipitated by GST-SRp38 RBD domain (Figure 2C), indicating that the RS domain of SRp38 is required for its association with PP1. We also performed GST pull-downs with GST-PP1 and in vitro-translated SRp38 RS domain or the full-length protein (Figure 2D). Similar amounts of

SRp38 RS domain and full-length SRp38 were precipitated by GST-PP1, suggesting that the RS domain of SRp38 is both necessary and sufficient for PP1 binding. Because the RS domain is a common feature of all members of the SR protein family, these results raise the question of whether PP1 binds to other SR proteins as well. When in vitrotranslated ASF/SF2 and SC35 were used in the same GST-PP1 pull-down assay, they were both precipitated, and dephosphorylated, by GST-PP1 (Figure 2D). These results indicate that the RS domain contains or constitutes a unique PP1-binding motif and that PP1 has the potential to be a general regulator of SR proteins. Specificity of SRp38 Dephosphorylation during Heat Shock Is Determined by Differential Kinase Activities SRp38 is specifically dephosphorylated during heat shock such that other SR proteins are affected only modestly (Shin et al., 2004). Thus, our observation that PP1 can directly bind and dephosphorylate SR proteins in addition to SRp38 raises the question of how specific dephosphorylation of SRp38 is achieved during heat shock. A clue came from comparison of in vitro and in vivo heat shock, which revealed an interesting difference in the

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Figure 2. PP1 Can Directly Interact with SRp38 via Its RS Domain (A) SRp38 associates with PP1 in vivo. NE from HEK293 cells ( ) or HEK293 cells transfected with an SRp38-3Flag construct (+) was used for IP with anti-Flag antibodies. SRp38, PP1, and PP2A levels in the input (2%) and IPs were monitored by western blotting. (B) Direct interactions between SRp38 and PP1 in vitro. GST pull-down assays were performed using GST or GST-PP1a, b, or g with recombinant 6His-SRp38. 6His-SRp38 was monitored by western blotting. (C) GST pull-down assays were performed using GST, GST-RBD, or GST-SRp38 with recombinant 6His-PP1a. 6His-PP1 was monitored by western blotting. (D) GST pull-down assays were performed using GST or GST-PP1g with in vitro-translated 35S-labeled proteins as indicated. Proteins were detected by autoradiography.

specificity of SR protein dephosphorylation. After HeLa cells were subjected to a heat shock (45 C for 40 min), western blotting of cell lysates (Figure 3A) revealed that more than 50% of SRp38 was fully dephosphorylated and the rest was partially dephosphorylated. In contrast, the great majority of ASF/SF2 was unaltered, with only a small percentage partially dephosphorylated. Western blotting with the mAb104 antibody, which specifically recognizes phosphorylated SR proteins, revealed that the overall phosphorylation status of the major SR proteins changed very modestly, results consistent with our previous study (Shin et al., 2004). When we examined the SR protein phosphorylation status during in vitro heat shock, however, the results were in sharp contrast. As shown in Figure 3B (lane 3), SRp38 was almost fully dephosphorylated after 30 min at 45 C. But unexpectedly, the majority of ASF/SF2 and SRp55 were also fully dephosphorylated under these conditions (Figure 3B, lanes 3 of the middle and bottom panels), indicating that the specificity of dephosphorylation observed in vivo was absent in vitro. The above results on the one hand establish a difference between in vitro and in vivo heat shock, but on the other suggest a possible explanation for why SRp38 is specifically targeted. As described in the Introduction, both kinases and phosphatases are known to control SR protein

phosphorylation status in vivo. However, because ATP was not added in the in vitro heat shock assays, kinase activities were excluded. Thus, it is possible that kinases are involved in determining the apparent specificity of SRp38 dephosphorylation. By this model, SRp38 would be a relatively poor substrate for SR protein kinases, compared to other SR proteins. To test this possibility, we performed in vitro heat shock in the presence of Mg2+ and ATP (Figure 3B, lanes 4 and 5). Under these conditions, the majority of SRp38 was still dephosphorylated (lane 5), somewhat less completely than in the absence of ATP, but similarly to the level of dephosphorylation detected in vivo (compare lane 5 of Figure 3B and lane 2 of Figure 3A). Strikingly, however, ASF/SF2 and SRp55 remained almost fully phosphorylated after heat shock (Figure 3B, lane 5 of the middle and bottom panel), very similar to what was observed in vivo. SRp40 though was partially dephosphorylated under these conditions. These results suggest that SR protein kinases are involved in determining the specificity of the heat-induced dephosphorylation. Because PP1 targets SRp38 and other SR proteins similarly, our hypothesis is that kinase activities targeting ASF/SF2 and other SR proteins are high enough to overcome PP1 activity and keep them in a phosphorylated form. On the other hand, kinase activity directed against SRp38 must be

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Figure 3. The Specificity of SRp38 Dephosphorylation Following Heat Shock Is Principally Determined by Differential Kinase Activities (A) Change of phosphorylation status of SRp38 and other SR proteins during heat shock in vivo. HeLa cells were incubated at 37 C ( HS) or at 45 C for 30 min (+HS). SRp38, ASF, and phospho-SR proteins were monitored by western blotting. (B) Change of phosphorylation status of SRp38 and other SR proteins during heat shock in vitro. HeLa NE was heat-shocked at 45 C for 15 or 30 min with or without 1 mM ATP. SRp38, ASF, and phospho-SR proteins were monitored by western blotting. Dephosphorylated ASF was marked as dASF. (C) Dephosphorylated SRp38 (d6His-SRp38) and ASF (d6His-ASF) were resolved by SDS-PAGE and stained with Coomassie. (D) d6His-ASF (left panel) or d6His-SRp38 (50 ng) was incubated with GST-Clk1 (50 ng) for 0, 30, or 60 min. Phosphorylated ASF or SRp38 was loaded for comparison. (E) d6His-ASF (left panel) or d6His-SRp38 (right panel) was incubated with 6His-SRPK1 (200 ng) for 0, 30, or 60 min. Phosphorylated ASF or SRp38 was loaded for comparison. (F) d6His-ASF (left panel) or d6His-SRp38 (right panel) was incubated with 6His-SRPK1 (200 ng) in the presence of [g-32P]ATP for the indicated time. Incorporation of 32P was monitored by autoradiography.

lower and insufficient to rephosphorylate SRp38 during heat shock. To test the above hypothesis, we compared the activities of the two major SR protein kinases, Clk/Sty and SRPK, toward SRp38 and ASF/SF2. We first prepared unphosphorylated SRp38 (dSRp38) and ASF/SF2 (dASF) by treating 6His-tagged SRp38 and ASF/SF2 purified from baculovirus-infected insect cells with alkaline phosphatase (CIP), and the fully dephosphorylated proteins were repurified using nickel beads (Figure 3C). Equivalent amounts of unphosphorylated SRp38 and ASF/SF2 were then incubated with purified recombinant Clk/Sty or SRPK1 kinases in in vitro kinase assays. Upon incubation with Clk/Sty, ASF/SF2 was rapidly and efficiently phosphorylated, as indicated by its mobility shift on SDS-PAGE relative to fully

phosphorylated ASF/SF2 (Figure 3D, left panel). Under the same conditions, however, SRp38 mobility remained largely unchanged (Figure 3D, right panel), suggesting that SRp38 is a poor substrate for Clk/Sty kinase. When ASF/SF2 was incubated with SRPK1, a small shift in mobility was observed (Figure 3E, left panel), consistent with previous studies showing that SRPK only partially phosphorylates ASF/SF2 (Velazquez-Dones et al., 2005). SRp38 mobility was essentially unchanged after incubation with SRPK1 (Figure 3E, right panel). To confirm this result, [g-32P]ATP was used in the SRPK kinase assay (Figure 3F). ASF/SF2 was efficiently labeled but SRp38 was not, establishing that SRp38 is, indeed, a poor substrate for SRPK. These results suggest that SR proteins are dynamically regulated by both kinases and phosphatases, and that

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Figure 4. Induction of PP1 Activity against SRp38 by Heat Shock (A) Immunopurification of PP1g complexes. IP was performed with NE made from HEK293 or an HEK293 cell line that stably expresses Flag-PP1g. Immunoprecipitated proteins were resolved by SDS-PAGE and stained with silver (left panel) or subjected to western blotting (right panel). (Asterisk [*], nonspecific bands from IPs; arrows specify the bands corresponding to PNUTS, NIPP1, and PP1). (B) Phosphatase assay using immunopurified PP1g complexes. PP1g complexes were incubated with 6His-SRp38 at 30 C or 45 C for 30 min. SRp38 was detected by western blotting. (C) NIPP1 inhibits PP1 activity toward SRp38. 6His-SRp38 was incubated with 6His-PP1 (50 ng) alone (lane 2) or with increasing amounts of GST-NIPP1 (25, 50, 100, or 200 ng) for 30 min at 30 C. (D) Changes in PP1 complexes during heat shock in vivo. NE was prepared from HeLa cells that have been heat-shocked for the indicated time, and PP1 was precipitated with microcystin beads. PP1, NIPP1, and PNUTS present in the pull-down (PD, 40% was loaded) and supernatants (SN) were monitored by western blotting. (E) Overexpression of NIPP1 inhibits SRp38 dephosphorylation. HeLa cells transfected with empty vector ( ) or a NIPP1-3Flag construct were untreated or subjected to heat shock. NIPP-1 and SRp38 were monitored by western blotting. Relative amounts of SRp38 and dSRp38 were quantified with the Scion Image program, and the ratio (dSRp38/SRp38) was listed.

the specificity of heat-induced dephosphorylation of SRp38 is determined largely by differential kinase activities toward individual SR proteins.

Induction of PP1 Activity during Heat Shock How is PP1 activity toward SRp38 induced by heat shock? Because PP1 activity is often regulated by its associated factors, we first isolated and characterized the major nuclear PP1-containing complexes. To this end, we prepared NE from an HEK293 cell line that stably expresses Flag-PP1g, and immunoprecipitated PP1 complexes using anti-Flag antibodies under stringent conditions (Figure 4A, left panel). In addition to Flag-PP1g, two other prominent bands of 40 and 110 kDa were detected. The sizes of these species were very close to those of NIPP1 and PNUTS, two well-known nuclear PP1 regulators (Cohen, 2002). Western blotting confirmed that indeed the 40

and 110 kDa species correspond to NIPP1 and PNUTS, respectively (Figure 4A, right panel). To examine whether the immunopurified PP1 complex can dephosphorylate SRp38, we incubated the PP1 complexes with recombinant SRp38 at 30 C or 45 C. As shown in Figure 4B, a relatively low level of SRp38 dephosphorylation occurred at 30 C, but SRp38 was extensively dephosphorylated at 45 C, similar to what was observed in heat-treated NE (Figure 2B) or following heat shock in vivo (Figure 2A). These results suggest that the immunopurified PP1 complexes are active against SRp38. Furthermore, a temperature-sensing factor may be present in these PP1 complexes. Because the phosphatase activity of the immunopurified PP1 complexes is relatively low at 30 C, we wondered if this was due to the presence of inhibitors, NIPP1 and PNUTS. To test this, we examined the effect of recombinant GST-NIPP1 in a PP1 phosphatase assay. As shown in Figure 4C,

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recombinant PP1 efficiently dephosphorylated SRp38 at 30 C (lane 2) and NIPP1 inhibited SRp38 dephosphorylation in a dose-dependent manner (lanes 3–6). Therefore, NIPP1 keeps PP1 activity toward SRp38 low under normal conditions. Next we wished to examine whether the association of PP1 with NIPP1 and/or PNUTS changes during heat shock, which could potentially serve as a heat-sensing mechanism. To test this, we incubated HeLa cells under heat shock conditions for 0, 20, 40, and 60 min, and prepared NE from these cells. Then, endogenous PP1 complexes in these extracts were precipitated by using microcystin-conjugated beads (Figure 4D). Microcystin is a small-molecule inhibitor that binds strongly and specifically to PP1/PP2A phosphatases (MacKintosh et al., 1990), and microcystin beads have been used to precipitate or even deplete PP1/PP2A phosphatases from cell extracts (Shi et al., 2006). In untreated cells or cells that were heat-shocked for a short period of time (20 min), essentially all NIPP1 was precipitated by microcystin beads (Figure 4D, lanes 1–4), indicating that almost all NIPP1 was associated with PP1. In NE made from cells that were heat-shocked for 40 or 60 min, however, significantly less NIPP1 was precipitated by microcystin beads and concurrently, NIPP1 was now detected in supernatants (Figure 4D, lanes 5–8). This suggests that during heat shock, NIPP1 dissociates from PP1. In comparison, the association between PNUTS and PP1 did not change as significantly during heat shock in this assay (Figure 4D, bottom panel). If the dissociation of NIPP1 from PP1 contributes to activation of PP1 in vivo, then overexpression of NIPP1 should inhibit SRp38 dephosphorylation during heat shock. To test this, we transfected HeLa cells with a plasmid encoding Flag-tagged NIPP1 and examined its effect on SRp38 dephosphorylation (Figure 4E). The results showed that, indeed, significantly less SRp38 dephosphorylation occurred in NIPP1-overexpressing cells (compare lanes 3 and 4). Therefore, our results showed that NIPP1 inhibits PP1 activity toward SRp38 in vitro and in vivo, and NIPP1 dissociates from PP1 during heat shock. Together, these data suggest that during heat shock, PP1 is activated by dissociation of inhibitors, including NIPP1. 14-3-3 Proteins Associate with SRp38 and Protect It from Dephosphorylation Given that SRp38 is such a potent splicing repressor when dephosphorylated, we wondered whether there are mechanisms to ensure that the protein is not inappropriately dephosphorylated. To investigate whether other factors might bind to and regulate SRp38, we expressed Flag-tagged SRp38 in HeLa cells and immunoprecipitated SRp38-containing protein complexes with anti-Flag antibodies (Figure 5A). Coimmunoprecipitated proteins were subjected to mass spectrometry analysis. Among the factors identified were 14-3-33 and z (Figure 5A). Presence of the 14-3-3 proteins in the coimmunoprecipitated proteins

was confirmed by western blotting using an antibody that recognizes all 14-3-3 isoforms (Figure 5B). To examine whether endogenous SRp38 is associated with 14-3-3 proteins, we performed IPs using control or anti-14-3-3 antibodies with HeLa NE, and found that SRp38 was in fact coimmunoprecipitated (Figure 5C). We therefore conclude that SRp38 is associated with 14-3-3 proteins in vivo. Consistent with our conclusion, two recent proteomic analyses of 14-3-3-associated factors both identified SRp38 (Jin et al., 2004; Benzinger et al., 2005). To confirm the specific association between SRp38 and 14-3-33 and z, plasmids encoding Flag-SRp38 and HA-14-3-33 or z were cotransfected into HeLa cells and SRp38 immunoprecipitated using anti-Flag antibodies. Both 14-3-33 and z were specifically immunoprecipitated (Figure 5D). To examine whether these associations are direct, we performed GST pull-down assays using GST or GST-14-3-33 and z isoforms with purified 6xHis-SRp38. As shown in Figure 5E, SRp38 was specifically precipitated by GST-14-3-33 and z, suggesting that SRp38 can directly bind to these two isoforms. We next wished to identify the region of SRp38 required for 14-3-3 association. By examining the SRp38 amino acid sequence, we found two consensus 14-3-3binding motifs (Yaffe, 2002). We therefore mutated two critical residues in each motif (S185A/P187A and T255V/ P257A; see Figure S1 in the Supplemental Data available with this article online), and tested the effect of these mutations on 14-3-3 binding by using GST pull-down assays under relatively stringent conditions (500 mM NaCl in wash buffer) (Figure 5F). Equivalent amounts of SRp38 (S185A/P187A) and wild-type SRp38 were precipitated by GST-14-3-3z (lanes 6 and 7). However, significantly less SRp38 (T255V/P257A) was recovered (lane 8), indicating that the T255 site is a major 14-3-3-binding site in SRp38. To examine whether 14-3-3 proteins are involved in regulating SRp38 phosphorylation, we first tested the effect of 14-3-3 in a PP1 phosphatase assay (Figures 6A and 6B). Significantly, 14-3-3z inhibited SRp38 dephosphorylation by PP1 in a dose-dependent manner (Figure 6A, lanes 3–5), indicating that 14-3-3 can protect SRp38 from dephosphorylation in vitro. At 45 C, however, the protective effect of 14-3-3 was lost and SRp38 was dephosphorylated by PP1 (Figure 6B, lanes 6 and 7). One possible reason for the loss of protection by 14-3-3 at higher temperature is that 14-3-3 proteins might dissociate from SRp38 during heat shock. To examine whether this occurs in vivo, we transfected HEK293 cells with an SRp38-3Flag-encoding plasmid and immunoprecipitated SRp38 before and after the cells were subjected to heat shock. As shown in Figure 6C, equal amounts of SRp38 were immunoprecipitated before and after heat shock, and 14-3-3 proteins were coimmunoprecipitated before heat shock. However, significantly fewer 14-3-3 proteins were recovered after heat shock, suggesting that indeed 14-3-3 proteins dissociate from SRp38 during heat shock in vivo.

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Figure 5. SRp38 Associates with 14-3-3 Proteins In Vivo and In Vitro (A) Identification of SRp38-associated proteins. NE was made from HeLa cells transfected with an empty vector ( ) or an SRp38-3Flag construct (+), and SRp38-3Flag was immunoprecipitated using anti-Flag antibodies. Immunoprecipitated proteins were resolved by SDS-PAGE and stained with Coomassie. Proteins identified by MALDI-TOF were marked. (B) 14-3-3 proteins coimmunoprecipitated with SRp38. Immunoprecipitated proteins described in (A) were subjected to western blotting using anti-Flag (M2) and anti-14-3-3 pan antibodies. (l.c. is the light chain.) (C) Endogenous 14-3-3 and SRp38 associate in vivo. IP was performed with HeLa NE using control or 14-3-3 antibodies, and 14-3-3 and SRp38 were monitored by western blotting. (D) SRp38 associates with 14-3-33 and z. 293T cells were cotransfected with SRp38-3Flag and HA-14-3-3 plasmids as indicated. Flag-IPs were performed, and SRp38 and 14-3-3 were monitored by western blotting using anti-Flag or anti-HA antibodies. (E) Direct interactions between SRp38 and 14-3-3 in vitro. GST pull-down assays were performed using GST or GST-14-3-33 and z with purified 6His-SRp38. 6His-SRp38 was monitored by western blotting. (F) Mapping 14-3-3-binding site(s) in SRp38. GST pull-down assays were performed using GST or GST-14-3-3z with 35S-labeled in vitro-translated wild-type or mutant SRp38. SRp38 was visualized by autoradiography.

Finally, we wished to examine whether 14-3-3 plays a role in regulating heat-induced SRp38 dephosphorylation in vivo. We reasoned that if 14-3-3 protects SRp38 from dephosphorylation in vivo as it does in vitro, then the SRp38 mutant T255V/P257A, which has lower affinity for 14-3-3, should be dephosphorylated more efficiently than the wild-type SRp38 during heat shock. To test this, we transfected plasmids encoding wild-type SRp38 or the S185A/P187A and T255V/P257A mutants into HeLa cells, subjected these cells to heat shock, and analyzed lysates by western blotting (Figure 6C). Significantly, even without heat shock, T255V/P257A was partially dephosphorylated. Upon heat shock, wild-type SRp38 and S185A/P187A mutant were both partially dephosphorylated (lanes 4 and 5, Figure 6C). In contrast, T255V/ P257A was almost fully dephosphorylated (lane 6). Together, our results strongly suggest that 14-3-3 directly binds to SRp38 and plays a role in protecting SRp38 from dephosphorylation in vivo.

DISCUSSION Our results have elucidated a complex mechanism for the regulation of SRp38 phosphorylation. Under normal conditions, phosphorylated SRp38 is associated with 14-3-3 proteins, which help protect it from dephosphorylation. PP1 activity toward SRp38 is inhibited by its associated proteins, including NIPP1. During heat shock, PP1 dissociates from NIPP1 and directly binds to and dephosphorylates SRp38. Although ASF/SF2 and other SR proteins can also be targeted by PP1, the activities of SR protein kinases toward them are high enough to maintain them in a phosphorylated state, thereby establishing the observed specificity for SRp38. Below, we discuss this model (illustrated in Figure 7), the proteins involved, and its implications for other signaling pathways that involve regulated dephosphorylation. Three closely related isoforms of PP1 (a, b, and g) exist in mammalian cells (Cohen, 2002). These isoforms are

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Figure 6. 14-3-3 Proteins Protect SRp38 from Dephosphorylation In Vitro and In Vivo (A) 14-3-3 protects SRp38 from dephosphorylation by PP1 in vitro. Fifty nanograms purified 6His-SRp38 was mixed with 50 ng 6His-PP1 and increasing amounts of recombinant 14-33z (25, 50, and 100 ng) and incubated at 30 C for 30 min. (B) Protection by 14-3-3 is lost at heat shock temperature. 6His-SRp38 and 6His-PP1 were incubated without (lanes 2 and 3) or with 100 ng 14-3-3z (lanes 4–7) at indicated temperatures for 15 or 30 min. (C) 14-3-3 proteins dissociate from SRp38 during heat shock. HEK293 cells transfected with empty vector ( ) or SRp38-3Flag (+) constructs were untreated or heat-shocked, and SRp38-3Flag was immunoprecipitated. SRp383Flag and 14-3-3 proteins in lysates and IPs were monitored by western blotting. (D) 14-3-3 proteins protect SRp38 from dephosphorylation in vivo. HeLa cells transfected with HA-tagged wild-type or mutant SRp38 were untreated ( ) or heat-shocked (+HS). Transfected SRp38 was detected by western blotting with anti-HA antibodies.

more than 89% identical in amino acid sequences and have similar biochemical properties. Indeed, we found that recombinant versions of all three isoforms can efficiently dephosphorylate SRp38 in vitro. However, PP1b and g appear to be the major SRp38-directed phosphatases during heat shock. What might be the basis for this isoform specificity? The three PP1 isoforms have different subcellular localization patterns (Trinkle-Mulcahy et al., 2001), possibly due to associations with distinct targeting factors (Lesage et al., 2004). PP1a displays a diffused localization pattern while both PP1b and g are highly

enriched in the nucleoli. Interestingly, SRp38 is localized predominantly in small perinucleoli foci (Shin et al., 2005). Therefore, the spatial proximity of SRp38 and PP1b and g may explain the apparent PP1 isoform specificity in dephosphorylating SRp38. A major question in PP1-mediated protein dephosphorylation is how the phosphatase is targeted to specific substrates. Our results suggest that PP1 targets SRp38 through direct interaction with its RS domain. Furthermore, PP1 also binds to other SR proteins such as ASF/ SF2 and SC35 with similar affinity. Two highly degenerate

Figure 7. A Complex Pathway Controls SRp38 in Response to Heat Shock Under normal conditions, phosphorylated SRp38 is associated with 14-3-3 proteins, which help protect it from dephosphorylation. PP1 activity toward SRp38 is inhibited by its associated proteins, including NIPP1. During heat shock, PP1 dissociates from NIPP1 and directly binds to and dephosphorylates SRp38. Although ASF/SF2 and other SR proteins can also be targeted by PP1, the activities of SR protein kinases toward them are high enough to maintain them in a phosphorylated state, thereby establishing the observed specificity for SRp38.

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PP1-binding motifs have been characterized, the RVxF motif and the FxxR/KxR/K motif (Cohen, 2002). Given the large number of arginine residues present in RS domains, it is possible that RS domains contain a variant of one of these motifs. Alternatively, the RS domain itself may constitute a unique PP1-binding motif. The direct association between PP1 and RS domains could have significant implications for regulation of SR protein phosphorylation. Earlier studies indicated that ASF/SF2 and U1-70K (which also contains an RS domain) must be dephosphorylated during splicing (Cao et al., 1997; Tazi et al., 1993). Recently, it has been shown more directly that several SR proteins, such as 9G8, SRp20, and ASF/SF2, are specifically dephosphorylated during mRNA maturation, most likely in a splicing-dependent manner (Huang et al., 2004). It has been reported that 9G8 and ASF/SF2, but not SC35, relocalize to stress-induced nuclear bodies during heat shock (Denegri et al., 2001). It remains to be determined whether PP1 plays a role in regulating phosphorylation of these SR proteins during mRNA maturation and in response to stress. How does heat shock activate PP1? It has been previously suggested that, in certain cell types, heat shock can lead to acute de novo synthesis of ceramide, which in turn activates PP1 to dephosphorylate SR proteins (Chalfant et al., 2001). However, addition of exogenous ceramide did not affect dephosphorylation of SRp38 during heat shock in our experiments (unpublished data). Also, the fact that the heat-induced dephosphorylation of SRp38 can be reproduced in vitro suggests that new protein or lipid synthesis may not be necessary for PP1 activation by heat shock. In contrast, studies on many PP1 regulators suggest that the major mechanism for PP1 activation is through changes in the binding of regulatory factors (Cohen, 2002). Consistent with this, our results suggest that PP1 is activated during heat shock by dissociation of its inhibitors, such as NIPP1. The association of PP1 with both PNUTS and NIPP1, the two major nuclear PP1-interacting factors, is regulated by reversible phosphorylation and, intriguingly, by RNA binding (Jagiello et al., 1995; Kim et al., 2003). Heat shock is known to activate several mitogen-activated protein (MAP) kinase pathways (Dorion and Landry, 2002). It is thus possible that NIPP1 dissociation from PP1 during heat shock is induced by its phosphorylation by a MAP kinase. However, we found that inhibitors of major known MAP kinase pathways had no effect on SRp38 dephosphorylation (unpublished data). Another possibility is that the dissociation of PP1 and NIPP1 is triggered directly by heat-induced conformational changes. Previous studies showed that trimerization and DNA binding of the heat shock transcription activator, HSF1, can be directly induced by elevated temperatures (Zhong et al., 1998). A third possibility is that during heat shock NIPP1 may become associated with RNA molecules, which then alleviate NIPP1 inhibition of PP1. In fact, a recent study demonstrated that a noncoding RNA functions as an essential cofactor in fully activating HSF1 (Shamovsky et al., 2006).

An unexpected conclusion of our work is that the specificity of SRp38 dephosphorylation is due in large part to the activities of kinases. PP1 displays broad substrate specificity, and it is generally believed that this specificity is achieved through association with various targeting factors (Cohen, 2002). Over 45 putative targeting factors have been identified, although in most cases the specific targeted substrate(s) are not known. In this study, we observed that PP1 directly binds to and dephosphorylates SRp38 and other SR proteins with similar efficiency, and specific dephosphorylation of SRp38 is then determined by differential kinase activities. These observations underscore the importance of the balance between kinases and phosphatases in specifically controlling phosphorylation status. Our data also raise the question of which kinase(s) are responsible for phosphorylating SRp38. Analysis of the amino acid sequence of SRp38 revealed the presence of potential target sites for Akt and PKC kinases, all of which are located in the RS domain (unpublished data). In fact, two recent studies suggested that Akt is involved in phosphorylation of SR proteins and, in turn, alternative splicing (Blaustein et al., 2005; Patel et al., 2005). It will be interesting to determine whether Akt or PKC, or another kinase, is responsible for phosphorylating SRp38 in vivo. Our study has provided the first functional evidence that 14-3-3 proteins play a role in splicing regulation. 14-3-3 proteins constitute a family of highly conserved proteins well known for their ability to bind phospho-serine/threonine residues in a variety of proteins (Yaffe, 2002). There are at least seven isoforms in mammals encoded by different genes, and they form homo- or heterodimers. Binding of 14-3-3 proteins to target proteins can affect their enzymatic activities, stability, intracellular localization, and interactions with other proteins (Yaffe, 2002). Recently, two proteomic analyses of 14-3-3-associated factors identified a number of splicing factors (Jin et al., 2004; Benzinger et al., 2005), indicating that 14-3-3 proteins may also function in splicing regulation. It is highly likely that more functions of 14-3-3 proteins in splicing regulation will be uncovered in future studies. Comparison of SRp38 with another 14-3-3-associated protein, Cdc25, reveals similarities that may have implications for understanding the regulation of SRp38 phosphorylation during the cell cycle. Similar to SRp38, Cdc25 (Ser216) is dephosphorylated in mitosis. As with SRp38, the association with 14-3-3 proteins prevents inappropriate dephosphorylation of Cdc25 by PP1 (Margolis et al., 2003). These similarities suggest that SRp38 phosphorylation might be regulated by similar mechanisms as Cdc25 in mitosis. In the case of Cdc25, 14-3-3 release precedes Cdc25 dephosphorylation by PP1 prior to mitosis, and is induced by phosphorylation of Cdc25 at another residue by Cdk2 (Margolis et al., 2003). Interestingly, a potential Cdk site (SP) is found in SRp38 that in fact overlaps the 14-3-3-binding motif. Therefore, it will be of considerable interest in the future to determine whether this residue is phosphorylated by Cdk2 prior to mitosis, and whether

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the dissociation of 14-3-3 is involved in regulating SRp38 phosphorylation in M phase. In summary, our results elucidated the detailed mechanisms for regulation of SRp38 phosphorylation, and revealed a network involving kinases, phosphatases, and other factors that regulates pre-mRNA splicing in response to cellular stress. Heat shock-induced splicing inhibition has been observed from yeast to human, suggesting that this is an important survival mechanism throughout evolution. Interestingly, most of the factors involved in regulating SRp38, such as PP1, SR protein kinases, and 14-3-3, are conserved from yeast to human. There is also an SR-like protein in yeast, Npl3p, whose phosphorylation is known to be regulated by Glc7 (PP1 homolog in yeast) and Sky1 (SRPK homolog in yeast) (Gilbert and Guthrie, 2004). Therefore, this ancient signaling network might have evolved to play important roles in regulation of RNA metabolism both under normal conditions and in response to stress. EXPERIMENTAL PROCEDURES

Supplemental Data Supplemental Data include Supplemental Experimental Procedures and one figure and can be found with this article online at http:// www.molecule.org/cgi/content/full/28/1/79/DC1/.

ACKNOWLEDGMENTS We thank Drs. L. Trinkle-Mulcahy, P. Greengard, A. Nairn, H. Fu, M. Yaffe, T. Pawson, X-D. Fu, and J. Adams for providing reagents; and members of the Manley lab for helpful discussions. Y.S. was supported by an NIH postdoctoral fellowship (F32GM070113), and this work was supported by NIH grant R01 GM 48259. Received: May 13, 2007 Revised: July 20, 2007 Accepted: August 2, 2007 Published: October 11, 2007 REFERENCES Benzinger, A., Muster, N., Koch, H.B., Yates, J.R., III, and Hermeking, H. (2005). Targeted proteomic analysis of 14-3-3 sigma, a p53 effector commonly silenced in cancer. Mol. Cell. Proteomics 4, 785–795. Blaustein, M., Pelisch, F., Tanos, T., Munoz, M.J., Wengier, D., Quadrana, L., Sanford, J.R., Muschietti, J.P., Kornblihtt, A.R., Caceres, J.F., et al. (2005). Concerted regulation of nuclear and cytoplasmic activities of SR proteins by AKT. Nat. Struct. Mol. Biol. 12, 1037–1044.

Plasmids, Recombinant Proteins, and Antibodies A complete list of materials can be found in the Supplemental Data. Cell Culture and Transfections HeLa, HEK293, and 293T cells were maintained in DMEM plus 10% fetal bovine serum. DNA transfections were performed using calcium phosphate method or Lipofectamine 2000 (Invitrogen) following manufacturer’s instructions. For making stable cell lines, Flag-PP1gpCDNA3 was transfected into HEK293 cells and stable transfectants were selected with Geneticin (Invitrogen). For RNAi, siRNA duplexes were synthesized by Dharmacon, and transfections were performed for two rounds using Lipofectamine 2000 (Invitrogen). Heat shock was performed 48–60 hr after the second transfection as described (Shin et al., 2004).

Blencowe, B.J. (2006). Alternative splicing: new insights from global analyses. Cell 126, 37–47.

Immunoprecipitation and Protein Binding Assays For immunoprecipitations (IPs), NE was incubated with antibody-conjugated beads and for 4 hr with rotation at 4 C, followed by three washes with buffer D-300 mM NaCl. To immunopurify Flag-PP1g complex for activity assays, more stringent washing buffer (buffer D-1 M NaCl, 1% NP-40) was used and Flag-PP1g eluate was concentrated using Centricon filters (Millipore). For identification of SRp38-associated factors, immunoprecipitated proteins were resolved by SDSPAGE and stained with Coomassie. Protein bands were excised, ingel trypsin digested, and analyzed by MALDI-TOF using a PerSeptive Biosystems Voyager-DE. Peptide peaks were searched against protein databases using the MS-Fit program on the ProteinProspector server (UCSF). For GST pull-down assays, 4 mg of GST or GST fusion proteins were bound to glutathione beads before being mixed with in vitro-translated 35S-labeled proteins or purified recombinant proteins plus 1 mg/ml BSA in buffer D. The mixture was incubated for 2 hr with rotation at 4 C, followed by three washes with buffer D-300 or buffer D-500 (Figure 5F).

Cohen, P. (1991). Classification of protein-serine/threonine phosphatases: identification and quantitation in cell extracts. Methods Enzymol. 201, 389–398.

Phosphatase and Kinase Assays For phosphatase assays, 30 ng of 6His-SRp38 was mixed with indicated amounts of 6His-PP1a or GST-PP1g in buffer D plus 1 mg/ml BSA and 1 mM MnCl2, and incubated at 30 C or 45 C. For kinase assays, 30 ng d6His-SRp38 or d6His-ASF were incubated with indicated amounts of GST-Clk/Sty or 6His-SRPK1 in buffer D plus 1 mg/ml BSA, 1 mM ATP, and 10 mM MgCl2.

Cao, W., Jamison, S.F., and Garcia-Blanco, M.A. (1997). Both phosphorylation and dephosphorylation of ASF/SF2 are required for premRNA splicing in vitro. RNA 3, 1456–1467. Chalfant, C.E., Ogretmen, B., Galadari, S., Kroesen, B.J., Pettus, B.J., and Hannun, Y.A. (2001). FAS activation induces dephosphorylation of SR proteins; dependence on the de novo generation of ceramide and activation of protein phosphatase 1. J. Biol. Chem. 276, 44848–44855. Cohen, P. (1990). The structure and regulation of protein phosphatases. Adv. Second Messenger Phosphoprotein Res. 24, 230–235.

Cohen, P.T. (2002). Protein phosphatase 1—targeted in many directions. J. Cell Sci. 115, 241–256. Colwill, K., Pawson, T., Andrews, B., Prasad, J., Manley, J.L., Bell, J.C., and Duncan, P.I. (1996). The Clk/Sty protein kinase phosphorylates SR splicing factors and regulates their intranuclear distribution. EMBO J. 15, 265–275. Cowper, A.E., Caceres, J.F., Mayeda, A., and Screaton, G.R. (2001). Serine-arginine (SR) protein-like factors that antagonize authentic SR proteins and regulate alternative splicing. J. Biol. Chem. 276, 48908– 48914. Denegri, M., Chiodi, I., Corioni, M., Cobianchi, F., Riva, S., and Biamonti, G. (2001). Stress-induced nuclear bodies are sites of accumulation of pre-mRNA processing factors. Mol. Biol. Cell 12, 3502–3514. Dorion, S., and Landry, J. (2002). Activation of the mitogen-activated protein kinase pathways by heat shock. Cell Stress Chaperones 7, 200–206. Gilbert, W., and Guthrie, C. (2004). The Glc7p nuclear phosphatase promotes mRNA export by facilitating association of Mex67p with mRNA. Mol. Cell 13, 201–212. Graveley, B.R. (2000). Sorting out the complexity of SR protein functions. RNA 6, 1197–1211.

Molecular Cell 28, 79–90, October 12, 2007 ª2007 Elsevier Inc. 89

Molecular Cell Complex Regulation of SRp38 Phosphorylation

Gui, J.F., Lane, W.S., and Fu, X.D. (1994). A serine kinase regulates intracellular localization of splicing factors in the cell cycle. Nature 369, 678–682.

Murray, M.V., Kobayashi, R., and Krainer, A.R. (1999). The type 2C Ser/Thr phosphatase PP2Cgamma is a pre-mRNA splicing factor. Genes Dev. 13, 87–97.

Huang, Y., Yario, T.A., and Steitz, J.A. (2004). A molecular link between SR protein dephosphorylation and mRNA export. Proc. Natl. Acad. Sci. USA 101, 9666–9670.

Patel, N.A., Kaneko, S., Apostolatos, H.S., Bae, S.S., Watson, J.E., Davidowitz, K., Chappell, D.S., Birnbaum, M.J., Cheng, J.Q., and Cooper, D.R. (2005). Molecular and genetic studies imply Akt-mediated signaling promotes protein kinase CbetaII alternative splicing via phosphorylation of serine/arginine-rich splicing factor SRp40. J. Biol. Chem. 280, 14302–14309.

Jagiello, I., Beullens, M., Stalmans, W., and Bollen, M. (1995). Subunit structure and regulation of protein phosphatase-1 in rat liver nuclei. J. Biol. Chem. 270, 17257–17263. Jenkins, G.M., Cowart, L.A., Signorelli, P., Pettus, B.J., Chalfant, C.E., and Hannun, Y.A. (2002). Acute activation of de novo sphingolipid biosynthesis upon heat shock causes an accumulation of ceramide and subsequent dephosphorylation of SR proteins. J. Biol. Chem. 277, 42572–42578. Jin, J., Smith, F.D., Stark, C., Wells, C.D., Fawcett, J.P., Kulkarni, S., Metalnikov, P., O’Donnell, P., Taylor, P., Taylor, L., et al. (2004). Proteomic, functional, and domain-based analysis of in vivo 14-3-3 binding proteins involved in cytoskeletal regulation and cellular organization. Curr. Biol. 14, 1436–1450. Jurica, M.S., and Moore, M.J. (2003). Pre-mRNA splicing: awash in a sea of proteins. Mol. Cell 12, 5–14. Kim, Y.M., Watanabe, T., Allen, P.B., Kim, Y.M., Lee, S.J., Greengard, P., Nairn, A.C., and Kwon, Y.G. (2003). PNUTS, a protein phosphatase 1 (PP1) nuclear targeting subunit. Characterization of its PP1- and RNA-binding domains and regulation by phosphorylation. J. Biol. Chem. 278, 13819–13828. Lesage, B., Beullens, M., Nuytten, M., Van Eynde, A., Keppens, S., Himpens, B., and Bollen, M. (2004). Interactor-mediated nuclear translocation and retention of protein phosphatase-1. J. Biol. Chem. 279, 55978–55984. Licatalosi, D.D., and Darnell, R.B. (2006). Splicing regulation in neurologic disease. Neuron 52, 93–101. MacKintosh, C., Beattie, K.A., Klumpp, S., Cohen, P., and Codd, G.A. (1990). Cyanobacterial microcystin-LR is a potent and specific inhibitor of protein phosphatases 1 and 2A from both mammals and higher plants. FEBS Lett. 264, 187–192. Maniatis, T., and Tasic, B. (2002). Alternative pre-mRNA splicing and proteome expansion in metazoans. Nature 418, 236–243. Manley, J.L., and Tacke, R. (1996). SR proteins and splicing control. Genes Dev. 10, 1569–1579.

Prasad, J., Colwill, K., Pawson, T., and Manley, J.L. (1999). The protein kinase Clk/Sty directly modulates SR protein activity: both hyper- and hypophosphorylation inhibit splicing. Mol. Cell. Biol. 19, 6991–7000. Shamovsky, I., Ivannikov, M., Kandel, E.S., Gershon, D., and Nudler, E. (2006). RNA-mediated response to heat shock in mammalian cells. Nature 440, 556–560. Shin, C., and Manley, J.L. (2002). The SR protein SRp38 represses splicing in M phase cells. Cell 111, 407–417. Shin, C., and Manley, J.L. (2004). Cell signalling and the control of pre-mRNA splicing. Nat. Rev. Mol. Cell Biol. 5, 727–738. Shin, C., Feng, Y., and Manley, J.L. (2004). Dephosphorylated SRp38 acts as a splicing repressor in response to heat shock. Nature 427, 553–558. Shin, C., Kleiman, F.E., and Manley, J.L. (2005). Multiple properties of the splicing repressor SRp38 distinguish it from typical SR proteins. Mol. Cell. Biol. 25, 8334–8343. Shi, Y., Reddy, B., and Manley, J.L. (2006). PP1/PP2A phosphatases are required for the second step of pre-mRNA splicing and target specific snRNP proteins. Mol. Cell 23, 819–829. Tamrakar, S., and Ludlow, J.W. (2000). The carboxyl-terminal region of the retinoblastoma protein binds non-competitively to protein phosphatase type 1alpha and inhibits catalytic activity. J. Biol. Chem. 275, 27784–27789. Tazi, J., Kornstadt, U., Rossi, F., Jeanteur, P., Cathala, G., Brunel, C., and Luhrmann, R. (1993). Thiophosphorylation of U1-70K protein inhibits pre-mRNA splicing. Nature 363, 283–286. Trinkle-Mulcahy, L., Sleeman, J.E., and Lamond, A.I. (2001). Dynamic targeting of protein phosphatase 1 within the nuclei of living mammalian cells. J. Cell Sci. 114, 4219–4228.

Margolis, S.S., Walsh, S., Weiser, D.C., Yoshida, M., Shenolikar, S., and Kornbluth, S. (2003). PP1 control of M phase entry exerted through 14-3-3-regulated Cdc25 dephosphorylation. EMBO J. 22, 5734–5745.

Velazquez-Dones, A., Hagopian, J.C., Ma, C.T., Zhong, X.Y., Zhou, H., Ghosh, G., Fu, X.D., and Adams, J.A. (2005). Mass spectrometric and kinetic analysis of ASF/SF2 phosphorylation by SRPK1 and Clk/Sty. J. Biol. Chem. 280, 41761–41768.

Mermoud, J.E., Cohen, P.T., and Lamond, A.I. (1994). Regulation of mammalian spliceosome assembly by a protein phosphorylation mechanism. EMBO J. 13, 5679–5688.

Xiao, S.H., and Manley, J.L. (1997). Phosphorylation of the ASF/SF2 RS domain affects both protein-protein and protein-RNA interactions and is necessary for splicing. Genes Dev. 11, 334–344.

Misteli, T. (1999). RNA splicing: what has phosphorylation got to do with it? Curr. Biol. 9, R198–R200.

Yaffe, M.B. (2002). How do 14-3-3 proteins work?—gatekeeper phosphorylation and the molecular anvil hypothesis. FEBS Lett. 513, 53–57.

Misteli, T., and Spector, D.L. (1996). Serine/threonine phosphatase 1 modulates the subnuclear distribution of pre-mRNA splicing factors. Mol. Biol. Cell 7, 1559–1572.

Zhong, M., Orosz, A., and Wu, C. (1998). Direct sensing of heat and oxidation by Drosophila heat shock transcription factor. Mol. Cell 2, 101–108.

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