Studies on Mammalian Pre-Mrna Splicing: Connections to Transcription and Cancer

Studies on mammalian pre-mRNA splicing: connections to transcription and cancer

Charles J. David

Submitted for the partial fulfillment of the requirements

for the degree of Doctor of Philosophy in the

Graduate School of Arts and Sciences

Columbia University

2011

Studies on mammalian pre-mRNA splicing: connections to transcription and cancer

Charles J. David

This thesis presents two separate pieces of work pertaining to pre-mRNA splicing in mammalian cells. The first part examines the regulation of the alternative splicing of the PKM gene in cancer cells, while the second part investigates the physical connections between the transcriptional apparatus and splicing factors. Cancer cells uniformly alter key aspects of their metabolism, including their glucose usage. In contrast to quiescent cells, which use most of their glucose for oxidative phosphorylation when oxygen is present, under the same conditions, most of the glucose consumed by cancer cells is converted to lactate. This phenomenon is known as aerobic glycolysis, and is critical for cancer cell growth. The pyruvate kinase isoform expressed by the cell is a key determinant of glucose usage. Pyruvate kinase in most tissues is produced from the PKM gene, which is alternatively spliced to produce to produce the PKM1 or PKM2 isoforms, which contain exons 9 or 10 respectively. Adult tissues express predominantly the

PKM1 isoform, which is universally reverted to the embryonic PKM2 isoform in cancer cells.

PKM2 expression promotes aerobic glycolysis. In Chapter 3, I describe a mechanism by which cancer cells promote switching to PKM2. We show that PKM exon 9 is flanked by binding sites

for the RNA-binding proteins hnRNP A1/A2 and PTB. These proteins bind to exon 9 and repress

its inclusion in the mRNA, resulting in PKM2 production. Additionally, we show that hnRNP

A1/A2 and PTB are all overexpressed in cancers in a way that precisely correlates with the

expression of PKM2. Finally, we show that the oncogenic transcription factor c-Myc promotes

PKM2 expression by transcriptionally upregulating the genes encoding hnRNP A1/A2 and PTB.

In the second part of my work, presented in Chapter 5, I examine the coupling of transcription and splicing. The RNA polymerase II C-terminal domain (CTD) plays an important role in ensuring that pre-mRNA transcripts are efficiently spliced, most likely through interactions between splicing factors and the CTD. We have established a biochemical complementation system that has facilitated the identification of a splicing factor that binds to the CTD. Surprisingly, purification of the factor revealed it to be a complex containing U2AF65 and the Prp19 complex, two central splicing factors that had not previously been shown to interact. This complex is functional: I present evidence that the two factors can only activate splicing of the IgMA3 pre-mRNA when they are engaged in a complex. I go on to show that

U2AF65 binds directly to the CTD, and this interaction stimulates the RNA binding of U2AF65.

TABLE OF CONTENTS

Table of Contents ………………………………………………………………………………. i

Preface ...... iv

Acknowledgements …………………………………………………………………………….. v

Dedication ……………………………………………………………………………………… vi

Chapter 1: Alternative pre-mRNA splicing regulation in cancer: pathways and programs unhinged ………………………………………………………………………………... 1

Abstract ………………………………………………………………………………………… 3

Regulation of AS by RNA-binding proteins ……………………………………………………. 4

Splicing and apoptosis ……………………………………………………………………...... 6

Biological functions of apoptosis-regulating RBPs ……………………………………………. 12

AS regulation of metabolism – pyruvate kinase M ……………………………………………..16

Regulation of proto-oncogenes by AS …………………………………………………………. 19

Invasion and metastasis………………………………………………………………………..... 23

Regulation of angiogenesis: VEGFA …………………………………………………………... 30 hnRNP and SR proteins in proliferation and cancer …………………………………………… 33

Conclusions …………………………………………………………………………………….. 39

References ……………………………………………………………………………………… 41

Chapter 2: The search for alternative splicing regulators: new approaches offer a path to a splicing code …………………………………………………………………………… ………72

Known AS regulators …………………………………………………………………………. ..75

A genetic approach to identifying AS regulators ………………………………………………. 79

Three conserved splicing regulators identified for another AS event ………………………….. 83

From the identification of AS regulators to a cellular code of AS regulation …………………. 84

Conclusions and perspectives …………………………………………………………….……. 85

References ……………………………………………………………………………………… 86

Chapter 3: HnRNP proteins controlled by c-Myc deregulate pyruvate kinase mRNA splicing in cancer ……………………………………………………………………… ……... 96

Abstract ……………………………………………………………………………………….... 98

Methods summary ……………………………………………………………………………...105

References ……………………………………………………………………………………...106

Full Methods ………………………………………………………………………………….. 112

Chapter 4: RNA polymerase II and pre-mRNA splicing …………………………………..121

The spliceosome assembly pathway ………………………………………………………… 122

Evidence for coupling of transcription and splicing ………………………………………… 124

The RNAPII CTD: a critical mediator of transcription/splicing coupling …………………….126

CTD structure and modification ……………………………………………………………….127

Physical links between the CTD and splicing …………………………...... 128

References …………………………………………………………………………………… 129

Chapter 5: The RNA polymerase II C-terminal domain promotes splicing activation through recruitment of a U2AF65-Prp19 complex …………………………………………133

Abstract ……………………………………………………………………………………… 135

Introduction …………………………………………………………………………………….136

Results ……………………………………………………………………………………….....139

Discussion …………………………………………………………………………………… 147

Materials and methods …………………………………………………………………………152

References …………………………………………………………………………………… 156

Figure legends ………………………………………………………………………………….163

Figures ………………………………………………………………………………………… 169

iii

Preface

This thesis is divided into five chapters. The Chapters 1 and 2 are review articles on the regulation of alternative splicing written for Genes and Development, provided in their original print format. Chapter 3, about the regulation of PKM alternative splicing is provided in its original form as published in Nature. Chapter 4 is a brief introduction to spliceosome assembly and transcription/splicing coupling. Chapter 5 is a recently submitted manuscript describing the purification of a CTD-dependent splicing activity.

Acknowledgements

I would like to first thank my advisor Jim Manley for years of patient support. I have

learned a great deal from him over the past several years. I also have to thank Jim for introducing

me to a couple of very interesting projects. My committee members Larry Chasin and Carol

Prives have been the source of helpful suggestions, for which I think them. I would also like to thank my outside committee members, Magda Konarska and Kristen Lynch for taking the time to be on my committee.

Members of the Manley lab have been indispensable during my PhD. I learned in vitro splicing and baculovirus expression from Scott Millhouse, who also initiated the project described in Chapter 5. Discussions with Yongsheng Shi, Tsuyoshi Kashima, Jing-ping Hsin were also helpful. I am very indebted to Emanuel Rosonina and Patricia Richard, who spent a

huge amount of time listening to me talk, and teaching me how to present my results in a

professional manner. Tristan Coady and Dafne Campigli Di Giammartino were also very helpful

in this regard.

Finally, I wouldn’t be completing a PhD if it weren’t for members of my family. My dad

passed on a love of science to me. My mother has been extremely supportive, while providing invaluable advice on the route to a PhD. Finally, I owe the biggest debt of gratitude scientifically

and personally to my fiancée, Mo Chen. I would certainly not be completing this without her!

To Mo

Chapter 1: Alternative pre-mRNA splicing regulation

in cancer: pathways and programs unhinged.

Alternative pre-mRNA splicing regulation in cancer: Pathways and programs unhinged

Charles J. David and James L. Manley

Department of Biological Sciences

Columbia University, New York, NY 10027

3 Abstract

Alternative splicing of mRNA precursors is a nearly ubiquitous and extremely flexible point of gene control in humans. It provides cells with the opportunity to create protein isoforms of differing, even opposing, functions from a single gene. Cancer cells often take advantage of this flexibility to produce proteins that promote growth and survival. Many of the isoforms produced in this manner are developmentally regulated, and are preferentially re-expressed in tumors. Emerging insights into this process indicate that pathways that are frequently deregulated in cancer often play important roles in promoting aberrant splicing, which in turn contributes to all aspects of tumor biology.

Each regulatory point in the control of gene expression (which includes chromatin structure, splicing and polyadenylation of mRNA precursors, translation, and mRNA and protein stability), is subject to profound alterations during the development of most, if not all cancers

(Venables 2006; Mayr and Bartel 2009; Chi et al. 2010; Silvera et al. 2010). Of all these points in the gene expression pathway, none provides the potential for more diverse outcomes than alternative splicing (AS). AS, the alternative selection of splice sites present within a pre-mRNA, leads to the production of multiple mRNAs from a single gene. AS thus has the capacity to radically alter the composition and function of the encoded protein. For example, a frequent outcome of AS is the production of proteins with opposing functions, a phenomenon illustrated perhaps most dramatically by the fact that a large majority of genes encoding proteins that function in apoptotic cell death pathways give rise to either pro- or anti-apoptotic isoforms by

AS (Schwerk and Schulze-Osthoff 2005).

The plasticity offered by AS to remodel the proteome means that this process is rich with opportunities for cancer cells to subvert the process to produce proteins that suit the needs of the growing and spreading tumor. All areas of tumor biology appear to be affected by changes in AS, including metabolism, apoptosis, cell cycle control, invasion and metastasis, as well as

4 angiogenesis (Venables 2004; Ghigna et al. 2008). Many of these events appear to represent a return to isoforms normally expressed in a tightly controlled manner during development, but downregulated in most adult cells. Therefore, as in many other areas of tumor biology, the regulation of these AS events in cancer (often by a recurring cast of splicing factors, as will be discussed below) can be understood as a consequence of the deregulation of important developmental pathways.

Genome-wide approaches have revealed that tumorigenesis often involves large scale alterations in AS (Venables et al. 2009). Such approaches have been valuable in providing insight into the regulation of splicing in cancer, and have even proven useful in the classification of tumors (Venables 2006; Skotheim and Nees 2007; Omenn et al. 2010). While the number of

AS events observed to differ in cancer has grown quickly with the use of such methods, relatively few have been demonstrated to be functionally important. Most such events have instead been discovered, often serendipitously, by investigators working in various areas of cancer biology. After a brief discussion of general splicing regulatory strategies, this review will focus on AS events for which a functional significance has been established in processes relevant to cancer biology, and for which the underlying regulatory mechanisms have been investigated.

Several excellent reviews that deal with related topics have been published recently, and the reader is referred to these for additional insight into the importance of AS in development and disease (Skotheim and Nees 2007; Ghigna et al. 2008; Grosso et al. 2008; Cooper et al. 2009).

Regulation of AS by RNA Binding Proteins

Pre-mRNA splicing, the joining of two exons accompanied by the removal of intronic sequence, requires a very large ribonucleoprotein (RNP) complex termed the spliceosome (Wahl et al. 2009). The spliceosome must be guided to the correct splice sites, so what determines which sequences will be included in the mRNA, and which will be excised and degraded in the nucleus? The answer to this question has turned out to be quite complex, especially in light of

5 recent revelations that almost all genes produce transcripts that undergo AS, with a great deal of cell-type variation in what sequences are defined as exons and included in the final mRNA

(Wang et al. 2008a). Most of the information necessary to decide which sections of pre-mRNA will be included when and where is present in the sequence of the pre-mRNA (Barash et al.

2010). This information is “read” by RNA-binding proteins (RBPs) that bind to RNA with varying degrees of sequence specificity, and dictate the fate of the surrounding RNA sequences

(Chen and Manley 2009; Nilsen and Graveley 2010). Alterations in the levels and activity of these RBPs thus provide the primary means of AS regulation.

A number of RBPs that function to control AS have been well studied. The classical regulators of splice site choice are serine/arginine-rich (SR) proteins, which, when bound to exonic sequences known as exonic splicing enhancers (ESEs), tend to promote exon inclusion, and heterogeneous ribonucleoproteins (hnRNPs), which frequently bring about exon exclusion when bound to exonic and/or intronic splicing silencers (E/ISSs; Figure 1). These proteins are joined by several dozen additional RBPs, some with more restricted cell-type expression patterns that play important roles in a more limited number of tissue-specific AS events (for review, see

Chen and Manley, 2009). Many RBPs can act positively or negatively on exon inclusion depending on the location of their binding sites relative to the regulated exon. This principle was demonstrated systematically by Darnell and colleagues, who showed that the brain-specific Nova proteins inhibit exon inclusion when their binding sites are located within the exon, while Nova binding sites in the adjacent intron tend to promote exon inclusion (Ule et al. 2006).

In light of their crucial role in regulating AS, it follows that aberrant expression and regulation of RBPs likely results in the deregulation of splicing observed in cancer. As will be discussed below, the production of critical AS isoforms can be a matter of life or death for cancer cells, meaning that there is strong selection for the expression of certain variants. However, as will also be discussed below, most of the RBPs that regulate these important AS events have pleiotropic effects on splicing and other processes (especially translation), meaning that the

6 critical changes in AS come as part of a wider program of RBP-mediated changes in gene expression. Most of the additional activities of these RBPs are generally consonant with the outcomes of the functionally important splicing events they govern. This indicates that cancer- associated RBPs are normally regulated as part of developmental pathways that are dysregulated at various stages of tumorigenesis. While the regulation of these RBPs is still not well understood, some new insights are available, as will be discussed below.

Splicing and Apoptosis

Apoptosis, or programmed cell death, occurs through activation of one of several pathways present in normal cells. Because cancer cells display behavior that would normally elicit apoptosis, these cells must in some way or another suppress this process (Letai 2008). As mentioned above, transcripts from a significant number of genes involved in apoptosis are alternatively spliced, often resulting in isoforms with opposing roles in promoting or preventing cell death (Schwerk and Schulze-Osthoff 2005). Apoptotic signaling pathways have been shown to alter the balance of some of these isoforms in favor of pro-apoptotic isoforms, indicating that altering the balance of pro/anti-apoptotic isoforms is a normal part of programmed cell death (see below).

Given the apparent importance of particular splicing decisions in regulating apoptosis, it is not surprising that the levels of certain RBPs that control these events are a determinant of whether a cell undergoes apoptotic death; the biology of some proteins implicated in the control apoptotic AS events will be discussed at the end of the section. While the number of genes involved in apoptosis that are alternatively spliced is large (see Schwerk and Schulze-Osthoff,

2005), here we focus on a few well-studied examples for which insight into the relevant splicing regulatory mechanisms is available.

Bcl-x

7 One of the earliest discovered examples of AS creating opposing isoforms in apoptosis is

Bcl-x. The Bcl-x pre-mRNA can be alternatively spliced to produce two isoforms, Bcl-x(L), which has antiapoptotic effects, and Bcl-x(s) which promotes apoptosis (Boise et al. 1993). The two isoforms arise from alternative splicing at two competing 5’ splice sites in exon 2, the first coding exon of the Bcl-x transcript (Figure 2a). Expression of Bcl-x(L), like Bcl-2, was found to prevent cell death after growth factors were removed from the medium (Boise et al. 1993). The shorter isoform antagonized the protective effects of both Bcl-2 and Bcl-x(L), and the expression of Bcl-x(s) alone was sufficient to induce apoptotic cell death in a wide range of cancer cell types (Boise et al. 1993; Clarke et al. 1995; Minn et al. 1996).

High Bcl-x(L)/Bcl-x(s) ratios are observed in a variety of cancer types, consistent with an important role for the long isoform in cancer cell survival (Xerri et al. 1996; Olopade et al. 1997; Takehara et al. 2001). This likely reflects a reduction in Bcl-x(s) in cancer, as an examination of endometrial carcinomas showed a downregulation of Bcl-x(s) mRNA compared to normal endometrial tissue, with the extent of Bcl-x(s) downregulation correlated with clinical staging. (Ma et al. 2010). Illustrating the importance of the this AS event to cancer cells, an antisense oligonucleotide complementary to the Bcl-x(L) isoform 5’ splice site shifted splicing of Bcl-x to the Bcl-x(s) isoform, and was sufficient to induce apoptosis in a prostate cancer cell line (Mercatante et al. 2002). A variety of signals and effectors that regulate Bcl-x AS have been identified. Prompted by reports that Sam68 overexpression can result in apoptosis in NIH 3T3 cells (Babic et al. 2006), and their observation that Sam68 interacts with the Bcl-x mRNA, Sette and colleagues investigated a possible role for this protein in Bcl-x splicing (Paronetto et al. 2007).

Overexpression of Sam68 in 293 cells resulted in an increase Bcl-x(s) isoform, consistent with the pro-apoptotic effects observed upon Sam68 overexpression (Taylor et al. 2004; Paronetto et al. 2007). Interestingly, Sam68 phosphorylation by the Src-like tyrosine kinase Fyn reversed the effects of Sam68 overexpression, switching splicing of Bcl-x back to the long isoform. This result indicates that in the presence of Sam68, growth factors or other signals that activate Fyn,

8 or other tyrosine kinases, are necessary to maintain expression of Bcl-x(L), providing an additional connection between mitogenic signaling pathways and regulation of apoptosis.

While mitogenic signaling pathways have been implicated in maintaining high levels of

Bcl-x(L), a pro-apoptotic pathway initiated by the sphingolipid ceramide has been suggested to promote Bcl-x(s) splicing (Chalfant et al. 2002; Pettus et al. 2002). Ceramide activates the serine/threonine phosphatases PP1 and PP2A, and treatment of cells with an inhibitor of PP1 negated the effects of ceramide on Bcl-x splicing (Chalfant et al. 2002). Ceramide-induced activation of PP1 has been shown to result in widespread dephosphorylation of SR proteins, although no direct connection between this event and Bcl-x splicing has been established

(Chalfant et al. 2001). The RNA-binding protein SAP155, best known as a member of the SF3b complex that associates with the U2 snRNP, has been shown to bind to a ceramide responsive element present in the Bcl-x pre-mRNA and is necessary for the effects of ceramide on Bcl-x splicing (Massiello et al. 2006). Incidentally, SAP155 is a known target of PP1/PP2A prior to the second step of splicing (Shi et al. 2006). It is tempting to speculate that dephosphorylation of

SAP155 by PP1 has a role in regulating Bcl-x splicing. In a separate pathway, expression of the transcription factor E2F1, which can promote apoptosis, resulted in an increase in Bcl-x(s)

(Merdzhanova et al. 2008). Depletion of the SR protein SRSF2 (formerly SC35; Krainer and

Manley, 2010), which is specifically induced by E2F1, reversed this effect, implicating SRSF2 in regulation of Bcl-x splicing.

Caspase-2

Caspase-2 is a highly conserved cysteine protease first identified as a mammalian homolog of the CED-3 caspase in C. elegans (Wang et al. 1994). While it was first implicated in apoptosis on the basis of its similarity to CED-3, it has since been shown to act as a tumor suppressor that participates in a wide variety of cellular processes (Ho et al. 2009; Kumar 2009).

Caspase-2 mRNA is alternatively processed to produce multiple isoforms (Figure 2b). The

9 predominant form in most tissues, caspase-2L, produces a full-length protein with pro-apoptotic properties (Wang et al. 1994). However, in certain differentiated tissues such as brain and skeletal muscle an additional mRNA isoform containing an additional 61 nt exon (exon 9) was detected (Wang et al. 1994). Inclusion of this exon results in a frameshift leading to the introduction of a premature termination codon (PTC) in the mRNA, creating a short-lived nonsense-mediated decay (NMD) substrate (Solier et al. 2005). It is unclear under what circumstances caspase-2S mRNA might be stabilized, and conclusive evidence that it is translated is still lacking (Kitevska et al. 2009). In any event, when expressed from cDNA, the truncated product of the caspase-2S mRNA was shown to protect against cell death in some contexts (Wang et al. 1994; Droin et al. 2001).

While the existence of the caspase-2S protein remains to be conclusively demonstrated

(Kitevska et al. 2009), alignment of caspase-2 gene sequences from multiple organisms reveals that E9 and the sequences flanking it are in fact highly conserved throughout vertebrates (in fact,

E9 is among the most highly conserved portions of the gene; Figure 3a). This leaves little doubt that E9 inclusion plays an important role at some point in development. One clue to a potential function of E9 inclusion comes from a recent study that identified a class of AS event in which increased inclusion in differentiated cells of a PTC-inducing exon acts as a means of post- transcriptional gene control, resulting in NMD-mediated downregulation of the gene product

(Barash et al. 2010). Caspase-2 expression has been shown to be reduced in the developing retina, where the 2L isoform is downregulated with a concomitant increase in a small amount of detectable 2S isoform, indicating that E9 inclusion may be a mechanism for developmental control (Kojima et al. 1998).

Whether exon 9 inclusion in caspase-2 results in the production of an anti-apoptotic protein or simply results in reduction of caspase-2 mRNA levels through NMD, it is clear that production of the 2S isoform would favor survival of cancer cells, making its regulation of interest. Early experiments showed that overexpression of hnRNP A1 promoted inclusion of E9

10 in a minigene construct, while overexpression of SRSF2 resulted in the opposite effect (Jiang et al. 1998). The RNA cis-elements responsible for the effects of hnRNP A1 and SRSF2 were not identified, but an element in intron 9 (I9) termed In100 was shown to contain binding sites for another hnRNP protein, polypyrimidine tract binding protein (PTB), from which it repressed E9 inclusion (Cote et al. 2001b). In100 also contains a “decoy” 3’ splice site, capable of forming non-productive spliceosome-like complexes with the E9 5’ splice site (Cote et al. 2001a; Cote et al. 2001b). E9 inclusion is promoted by the RBP RBM5. RBM5 binds to an element in I9 to promote E9 exclusion, producing the full-length pro-apoptotic isoform of caspase-2 (Fushimi et al. 2008).

In addition to RNA binding proteins, promoter choice appears to affect caspase-2 splicing.

Corcos and colleagues demonstrated that caspase-2S and caspase-2L mRNAs derive from different transcription start sites and contain different 5’ untranslated exons (Logette et al. 2003).

It is known that alternative promoters can have a profound impact on AS patterns (Kornblihtt

2005), and caspase-2 splicing provides another example of this phenomenon. The importance of this phenomenon in regulating caspase-2 splicing in cancer, as well as the mechanism of promoter-dependent differential E9 inclusion, have yet to be explored.

Fas

Alternative splicing of the Fas receptor pre-mRNA provides a potentially important means by which tumors cells can escape elimination by the immune system. The Fas protein

(also known as CD95) is a widely expressed cell-surface receptor, which when bound to Fas ligand (FasL) expressed on cytotoxic T-cells can initiate a cascade that eventually leads to cell death (Bouillet and O'Reilly 2009). In addition to producing the full-length mRNA, the Fas pre- mRNA can be alternatively spliced to produce a number of shorter products (Figure 2c). The most abundant of these is an isoform in which the 63 nt exon 6 (E6) is skipped, deleting the transmembrane domain (Cheng et al. 1994; Cascino et al. 1995). The protein produced by the

11 ΔE6 Fas isoform is soluble, and capable of inhibiting Fas-mediated cell death, presumably by binding to FasL and preventing the interaction of FasL with membrane-bound Fas.

Elevated production of soluble Fas (sFas) has been observed in a wide range of cancers, as determined by Fas serum concentrations, which show a strong correlation with tumor staging

(e.g., Sheen-Chen et al. 2003; Kondera-Anasz et al. 2005). Consistent with a role for AS in producing the soluble variants, examination of Fas mRNA isoforms in peripheral blood mononuclear cells (PBMCs) from patients with large granular lymphocyte leukemia revealed a large increase in the ΔE6 isoform compared with PBMCs from healthy individuals, consistent with increased serum concentrations of sFas in these patients (Liu et al. 2002). Studies examining the expression of Fas mRNA isoforms in other forms of cancer will be important to provide support for the idea that changes in Fas AS (as opposed to mechanisms such as proteolytic cleavage) indeed underlie the widespread appearance of sFas in cancer.

In light of the potential for Fas AS to play a role in the suppression of the anti-tumor immune response, the regulation of Fas AS has been extensively investigated. Several RBPs have been shown to be involved in promoting the production of full-length Fas mRNA. TIA-1 and TIAR, two closely related RNA recognition motif (RRM)-containing proteins involved in apoptosis (see below), bind to U-rich sequences downstream of Fas E6 and promote its inclusion in the mRNA (Izquierdo et al. 2005). TIA-1 binding downstream of E6 results in increased U1 snRNP recruitment, presumably through an interaction with the U1 snRNP protein U1C (Forch et al. 2002; Izquierdo et al. 2005). Interestingly, Fas receptor activation influences the splicing of its own pre-mRNA, through the activation of the Fas-associated S/T kinase (FAST K). FAST K can phosphorylate TIA-1/TIAR, potentiating their ability to activate E6 inclusion by increasing their ability to recruit U1 snRNP to the pre-mRNA (Figure 4; Izquierdo and Valcarcel 2007).

A number of repressors of E6 inclusion have also been identified. Valcarcel and colleagues found that PTB inhibits E6 inclusion by interfering with binding of the general splicing factor U2AF to the polypyrimidine tract upstream of E6 (Izquierdo et al. 2005). RBM5

12 was also identified as an inhibitor of Fas E6 inclusion. RBM5, unlike PTB, does not disrupt early events in the recognition of E6 by the splicing machinery, but rather inhibits the pairing between spliceosomal complexes assembled on E6 and those on the neighboring exons (Bonnal et al.

2008). Another RBP that promotes the expression of the sFas isoform is HuR, which binds to inhibitory sequences in E6 to promote its exclusion (Izquierdo 2008).

Biological functions of apoptosis-regulating RNA binding proteins

As just discussed, a number of RBPs that affect the splicing of apoptosis-related transcripts have been identified, highlighting the possibility that these proteins are in fact regulators of apoptosis. Below we discuss the function and biology of some of these proteins in normal development and cancer.

TIA-1/TIAR

T-cell intracellular antigen-1 (TIA-1) and TIA-1-related protein (TIAR) are highly conserved and widely expressed in mammals (Beck et al. 1996). The two proteins function in similar processes, and often exhibit partial redundancy (e.g., in Fas splicing as mentioned above).

However, they are not fully redundant, as knockout of either results in embryonic lethality (Beck et al. 1998; Piecyk et al. 2000). In addition, TIAR appears to have a unique role in germ cell development (Beck et al. 1998). Among the earliest functions attributed to TIA-1 and TIAR was the ability to bind mRNAs and recruit them to cytoplasmic bodies known as stress granules in response to cellular stress such as heat shock (Kedersha et al. 1999). Both proteins have also been found to bind AU-rich elements (AREs) and act as translational repressors (Piecyk et al.

2000; Kawai et al. 2006; Kim et al. 2007) TIA-1 and TIAR have been shown to be regulators of the inflammatory response that function by silencing translation of key mediators of inflammation such as TNF-α and COX-2 (Piecyk et al. 2000; Lopez de Silanes et al. 2005a).

13 Following the realization that TIA-1/TIAR show similarity to the S. cerevisiae U1 snRNP associated protein Nam8, it was demonstrated that the two proteins can function as regulators of pre-mRNA splicing (Gottschalk et al. 1998; Del Gatto-Konczak et al. 2000; Forch et al. 2000).

Notable splicing substrates for TIA-1/TIAR are Fas (discussed above), and the epithelial-specific exon IIIb in the fibroblast growth factor receptor 2 (FGFR2) pre-mRNA, which is frequently excluded during cancer progression (Del Gatto-Konczak et al., 2000; see below). Consistent with the fact that TIA-1/TIAR can promote production of the pro-apoptotic form of Fas, introduction of TIA-1/TIAR into cells promotes apoptosis (Tian et al. 1991; Iseni et al. 2002). In addition to their pro-apoptotic effects, TIA-1/TIAR depletion in HeLa cells was shown to result in increased proliferation (Reyes et al. 2009).

Given that TIA-1 and TIAR have functions associated with tumor-suppressor genes (i.e. promoting apoptosis and inhibiting proliferation), alterations in the expression or regulation of these proteins might be expected to contribute to tumorigenesis in some contexts. However, there is currently little evidence for widespread downregulation of TIA-1/TIAR in cancer. That being said, TIA-1 expression in tumors correlates strongly with responsiveness to immunotherapy for melanoma patients (Wang et al. 2002). This result supports the idea that downregulation of TIA-

1 is indeed a means by which tumors can evade the immune system. In light of its role in processes important to the growth and survival of cancer cells, an examination of changes in

TIA-1/TIAR expression and regulation during tumorigenesis is warranted.

RBM5

RBM5 (RNA binding motif 5) is a putative tumor suppressor that contains a large number of motifs, including a pair of RRMs, two Zn-finger motifs, an RS domain, and an

OCtamer REpeat (OCRE) motif, a newly identified domain now implicated in splicing regulation (Bonnal et al. 2008). RBM5, along with the less extensively studied proteins RBM6

14 and RBM10, comprise a small family of highly similar RNA binding proteins that in at least some activities can function redundantly (Bonnal et al. 2008).

Evidence exists supporting the idea that RBM5 is a tumor suppressor (Sutherland et al.

2010). An early suggestion for this came from the presence of the RBM5 gene on a piece of chromosome 3 (3p21.3) that is frequently deleted in lung cancer. Loss of heterozygosity at this locus occurs in 95% of small cell lung cancer (SCLC), as well as 70% of non-SCLC (Sutherland et al. 2010). Consistent with a role as a tumor suppressor, RBM5 (also known as LUCA-15 and

H37) was later shown to be downregulated in a high proportion of lung cancers (Oh et al. 2002).

RBM5 was also one of 9 genes downregulated as part of a “metastatic signature” identified by microarray (Ramaswamy et al. 2003). In stark contrast to observations that RBM5 functions as a tumor suppressor, RBM5 is consistently overexpressed in breast cancer (Rintala-Maki et al.

2007). Together, these results suggest an important, but likely complex, role for RBM5 in regulating genes important in several cancers.

Given the association with cancer, what are the biological functions of RBM5? A 326 bp fragment of cDNA from the RBM5 locus was identified as a suppressor of Fas-mediated apoptosis in a cDNA screen (Sutherland et al. 2000). It was soon realized that this encoded an inhibitory RNA complementary to the 3’ UTR of the RBM5 mRNA, and that full-length, sense- oriented RBM5 actually potentiates apoptosis initiated by Fas (Rintala-Maki and Sutherland

2004). In addition to its role in apoptosis, expression RBM5 has been found to inhibit proliferation when transfected into a number of different cell lines (Edamatsu et al. 2000; Oh et al. 2006). RBM5 also appears to stimulate p53 transcription and to promote higher levels of p53 transcripts through an unknown mechanism (Kobayashi et al. 2010).

While the results described above all support an important role for RBM5 in apoptosis and cell cycle regulation, apart from its role in Fas and caspase-2 splicing, very little is known about its biochemical functions. Judging from its large number of domains, these are no doubt diverse. In addition, while the effect of RBM5 on caspase-2 splicing (promoting the 2L isoform)

15 appears to be in line with its role in promoting apoptosis, the inhibition of Fas E6 inclusion would be predicted to protect against Fas-mediated apoptosis, which runs counter to its demonstrated role in promoting Fas-mediated apoptosis. Unraveling this apparent paradox, which may underlie the differential regulation of RBM5 in distinct cancer types, as well as identifying additional targets and functions of RBM5 (and of RBM6 and RBM10), will be of considerable future interest.

HuR

HuR is another multifunctional RRM-containing protein that is frequently upregulated in cancer (Blaxall et al. 2000; Lopez de Silanes et al. 2003; Denkert et al. 2004). HuR (unlike the related proteins HuB, HuC, and HuD) is ubiquitously expressed, and shuttles between the nucleus and the cytoplasm. In synchronized cells, the presence of HuR in the cytoplasm is cell- cycle dependent, while a fraction of HuR is constitutively nuclear (Wang et al. 2000). HuR has a relatively well-characterized role in the cytoplasm where it binds to mRNAs containing AREs and stabilizes, as well as affects translation of, a large number of mRNAs relevant to proliferation and apoptosis (Lopez de Silanes et al. 2005b; Hinman and Lou 2008). HuR is subject to regulation by phosphorylation as a result of a number of different signaling pathways, which result in changes in its shuttling ability, as well as its RNA binding (Doller et al. 2008).

HuR promotes cell proliferation, and is one of only a few RBPs for which an oncogenic function has been demonstrated. This was shown in a study in which HuR-overexpressing RKO colon cancer cells, when injected into nude mice, produced much larger tumors than control cells

(Lopez de Silanes et al. 2003).

As mentioned above, HuR functions as a suppressor of apoptosis in cancer cells. HuR knockdown in HeLa cells resulted in apoptosis, while HuR overexpression promoted survival of

UV-irradiated HeLa cells (Lal et al. 2005). HuR binds the mRNA encoding prothymosin alpha, an inhibitor of apoptosis, as well as that of Bcl-2, stabilizing both transcripts (Lal et al. 2005;

16 Ishimaru et al. 2009). While the nuclear role for HuR is much more poorly understood than its cytoplasmic function, a role for HuR in regulating splicing and polyadenylation is beginning to emerge (Hinman and Lou 2008). The identification of Fas exon 6 as a target of HuR repression is in general agreement with observations that HuR is a repressor of apoptosis (Izquierdo 2008).

This finding may portend a more widespread role for HuR in the regulation of post- transcriptional processing of genes involved in apoptosis and proliferation.

AS regulation of metabolism – Pyruvate Kinase M

AS also plays an important role in the control of metabolism in cancer, through the regulation of a key metabolic gene, pyruvate kinase M. One of the earliest observations of the molecular differences between cancer cells and non-cancerous tissue was that cancer cells consume large amounts of glucose and produce prodigious amounts of lactate, even in the presence of oxygen, a process referred to as aerobic glycolysis (also known as the Warburg effect, named for its discoverer) (Warburg 1956). This effect was later shown to be shared by non-cancerous cells induced to proliferate (Wang et al. 1976). From an energy production standpoint, the use of glucose in proliferating cells appears wasteful, as glycolytic conversion of glucose to lactate produces only two molecules of ATP per glucose, while oxidative phosphorylation is capable of producing 36 molecules of ATP per glucose molecule (Vander

Heiden et al. 2009). Why do tumor cells forsake the efficient extraction of energy from glucose instead opting for high glucose consumption and high glycolytic flux? One explanation is that this allows proliferating cells to use carbon derived from glucose for biosynthetic processes necessary for proliferation (Mazurek et al. 2005; Jones and Thompson 2009; Vander Heiden et al.

2009). Multiple glycolytic intermediates represent precursors for the production of nucleotides, lipids, and amino acids, so high glycolytic flux may mean an increased supply of intermediates necessary for growth.

17 How do proliferating cells reprogram their metabolism to engage in aerobic glycolysis?

The Warburg effect requires the shunting of pyruvate from the mitochondria, where, in differentiated cells, it provides the substrate for oxidative phosphorylation, to cytoplasmic conversion to lactate, catalyzed by lactate dehydrogenase. One important part of this process is increased production of lactate dehydrogenase-A (LDH-A), necessary for the final step in aerobic glycolysis, the production of lactate from pyruvate (Fantin et al. 2006). The oncogenic transcription factor c-Myc is known to promote the upregulation of LDH-A, as well as of several other glycolytic enzymes, an activity that appears to underlie, in part, the effect of Myc on aerobic glycolysis (Shim et al. 1997; Kim et al. 2004; Dang et al. 2009).

AS of the pre-mRNA encoding the enzyme that produces pyruvate, pyruvate kinase, is also an important determinant of how glucose is used in cancerous versus differentiated cells.

Pyruvate kinase M (PKM) is the pyruvate kinase gene expressed in all mammalian tissues except liver and erythrocytes (Mazurek et al. 2005). It is subject to mutually exclusive AS, with exon 9

(E9) or exon 10 (E10) included to produce either the adult isoform, PKM1 (E9) or the embryonic version, PKM2 (E10; Figure 2d). Cantley and colleagues found that in tumors, which uniformly express PKM2, replacing PKM2 with PKM1 reduced the production of lactate and increased oxidative phosphorylation, implicating the PKM2 isoform as a promoter of the Warburg effect

(Christofk et al. 2008a). Importantly, and consistent with the idea that aerobic glycolysis is vital for cell growth, replacing PKM2 with PKM1 in cancer cells resulted in impaired growth and reduced ability to form tumors when injected into nude mice. An additional property specific to

PKM2 is its ability to bind phosphotyrosine residues, which frequently result from the activation of mitogenic signaling cascades. This results in the release of its allosteric activator, fructose 1-6 bisphosphate, and the transient inhibition of PK acitivty, which is proposed to promote accumulation of glycolytic intermediates which can then be used for biosynthetic processes

(Christofk et al. 2008b).

18 The universal switching of tumors to PKM2, along with its functional importance to tumor cells, makes the regulation of this AS event of great interest, because it likely reflects an alteration in the splicing regulatory machinery shared by all proliferating cells. Three hnRNP proteins that often act as splicing repressors, PTB, hnRNP A1 and hnRNP A2, were recently shown to promote the formation of the PKM2 isoform by binding to sequences upstream and downstream of E9 (David et al. 2010). SiRNA-mediated depletion of these proteins in a variety of cancer cells resulted in switching of the PKM splicing pattern to the production of PKM1, indicating that in the absence of the repressive hnRNP proteins, the derepressed E9 is able to outcompete E10 for splicing to E11, resulting in the default production of PKM1 (Clower et al.

2010). Supporting the idea that these three RNA binding proteins play an important role in the production of PKM2 in tumors, upregulation of all three proteins correlated perfectly with PKM2 expression in a panel of human gliomas (David et al. 2010). Consistent with the effect on PKM splicing, depletion of hnRNP A1/A2 or PTB also resulted in decreased lactate production in a glioblastoma cell line (Clower et al. 2010). The reduction in lactate production appeared greater than would be expected based solely on the effect of knockdown on PKM splicing, indicating that these proteins likely have additional targets relevant to aerobic glycolysis. One possibility is that knockdown of hnRNP A1/A2 and PTB results in impaired translation of c-Myc, a process promoted by both hnRNP A1 and PTB (Mitchell et al. 2005; Jo et al. 2008).

PKM2 expression appears to be universal in tumors, and likewise the upregulation of hnRNP A1/A2 and PTB is very widely observed in cancer (see below), suggesting that pathways shared by many tumor types promote the overexpression of these RBPs. ChIP-seq data revealed binding of c-Myc to the promoters of all three genes (Chen et al. 2008), suggesting that c-Myc may activate their transcription. Indeed, shRNA-mediated knockdown of c-Myc in NIH-3T3 cells resulted in decreased levels of hnRNP A1/A2 and PTB, and switched AS to favor accumulation of the PKM1 isoform (David et al. 2010). These data suggest that in addition to its role in upregulating glycolytic enzymes, c-Myc contributes to the Warburg effect by indirectly

19 regulating PKM splicing (Chen et al. 2010). However, the effect of c-Myc knockdown on hnRNP protein levels was not observed in all cells tested (David et al. 2010), meaning that additional proliferation- associated transcriptional pathways may play a larger role in inducing overexpression of these hnRNP A1/A2 and PTB in other contexts.

Regulation of proto-oncogenes by AS

Activation of proto-oncogenes is a classic initiating event in cancer. While this often happens through mutation, it is now clear that changes in AS that are unconnected to mutations can profoundly affect the activity of some oncogenes. Here we will discuss two examples of proto-oncogenes for which AS yields products with distinct functions.

Cyclin D1

Cyclin D1 was first identified as a regulator of cell cycle progression through its association with cyclin-dependent kinases 4 or 6 (CDK4/6) (reviewed by Knudsen et al. 2006).

One role of the CDK4/6/cyclin D1 complex is to phosphorylate the transcription factor and tumor suppressor RB, thus relieving RB repression of E2F target genes and promoting cell cycle progression. In addition to its association with CDKs, cyclin D1 was later shown to have CDK- independent nuclear functions as a transcriptional regulator (Knudsen et al. 2006).

It is now known that cyclin D1 is subject to alternative splicing/polyadenylation that results in a more oncogenic protein. Cyclin D1a, the more common full-length variant, contains

5 exons, whereas a variant isoform known as cyclin D1b is polyadenylated at a site in intron 4

(Figure 2e; Betticher et al. 1995). Cyclin D1b can be detected in non-cancerous cells, but is upregulated in some cancers such as breast and prostate (Burd et al. 2006; Wang et al. 2008b).

The D1b isoform, like D1a, associates with CDK4, and appears to regulate CDK activity similarly to D1a (Lu et al. 2003). Instead, a major difference between the D1a and D1b is that while D1a shuttles between the nucleus and cytoplasm in a cell-cycle dependent manner, cyclin

20 D1b is constitutively nuclear, likely due to loss of a glycogen synthase kinase 3β phosphorylation site present at the C-terminus of D1a (Lu et al. 2003; Solomon et al. 2003).

Phosphorylation at this site results in export from the nucleus and proteolytic degradation, while mutation of this residue results in constitutive nuclear localization and a more oncogenic protein

(Alt et al. 2000). Consistent with the greater transforming potential of the constitutively nuclear

D1a mutant, D1b transforms NIH-3T3 cells with much greater efficiency than does D1a

(Solomon et al. 2003). D1b has been shown to associate with the androgen receptor (AR), but unlike D1a, D1b fails to inhibit AR transcriptional activity, a phenomenon of particular relevance to prostate cancer, which shows frequent upregulation of cyclin D1b (Burd et al. 2006;

Comstock et al. 2009).

The choice between D1a and D1b production, unlike the other examples described thus far, represents a competition between the splicing of intron 4 and use of a polyadenylation site within the intron. Some insights into factors affecting this alternative processing event are available. A common polymorphism that occurs near the 5’ splice site of exon 4 influences the extent of cyclin D1b produced. The G870A polymorphism, with a reported allele frequency of

42% in one European population, has been associated with production of D1b (Betticher et al.

1995; Comstock et al. 2009). The polymorphism occurs at the very last nucleotide of exon 4, and may affect the recognition of exon 4 by the splicing machinery. A simple model for how G870A favors D1b production is that the G to A alteration results in impaired recognition of E4, resulting in slower kinetics of intron 4 splicing, thus favoring polyadenylation at the intronic site, resulting in production of D1b. Significantly, the G870A allele, which does not alter the encoded protein, is associated with increased risks for multiple cancers, supporting a role for D1b in tumorigenesis (Knudsen et al. 2006).

While the G870A polymorphism appears to affect the production of D1b in normal tissues, in cancer cells production of D1b occurs regardless of the identity of the final E4 nucleotide (Olshavsky et al. 2010). Consistent with this, RBPs that influence the choice between

21 D1a/D1b have also been identified. Sam68, described above for its role in Bcl-x splicing, has been implicated in favoring the production of D1b (Paronetto et al. 2010). Binding sites for

Sam68 were identified in intron 4, where it either inhibits E4 recognition or promotes polyadenylation in intron 4. A strong correlation between Sam68 expression levels and D1b production was also observed, underscoring the biological significance of these findings. The SR protein SRSF1 (formerly ASF/SF2) has also been shown to bind preferentially to the D1b transcript and promote the production of D1b, and SRSF1 levels were also found to correlate with D1b production (Olshavsky et al. 2010). The effects of SRSF1 on D1b production were enhanced with the 870G allele, potentially providing an explanation for the fact that in cancer both alleles result in similar D1b production. Finally, it has also been reported that an oncogenic fusion between the multifunctional RBP EWS and the transcription factor FLI1, which is found in Ewing’s sarcoma, can influence production of D1b. While EWS, itself, was shown to promote production of full-length D1a, the EWS-FLI1 fusion promoted production of D1b (Sanchez et al.

2008). This was proposed to occur due to impaired transcription elongation in the presence of the fusion protein, which would allow more time for polyadenylation to occur in intron 4 before transcription of E5.

H-Ras

The H-Ras gene appears to provide another example of an AS event that produces two proteins with entirely different activities with regard to proliferation. In the late 1980s Levinson and colleagues identified an intronic mutation in H-Ras that significantly enhanced H-Ras expression, significantly increasing the transforming potential of the gene (Cohen and Levinson

1988). The mutation was soon found to disrupt the 5’ splice site of a previously unknown alternatively spliced exon which the authors named IDX (Figure 2f; Cohen et al. 1989). The IDX exon and flanking intronic regions are highly conserved in mammals (Figure 3b), a strong

22 indication that IDX inclusion is a functionally important regulated AS event (Cohen et al. 1989;

Sorek and Ast 2003).

What is the function of IDX inclusion? The fact that mutation of the IDX splice site results in greatly increased production of full-length H-Ras mRNA led to the initial proposal that one function of the IDX-containing isoform could be to modulate levels of H-Ras production.

When included, an in-frame stop codon in IDX (the third-to-last exon if all downstream introns are removed, see below) results in a presumptive NMD product, which was first proposed to channel much of the H-Ras transcript into a degradation pathway (Cohen et al. 1989). This was confirmed by a later study (Barbier et al. 2007).

While IDX-containing H-Ras (p19) appears to be an NMD target, Guil et al, using antibodies to the divergent C-terminus of p19 H-Ras, found that p19 is indeed expressed in HeLa cells (Guil et al. 2003a). Specific knockdown of p19 H-Ras resulted in increased proliferation of multiple cell lines (Guil et al. 2003a; Jang et al. 2010). p19, in contrast with other Ras proteins, is strongly localized in the nucleus, and was found to bind to p53 and p73, and to promote transcription by the p73 isoform p73β, possibly by preventing an inhibitory interaction between

MDM2 and p73 (Jeong et al. 2006).

Because the functions of p19 H-Ras apparently differ greatly from full-length H-Ras, the regulation of IDX inclusion is likely important at some point in development, an idea reinforced by the fact that the IDX exon and its flanking introns are highly conserved in mammals (Figure

3b). A number of RBPs have been implicated in the control of IDX inclusion. Using an in vitro splicing assay, Guil et al. (2003b) identified an ISS downstream of IDX that was found to bind hnRNP A1, hnRNP H, and the RNA helicase p68 (Guil et al. 2003b). HnRNP A1 and p68 were both found to negatively regulate IDX splicing, while hnRNP H was shown to promote the production of IDX-containing mRNA. Sequences downstream of IDX are predicted to form an extensive stem-loop structure, leading to speculation that p68 helicase activity plays a role in negatively regulating IDX inclusion, which was proposed to interfere with binding of hnRNP H

23 to that region (Camats et al. 2008). In addition to investigating the contributions of the hnRNP proteins and helicase, these authors also provided evidence, using in vitro assays and transient transfections, that the SR proteins SRSF2 and SRSF5 (SRp40) can also function in promoting

IDX splicing.

Invasion and metastasis

AS regulates a number of genes that play important roles in promoting invasive behavior.

In addition, a process that often plays a role in the acquisition of invasive behavior in cancer cells, the epithelial to mesenchymal transition (EMT), is accompanied by a reprogramming of AS

(Thiery et al. 2009). Downregulation of a pair of epithelial-specific splicing factors may underlie many of the changes in splicing that occur during EMT (see below). In the following paragraphs, we discuss several genes in which alternative splicing to creates isoforms that are associated with metastasis and the acquisition of invasive properties.

CD44

The transmembrane protein CD44 was among the first genes for which specific splice variants were associated with metastasis. In a 1991 report, Herrlich and colleagues demonstrated that CD44 molecules containing variant exons v4-7 and CD44 v6-7 were expressed specifically in a metastasizing pancreatic carcinoma cell line, but not in the parental tumor (Gunthert et al.

1991). Indeed, the expression of these variants in cells derived from the parental tumor was sufficient to render them metastatic. CD44 pre-mRNA is subject to complex AS involving 10 adjacent variant exons that can be included singly or in combination (Figure 2g; Ponta et al.

2003). Later analyses demonstrated the presence of variant exon-containing CD44 molecules throughout adult mammals in a variety of contexts. For example, the variant exons 8-10 are frequently included in CD44 expressed in epithelial tissues (Galiana-Arnoux et al. 2003), while

24 the variant exon 6-containing CD44 first observed to be expressed on metastatic cells was shown to be transiently expressed on B and T lymphocytes after antigenic stimulation, an event necessary for lymphocyte activation (Arch et al. 1992). In an insight into the metastatic properties of CD44 v6, Orian-Rousseau et al. (2002) showed that this molecule is required for the assembly on cell membranes of a ternary complex containing the receptor tyrosine kinase

Met, its ligand hepatocyte growth factor (HGF), and CD44 v6 (Orian-Rousseau et al. 2002).

Formation of this complex is necessary for the activation of Met by HGF, an event that is strongly implicated in the acquisition of metastatic properties by cancer cells (Cecchi et al. 2010).

Additionally, siRNA-mediated knockdown of v5-containing CD44 in HeLa cells resulted in drastically reduced invasion in an in vitro assay, confirming the importance of v5 inclusion in metastatic behavior (Cheng and Sharp 2006).

Because of its long-known association with metastasis, the regulatory mechanisms that underlie the changes in CD44 splicing in cancer have been the subject of long-standing interest, particularly centering around variant exons most strongly associated with metastasis (v4-v7).

CD44 was among the first genes for which AS was shown to be altered by signaling pathways associated with growth. For example, inclusion of both exon v5 and v6 can be induced by Ras signaling (Konig et al. 1998; Weg-Remers et al. 2001; Cheng et al. 2006). Activation of v5 inclusion by Ras was shown to require the Ras-Raf-MEK-ERK pathway, and it was later demonstrated that ERK directly phosphorylates Sam68, which was shown to bind to v5 (Weg-

Remers et al. 2001; Matter et al. 2002). ERK phosphorylation was shown to be required for

Sam68 to activate v5 inclusion (Matter et al. 2002), a finding that for the first time showed a direct connection between a mitogenic signaling pathway and splicing control.

Fibroblast growth factor receptors

Splicing of transcripts of fibroblast growth factor receptor (FGFR) genes, which encode four closely related receptor tyrosine kinases, is closely connected to EMT, and is frequently observed to change during the course of tumorigenesis. The FGFR genes are highly conserved,

25 and FGFR1-3 share a tightly regulated mutually exclusive AS event that determines the composition of the third extracellular Ig domain (Turner and Grose 2010). For FGFR2, this event, involving the choice between exons IIIb and IIIc, is controlled in an exquisitely cell-type specific manner, with the IIIb exon included in epithelial cells and the IIIc exon included in mesenchymal cells, with nearly complete switching occurring during EMT (Figure 2h; Yan et al. 1993). EMT, as discussed above, is often a critical event during tumorigenesis, and likewise, switching between the FGFR2 IIIb/IIIc isoforms accompanies crucial changes in tumor behavior. One example of this occurs in prostate cancer (PCa) cells, which in early stages are dependent on androgen (androgen sensitive), and can be controlled by therapies that reduce the level of circulating androgen. However, in most cases the cancer will return in a form that is no longer sensitive to such androgen deprivation (androgen insensitive). Using a rat prostate cancer model that recapitulates the transition from androgen sensitive to androgen insensitive PCa, Garcia-

Blanco and colleagues showed that androgen sensitive tumors express almost exclusively the

FGFR2 IIIb isoform, while androgen insensitive cells exhibit complete switching to the IIIc isoform (Carstens et al. 1997).

The switch-like nature of this AS event implies an important function in the two different cell types. The mutually exclusive exons compose part of the ligand binding domain, and one functional consequence of AS is a change in ligand affinity. For example the FGFR2 IIIb isoform has high affinity for FGF7, which is secreted by stromal cells, while FGFR2 IIIc does not (Yan et al. 1993). Because of the role of FGF7 in mediating both proliferation and differentiation of FGFR2 IIIb expressing cells, it was proposed that loss of responsiveness to this ligand may be an important event in the development of androgen insensitive PCa (Carstens et al.

1997). Reintroduction of FGFR2 IIIb into a highly malignant PCa cell line reduced tumor formation and resulted in increased differentiation and apoptosis, suggesting that the switch to

FGFR2 IIIc may be important for maximal tumorigenicity (Yasumoto et al. 2004). However, a

26 direct comparison of effects of the two isoforms in cancer must be performed before any conclusions can be made about the functional importance of FGFR2 AS in tumor progression.

While much remains to be learned about the functions of FGFR IIIb/c splicing in development and cancer, a great deal is known about how the switch is regulated, which in turn is instructive of changes in splicing that occur during EMT. Two hnRNP proteins frequently overexpressed in cancer, hnRNP A1 and PTB, have been implicated in silencing of the epithelial-specific IIIb exon, while hnRNP H/F proteins can contribute to IIIc silencing (Del

Gatto-Konczak et al. 1999; Carstens et al. 2000; Mauger et al. 2008). There is currently no strong evidence to suggest that changes in the levels of these hnRNP proteins provides an important means of regulating switching from exon IIIb to IIIc. However, differences in recruitment of these proteins to genes offers an additional, and until recently, unexplored potential point of regulation. In a interesting report, Misteli and colleagues showed that differences in H3K36 tri- methylation can result in differential recruitment of PTB to genes through the histone H3 lysine

36 tri-methyl binding protein MRG15 (Luco et al. 2010). ChIP assays revealed that in mesenchymal stem cells that repress the IIIb exon, H3K36 tri-methylation is increased when compared to epithelial cells in which IIIb is included, a phenomenon proposed to result in increased recruitment of PTB to FGFR2 IIIb in mesenchymal cells.

While the proteins discussed above may play a role in FGFR2 AS regulation, the most important regulators of the IIIb/IIIc AS during EMT appear to be a pair of recently discovered cell type-specific RBPs, ESRP1 and ESRP2 (Warzecha et al. 2009). These two RRM-containing proteins, identified in a cDNA screen for splicing factors that promote the epithelial isoform of

FGFR2, are expressed exclusively in epithelial-type cells. The two proteins function redundantly, and simultaneous knockdown of the two in an epithelial cell line promoted a complete reversal of FGFR2 splicing to exon IIIc, while transfection of ESRP1 or ESRP2 expression vectors into cells that express the IIIc exon had the opposite effect. The complete nature of the IIIb/IIIc switching induced by ESRP1/2 knockdown in an epithelial cell line (from 95% IIIb to 96% IIIc),

27 along with the perfect correlation of ESRP1/2 expression and IIIb inclusion, indicates that expression of these RBPs is the primary determinant of FGFR2 IIIb/IIIc inclusion.

To examine FGFR2 splicing in vivo, Garcia-Blanco and colleagues developed a minigene reporter system designed to express RFP in cells in which exon IIIc is not included (Oltean et al.

2006). They injected mesenchymal AT3 prostate tumor cells expressing the minigene into mice, and found that after invading the lungs, the tumor cells frequently exhibited IIIc skipping, indicative of a mesenchymal to epithelial transition (MET), which was confirmed by the presence of the epithelial marker E-cadherin. This result indicates that in addition to EMT, the reverse transition MET also occurs during tumor progression, and this process entails a reversion to epithelial splicing patterns. While it remains to be examined directly, much of the altered AS that occurs during MET is likely the result of re-expression of ESRP1/2. FGFR1 is subject to another AS event that is of possible relevance to cancer. The α-exon, which comprises one of the extracellular Ig-like domains, is normally included in the brain, but in a variety of gliomas it was found to be skipped, producing an isoform named FGFR1β (Yamaguchi et al. 1994). The skipping of the α-exon is most drastic in the most aggressive glioma, glioblastoma multiforme

(GBM). The FGFR1β isoform has higher affinity for FGF1, a feature that might contribute to tumor growth (Wang et al. 1995). The inclusion of the α-exon was shown to be repressed by

PTB, which is overexpressed in GBMs (Jin et al. 2003).

Rac1

Rac1 is another Ras-superfamily GTPase that, as a result of alternative splicing, can exist as an alternative isoform with important functional consequences in cancer cells. Like Ras, Rac1 cycles between an active GTP-bound and inactive GDP-bound form. The best studied function of the canonical isoform of Rac1 is regulation of cell migration through its control of lamellipodial protrusion (Bosco et al. 2009). Additional roles in cell proliferation, and the creation of reactive oxygen species have also been uncovered (Kheradmand et al. 1998; Bosco et al. 2009). Rac1 has

28 been shown to activate transcription by NFκB, as well as the AKT kinase, both important pathways in many cancers (Keely et al. 1997; Perona et al. 1997). Unlike Ras, however, activating mutations of Rac1 are found infrequently in cancer, although mutations analogous to those that activate Ras have been engineered into Rac1 and the mutant protein found to be capable of transforming NIH3T3 fibroblasts (Qiu et al. 1995; Schnelzer et al. 2000a).

While mutation of Rac1 might be rare, in 1999 it was found that Rac1 pre-mRNA is frequently alternatively spliced in colorectal cancer to produce a previously unknown isoform named Rac1b, which contains a new internal 57 nt exon, 3b (Figure 2i; Jordan et al. 1999). This isoform showed a striking pattern of tumor-specific expression in a set of colorectal tumors when compared to neighboring non-cancerous tissue, and was most upregulated (with reference to the normal Rac1 isoform) in metastatic disease. This was soon followed by a report of the same isoform upregulated in breast cancer (Schnelzer et al. 2000b). Exon 3b and surrounding intronic sequences show considerable conservation in vertebrates, suggesting that 3b inclusion is an important event during development.

Inclusion of exon 3b maintains the reading frame and adds 19 amino acids to a region immediately C-terminal to the switch II region, which, along with switch I, is critical for relaying the GTP/GDP binding status of Rac1 to downstream effectors (Fiegen et al. 2004). Importantly, biochemical assays performed by multiple groups showed that Rac1b exhibits reduced intrinsic

GTPase activity, but increased GDP/GTP exchange, expected to result in a higher level of constitutive activation (Schnelzer et al. 2000b; Fiegen et al. 2004; Singh et al. 2004). Consistent with this observation, Rac1b, unlike Rac1, was able to cooperate with an activated mutant of

Raf-1 to transform NIH3T3 cells (Singh et al. 2004). Expression of Rac1b is critical for cell viability in at least one colorectal cancer cell line, Caco-2 (Matos and Jordan 2008). An important insight into the biological function of Rac1b in cancer was provided by a study showing that induction of Rac1b splicing is critical in mediating malignant transformation mediated by matrix metalloproteinase-3 (MMP-3) (Radisky et al. 2005). Addition of MMP-3 to

29 mammary epithelial cells causes them to undergo EMT, leading to increased invasiveness and to genomic instability. Significantly, this process also leads to the production of Rac1b, and isoform specific knockdown of Rac1b both reversed the effects of MMP3 on cell motility and also led to reduced production of reactive oxygen species which were shown to be responsible for the observed genomic instability (Radisky et al. 2005).

The generation of Rac1b is thus closely connected to fundamental changes, such as EMT, that occur during tumorigenesis, making the regulation of the splicing event extremely interesting. To investigate the regulation of Rac1 splicing, Jordan and colleagues transiently overexpressed a variety of well known splicing factors, and found that SRSF1 overexpression resulted in increased exon 3b inclusion, while SRSF3 (SRp20) and SRSF7 (9G8) had the opposite effect (Goncalves et al. 2009). These results were confirmed using siRNA knockdowns.

The authors went on to show that inhibition of the Wnt/β-catenin pathway resulted in decreased

SRSF3 levels and increased Rac1b. In accordance with this, the transcriptional activity of β- catenin negatively correlated with Rac1b in colorectal cancer cell lines (Goncalves et al. 2008).

Additionally, inhibition of the PI3K/AKT pathway was found to promote increased SRSF1 levels, as well as increased Rac1b. The fact that the Wnt/β-catenin pathway appears to inhibit

Rac1b splicing in colorectal cancer cell lines appears to pose a paradox since this pathway is upregulated during MMP3-mediated EMT (Radisky et al. 2005). Additional work is necessary to unravel the complexity of Rac1 splicing regulation in the various contexts it has been observed.

Ron

Like Rac1, alternative splicing of Ron transcripts, which encode the receptor of the macrophage stimulating protein (MSP), in cancer cells is closely tied to the invasive phenotype.

Ron is a heterodimeric transmembrane receptor tyrosine kinase, composed of an α and β chain, each derived from proteolytic cleavage of a common precursor (Lu et al. 2007). Under normal conditions, binding of MSP to Ron results in tyrosine autophosphorylation of the receptor,

30 leading to activation of signaling pathways that result in increased motility and invasive growth

(Ghigna et al. 2005; Wagh et al. 2008). In 1996, a Ron variant missing exon 11, termed Δ-Ron, was found to be expressed in the gastric carcinoma cell line KATO-III (Figure 2j; Collesi et al.

1996). The Δ-Ron isoform, as well as several additional AS-generated isoforms, have been found in additional epithelial cancers, including colorectal and breast cancer, and expression of these isoforms correlates with metastasis (Zhou et al. 2003; Ghigna et al. 2005). E11 skipping results in an in-frame deletion of 49 amino acids from the membrane proximal extracellular domain, leading to a failure to undergo proper proteolytic processing and constitutive activation of the protein (Collesi et al. 1996).While incapable of transforming NIH3T3 cells, expression of Δ-Ron in two different cell types resulted in increased motility in the absence of MSP, indicating that Δ-

Ron indeed functions as a constitutively active receptor in promoting invasive behavior.

Like the constitutively active Rac1b isoform described above, the production of Δ-Ron can be mediated by SRSF1 (Ghigna et al. 2005). SRSF1 was found to bind to an ESE present in

E12, and to promote E11 skipping when overexpressed. The importance of SRSF1 for Δ-Ron production was confirmed by knockdown experiments that demonstrated conclusively that

SRSF1 expression levels can be an important determinant of the ratio of full-length Ron to Δ-

Ron. Consistent with its effect on Ron splicing, SRSF1 overexpression had a profound effect on cell motility and morphology. Perhaps most interestingly, SRSF1 overexpression resulted in the hallmark changes of EMT, including downregulation of E-cadherin and changes in β-catenin localization. Active Ron is known to activate EMT, and accordingly, the effect of SRSF1 overexpression was reversed by Δ-Ron specific siRNAs, indicating that regulation of Ron splicing is the key event mediating the effect of SRSF1 on cell motility (Bardella et al. 2004;

Wang et al. 2004).

Regulation of angiogenesis: VEGFA

31 Angiogenesis is another area of cancer biology in which AS plays an important regulatory role. VEGFA transcripts, which encode the key ligand secreted by tumors in response to hypoxia to promote the formation of new blood vessels, are extensively alternatively spliced

(Harper and Bates 2008). Perhaps the alternate isoform with the most pertinence to cancer was discovered in 2002, when a variant differing only in the final six amino acids was identified

(Bates et al. 2002; Harper and Bates 2008). This six amino acid change occurred due to the choice of a distal 3’ splice site in the final exon (Figure 2k). The most abundant isoform resulting from use of this 3’ splice site, VEGF165b, was distributed throughout most adult tissues, in some cases representing the major VEGF isoform. While canonical VEGF was broadly upregulated in kidney carcinomas, VEGF165b was undetectable in most tumors examined. The same pattern of

VEGF165b downregulation was observed in a number of other cancer types, including prostate cancer and malignant melanoma (Woolard et al. 2004; Pritchard-Jones et al. 2007).

VEGF165b was quickly shown to be an inhibitor of angiogenesis, providing yet another example of an AS event that produces antagonistic isoforms (Bates et al. 2002; Woolard et al.

2004). How does what appears to be a minor modification in VEGF result in such drastically different properties? The canonical form of VEGFA binds to VEGF receptor 1 (VEGFR1) or

VEGFR2, as well as co-receptors neurophilin 1 (NRP1) and heparan sulfate proteoglycan (HSPG)

(Olsson et al. 2006). Binding to VEGFR2 expressed on endothelial precursors by VEGFA is a critical event in the formation of new blood vessels. VEGFA binding to VEGFR2 as well as

NRP1 results in dimerization of VEGFR2 and activation of receptor tyrosine kinase activity, leading to autophosphorylation of key residues necessary for signaling events required for formation of new vasculature (Olsson et al. 2006). VEGF165b, unlike VEGF165, fails to engage

NRP1 in a multimeric complex on the cell surface, required for full VEGFR-induced signal transduction (Kawamura et al. 2008). VEGF165b binding to VEGFR2 also fails to result in phosphorylation of the critical residue Y1054, which is necessary for full activation of VEGFR tyrosine kinase activity (Kawamura et al. 2008). VEGF165b was originally proposed to inhibit

32 angiogenesis by acting as a competitive inhibitor of isoforms that promote activation of

VEGFR2, although other possibilities, such as activation of alternate signal transduction cascades upon binding VEGFR, cannot be ruled out (Harper and Bates 2008).

Investigation into the regulation of VEGFA splicing has yielded some interesting insights into the pathways and splicing factors involved in regulating the choice between the proximal and distal sites in the final exon. Nowak et al. (2008) examined the effects of three growth factors known to upregulate VEGFA expression on VEGFA splicing (Nowak et al. 2008). They found that insulin-like growth factor 1 (IGF1) and TNFα promoted upregulation of the canonical isoform of VEGFA and downregulation of VEGF165b, while TGFβ1 also upregulated VEGFA expression but had the opposite effect on splicing. Inhibition of the p38 MAPK, activated by

TGFβ1 signalling, reversed the effect of TGFβ1 on VEGFA splice site selection. The increase in

VEGFA expression, as well as the splicing switch in favor of the canonical isoform in IGF1 treated cells was inhibited when cells were simultaneously treated with inhibitors of protein kinase C (PKC) or the SR protein kinases SRPK1/2 (Nowak et al. 2010).

That inhibition of SRPK1/2 affects splice site choice in the VEGFA transcript implies that SR proteins are involved in regulating the choice between the angiogenic and anti- angiogenic forms of VEGFA. Predicted binding sites for SRSF1 and SRSF5 were identified within exon 8a (included in the canonical isoform), upstream of the distal splice site, and overexpression of these two SR proteins indeed promoted an increase in VEGF165/VEGF165b ratio (Nowak et al. 2008). In addition, SRSF6 (SRp55) binding sites were found downstream of the DSS in exon 8b, and SRSF6 overexpression strongly promoted the use of the DSS and expression of VEGF165b (Nowak et al. 2008). Furthermore, IGF1 treatment of cells resulted in an increase in SRSF1 nuclear localization, a phenomenon dependent on SRPK1/2. Although it would be important to confirm these results with appropriate siRNA knockdowns, these findings suggest a pathway that connects a growth factor that promotes vascularization, IGF1, with the alteration of an AS event by several SR proteins in favor of a pro-angiogenic form of VEGFA.

HnRNP and SR proteins in proliferation and cancer

The first proteins identified to regulate AS belong to the SR and hnRNP protein families

(Dreyfuss et al. 1993; Fu 1995; Manley and Tacke 1996). As detailed above, several members of these two protein families have recently been shown to play important roles in proliferation and cancer. Here we will examine the regulation and functions of these proteins.

hnRNP A/B family proteins

HnRNP A1, A2, and A3 constitute the hnRNP A/B family of RBPs. With the exception of hnRNP A3, these proteins have been extensively studied and have been implicated in a variety of processes, and are expressed in a proliferation-associated manner. A connection between increased hnRNP A1 expression and proliferation was established well before the functions of

A1 in regulation of gene expression were understood (LeStourgeon et al. 1978). It was later demonstrated that transcription of the hnRNP A1 gene could be induced by growth factor stimulation of cells, while differentiation of some cell types resulted in decreased A1 levels

(Planck et al. 1988; Minoo et al. 1989; Biamonti et al. 1993). In healthy adult tissues, the expression of hnRNP A1/A2 appears confined to proliferating cells, such as those in the basal layer of the skin, although they are also expressed in some neurons (Patry et al. 2003). Extending the analysis of A1 levels to an in vivo model of mouse lung tumorigenesis, Zerbe et al. (2004) showed that nuclear A1 protein levels were dramatically increased in tumors compared to surrounding non-neoplastic cells (Zerbe et al. 2004). Furthermore, induction of non-neoplastic proliferation in the lung was sufficient to lead to increased hnRNP A1 levels, which were further increased in tumors. Importantly, hnRNP A1 has been shown to be upregulated in a wide variety of cancers, including breast, colorectal, and lung, and, as described above, gliomas (Pino et al.

2003; Ushigome et al. 2005; Li et al. 2009; David et al. 2010). Likewise, hnRNP A2 is consistently overexpressed in a wide range of cancers (Zhou et al. 2001; Yan-Sanders et al. 2002;

34 Wu et al. 2003). HnRNP A2 is expressed at high levels during mouse lung development, then downregulated in the adult lung, indicating an important role during highly proliferative periods of normal development, and confirming the connection between proliferation and hnRNP A2 upregulation (Montuenga et al. 1998).

Increased expression of hnRNP A/B proteins has important functional consequences. For one, it was found to have a protective effect against apoptosis, possibly in part through an effect on alternative splicing of caspase-2 pre-mRNA (Jiang et al., 1998). In addition, depletion of hnRNP A/B proteins in Colo16 cells resulted in reduced proliferation, with hnRNP A2 playing a particularly important role (Jiang et al. 1998; He et al. 2005). Consistent with an important role in tumors, siRNA-mediated knockdown of hnRNP A1/A2 in cancer cells, but not normal cells, can result in apoptosis (Patry et al. 2003).

HnRNP A/B proteins are multifunctional, and many of these functions appear consistent with a role in proliferation. The regulation of pre-mRNA splicing was one of the earliest known functions of hnRNP A1, a function it often carries out by binding to ESS sequences and repressing exon inclusion (Mayeda and Krainer 1992; Del Gatto-Konczak et al. 1999). HnRNP

A1 bound to ISS sequences can also promote exon exclusion, possibly through a mechanism that involves self-interaction of A1 molecules bound to distal sites and loop formation (Blanchette and Chabot 1999; Kashima et al. 2007). While best characterized as repressors of splicing, hnRNP A/B proteins can also activate inclusion of some exons (Martinez-Contreras et al. 2006;

Venables et al. 2008). As observed in PKM splicing regulation (David et al., 2010), hnRNP A1 and A2 frequently function redundantly in splicing regulation (Kashima and Manley 2003;

Licatalosi and Darnell 2010). However, a genome-wide approach indicated that hnRNP A1/A2 targets may in fact be quite divergent (Venables et al. 2008). Additional investigation is required to establish the extent of their redundancy, and whether any redundancy exists with a third family member, hnRNP A3, which shares more identity with hnRNP A1 than A2 does. A recent report demonstrated a unique function for hnRNP A2 connecting AS regulation with invasive

35 tumor cell behavior (Moran-Jones et al. 2009). These authors found that hnRNP A2 was required for two different cell lines to invade matrigel or to migrate on a cell derived matrix. Using an exon array, they also showed that hnRNP A2 promotes inclusion of exon 2 in transcripts of the gene encoding a p53 target, TP53INP2, in invasive cells, and that inclusion of this exon is important for invasive behavior.

Despite its important role as a splicing repressor, there have been relatively few reports implicating changes in hnRNP A/B levels in regulation of AS during normal development. In one example, hnRNP A1 and A2 were shown to repress exon 16 inclusion in the protein 4.1R transcript. Differentiation of erythroblasts accompanied an AS switch to exon 16 inclusion, a process that coincided with downregulation of hnRNP A1 and A2 (Hou et al. 2002). Because upregulation of hnRNP A1/A2 appears to be shared by most if not all proliferating cells, it will be interesting to see if additional proliferation/differentiation associated changes in AS are the result of changes in hnRNP A1/A2 levels.

In addition to its role in splicing, hnRNP A1 has numerous additional functions that involve nucleic acid binding. In another role that is likely significant in cancer, human A1 has been implicated in lengthening and maintaining telomeres, a function that is conserved in C. elegans, and possibly yeast (Lin and Zakian 1994; LaBranche et al. 1998; Joeng et al. 2004). A1 binds to the single-stranded telomeric DNA repeat sequence TAGGGT (a sequence almost identical to the high-affinity SELEX binding site identified for A1 using RNA), where it is capable of stimulating telomerase activity (Burd and Dreyfuss 1994; Zhang et al. 2006). A similar function has been shown for hnRNP A2 (Kamma et al. 2001). HnRNP A1 shuttles between the nucleus and cytoplasm, and also has a well-established role in regulating translation of mRNAs through its binding to internal ribosome entry sites (IRES), present in the 5’ untranslated region (UTR) of many mRNAs. Binding to such sites in the cyclin D1 and c-Myc mRNAs results in increased translation, making hnRNP A1 an important positive regulator of critical proto-oncogene expression (Jo et al. 2008). The fact that hnRNP A1 is transcriptionally

36 upregulated by c-Myc and itself promotes the expression of c-Myc (see above) indicates that these two proteins may constitute a positive feedback loop in some circumstances. An additional recently discovered activity of hnRNP A1 is in microRNA (miRNA) biogenesis. A1 was shown to promote processing of miRNA-18a, a miRNA frequently overexpressed in cancer (Guil and

Caceres 2007; Motoyama et al. 2009). In light of its newfound role in regulating expression of miRNAs, it will be interesting to see if changes in hnRNP A1 expression are responsible for some of the large-scale changes in miRNA expression observed in cancer.

PTB

PTB also has a myriad of functions in cancer. The splicing-repressive functions of PTB are best studied, although it is also clear that it can promote inclusion of some exons (Xue et al.

2009). Like hnRNP A/B proteins, PTB is expressed throughout development, then downregulated in many adult tissues, most notably brain and muscle (Boutz et al. 2007a;

Makeyev et al. 2007). In differentiating neurons, PTB is replaced by a related protein, nPTB, which binds to similar sequences but functions as a weaker splicing repressor than PTB

(Markovtsov et al. 2000; Boutz et al. 2007b). PTB has been shown to be overexpressed in ovarian cancer as well as gliomas (Jin et al. 2003; He et al. 2007; David et al. 2010). Like hnRNP A2, PTB promotes invasive behavior in a number of cancer cell types (He et al. 2007;

Cheung et al. 2009). In glioma-derived cells, the increased inclusion of exon 3 in the RTN4 transcript, which is inhibited by PTB, appears to underlie the adverse effects of PTB depletion on cell migration (Cheung et al. 2009). PTB also exerts an effect on cell migration by binding to mRNAs encoding vincluin and α-actinin 4 and localizing to focal adhesions upon cell adhesion

(Babic et al. 2009). PTB depletion reduced the number of cell protrusions, and reduced the amount of vinculin mRNA present at the cellular edge.

PTB also plays significant roles in translation. In another parallel with hnRNP A1, PTB has been found to bind to the c-Myc IRES sequence where it upregulates c-Myc translation

37 synergistically with another RNA-binding protein, YB-1 (Cobbold et al. 2010). These authors found a striking correlation between c-Myc and PTB protein levels, both of which were elevated in multiple myeloma cell lines compared to B-cell lines. A similar correlation between c-Myc and PTB (as well as hnRNP A1/A2) levels was observed in gliomas (David et al. 2010). Like hnRNP A1/A2, the level of PTB in some cell types is upregulated by c-Myc, again potentially forming the basis of a positive feedback loop. In addition to c-Myc, PTB was found to bind to most if not all putative IRES elements present in a large number of target mRNAs involved in proliferation and apoptosis (Mitchell et al. 2005). The translational activity of PTB was shown to be important for the upregulation of translation of a subset of apoptotic mRNAs upon the induction of TRAIL-induced apoptosis (Sawicka et al. 2008). In addition, PTB binds to the IRES of the cyclin-dependent kinase inhibitor p27Kip1, and promotes its translation (Cho et al. 2005).

Depletion of PTB by siRNA-mediated knockdown resulted in a shortened G1 phase, indicating that control of p27Kip1 may play an important role in cell cycle regulation. So, while PTB is frequently overexpressed in cancer, it can have pro-apoptotic and anti-proliferative effects through its effect on translation of specific mRNAs. cAMP-dependent protein kinase A phosphorylation of PTB results in its redistribution to the cytoplasm, a phenomenon shown to result in PTB-mediated stabilization of insulin mRNA in response to cAMP signaling (Xie et al.

2003; Knoch et al. 2006). It will be of interest to determine in more detail how the various activities of PTB are regulated during development and dysregulated in cancer.

SRSF1

The SR protein with the best-demonstrated role in cancer is SRSF1. As discussed above,

SRSF1 has been implicated in a number of cancer-associated changes in AS events related to cell motility (Rac1 and Ron) and proliferation (Cyclin D1). Consistent with this, Krainer and colleagues demonstrated that the gene encoding SRSF1 can function as a proto-oncogene (Karni et al. 2007). SFSR1 expression was shown to be upregulated frequently in cancer (sometimes by

38 amplification of its chromosomal locus). In addition, overexpression of SRSF1 resulted in transformation of immortalized cell lines while reduction of SRSF1 levels in transformed cells reversed the malignant phenotype. The authors hypothesized that some of the effects of SRSF1 overexpression on the transformed phenotype where due to altered AS, and examined candidate genes in signaling pathways frequently deregulated in cancer for changes in AS. For one gene, encoding the translational regulator S6K1, alternative splicing/polyadenylation creates a truncated product, dubbed isoform 2. This mRNA, and its truncated protein product, was upregulated in cells overexpressing SRSF1, and isoform 2 itself was found to be sufficient to transform cells. In addition, SRSF1 overexpression promoted AS of the mRNA encoding the tumor suppressor BIN1, producing an isoform with no tumor suppressor activity. SRSF1 also plays a more direct role in translation, promoting cap-dependent translation requiring eIF4E, the deregulation of which can have oncogenic consequences (Karni et al. 2008; Michlewski et al.

2008). This appears to occur in part due to the activation by SRSF1 of mTORC1 through an unknown mechanism (Karni et al. 2008) Like hnRNP A1, SFSR1 has been shown to have a splicing-independent role in promoting the maturation of several miRNAs, some of which are themselves upregulated in cancer (Wu et al. 2010).

Control of SR protein phosphorylation by signaling pathways

Phosphorylation of SR proteins, most notably on serines in the serine and arginine-rich domains that give them their name, is an important determinant of their localization and activity

(Shepard and Hertel 2009). Regulation of SR protein activity by phosphorylation has now been implicated in a cancer-associated AS event. Specifically, the EDA exon of the fibronectin (Fn) transcript, an extracellular matrix protein involved in cell adhesion and migration is expressed at low levels in adult tissues, and its inclusion has been shown to be a highly specific marker of liver metastasis (Rybak et al. 2007). Importantly, the EDA-containing variant appears to promote cellular proliferation to a greater extent than the EDA-lacking variant, indicating a potential

39 functional importance in cancer progression (Manabe et al. 1999). Srebrow and colleagues demonstrated that growth factors can promote EDA inclusion, and went on to demonstrate that this required the activity of PI3K (Blaustein et al. 2004). Furthermore, activation of the AKT kinase, a key mediator of PI3K signaling in response to growth signals, was capable of activating

EDA inclusion (Figure 4; Blaustein et al. 2005). AKT was shown to be capable of phosphorylating SR proteins SRSF1 and SRSF7 in vitro (Blaustein et al. 2005). These two proteins had been shown to activate splicing of the EDA exon, suggesting the possibility that direct AKT phosphorylation of SR proteins might underlie the change in Fn splicing in response to growth factors. AKT phosphorylation of SRSF1 and SRSF7 occurred mainly in the RS domain, which contain multiple consensus AKT phosphorylation sites (RXRXXS/T). However, the affect of AKT activation of Fn splicing was opposite the effect of overexpression of either

SRPK1 or another SR protein kinase, Clk/Sty (Colwill et al. 1996), and the intracellular distribution of the SR proteins was unchanged in response to AKT activation, in contrast to

SRPK1 and Clk/Sty activation. This indicates that SR protein phosphorylation by AKT differs from that by SRPK1 and Clk/Sty. Providing additional support for the notion that AKT activation can affect splicing through SR protein phosphorylation, another study suggested that

SRSF5 may similarly be regulated by AKT phosphorylation (Patel et al. 2005). Future studies will be necessary to show that the effects of AKT on AS are indeed mediated by direct SR protein phosphorylation, and if so, how AKT phosphorylation of SR proteins affects their splicing activities.

Conclusions

The above examination of a limited number of important AS events in cancer provides only a small window into the regulation of splicing in cancer. However, a few themes are apparent from these examples. For one, some growth pathways are able to influence splicing through their control of splicing factor expression or post-translational modifications. C-Myc-

40 mediated upregulation of PTB, hnRNP A1, and hnRNP A2, AKT phosphorylation of SR proteins, and the regulation of Sam68 activity by ERK and Fyn provide examples of this. The dysregulation of these pathways in cancer thus contributes to large-scale changes in AS as well as to alterations in other areas of gene expression controlled by these proteins. Gaining a better understanding of the pathways that lead to upregulation of key AS regulatory proteins will be of future interest. Because post-translational modifications often have profound effects on the activity of splicing factors, a full understanding of AS regulation in cancer will likely require additional discoveries in this challenging area. A second theme that emerges is that RBPs that regulate AS are all multifunctional, and many of their activities (with some exceptions) appear to be functionally aligned. Systematic analysis of cancer-associated RBPs through the use of genome-wide methods will identify additional targets for these proteins and provide further insight into the extent of “biological coherence” in the processes they regulate (Licatalosi and

Darnell 2010). But it should also be apparent that additional mechanistic studies will be need to understand fully the diverse functions of these proteins, and how they contribute to control of cell proliferation in normal and transformed cells.

There is certainly much more to learn about splicing regulation in cancer. The list of splicing regulatory proteins that control AS in cancer is very likely to expand, as is the number of genes affected. Nonetheless, significant insights have been obtained from examination of the relatively small number of splicing events reviewed here, and it is now apparent that dysregulation of alternative splicing plays critical roles in numerous cancers, and at multiple points in disease progression. It is hoped that future studies will not only provide a deeper understanding of the role of AS in cancer, but also suggest possible therapeutic approaches.

Acknowledgements

41 We thank Mo Chen and other members of the Manley laboratory for helpful discussions, and Jim Duffy for help with the figures. Relevant work in our laboratory is supported by grants from the NIH.

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Figure legends

Figure 1. Combinatorial control of splicing by RBPs. In this schematic, a regulated exon

(yellow) is flanked by two constitutively spliced exons (blue). Regulatory elements lying within the exon are known as exonic splicing enhancers (ESEs) and exonic splicing silencers (ESSs), while intronic regulatory elements are referred to as intronic splicing enhancers (ISEs) and intronic splicing silencers (ISSs). The trans-acting factors that bind to these elements can be regulated by changes in intracellular levels, as well as post-translational modifications that affect their cellular localization or activity. The balance of positive and negative-acting factors present in a given cell determine the extent of regulated exon inclusion.

Figure 2. Schematic representation of the AS events discussed in this review. In each case, isoforms that are upregulated in cancer, or otherwise shown to have positive effects on growth, survival, or invasive behavior are shown at the top of each diagram.

67 Figure 3. Caspase-2 and H-ras alternative exons lie in highly conserved regions. Images taken from UCSC genome browser, using the March 2006 assembly, available at http://genome.ucsc.edu/, (Kent et al. 2002) with exon/intron structure of each gene indicated at top, and the mammalian conservation of the corresponding regions indicated below. Conserved regions are indicated by blue bars. a. For caspase-2, the alternatively spliced exon 9 is highly conserved, and is also flanked by conserved regions, a hallmark of regulated alternatively spliced exons. b. The IDX exon of H-ras lies within a similarly highly conserved region.

Figure 4. Selected signal transduction pathways that affect AS of transcripts of genes important in cancer. From top left, hnRNP A1 promotes translation of c-Myc mRNA. Translated c-Myc protein can promote the transcription of PTB, hnRNP A1, and hnRNP A2, which in turn promote the production of PKM2. The PI3K pathway is activated downstream of ligand binding by receptor tyrosine kinases (RTK), activating AKT, which phosphorylates SR proteins such as SRSF1, leading to altered splicing of fibronectin (Fn) transcripts. IGF1 binding to the IGF receptor (IGF-R) results in activation of the SR protein kinases SRPK1/2, which phosphorylate SRSF1 and alter VEGF165 splicing in favor of the pro-angiogenesis isoform. Finally, Fas receptor activation by binding to FasL on another cell results in FAST K activation, which in turn results in TIA-1 phosphorylation and inclusion of exon 6 in the Fas transcript.

68 69 70 71

Chapter 2: The search for alternative splicing regulators:

new approaches offer a path to a splicing code.

The search for alternative splicing regulators: new approaches offer a path to a splicing code.

Charles J. David and James L. Manley

Department of Biological Sciences, Columbia University, New York, NY,

10027.

74 Complex multicellular organisms require a diverse set of proteins to set the form and function of specialized cell types. The availability of complete genomic sequences has revealed that instead of a large increase in the number of protein coding genes compared with unicellular organisms, more eukaryotes instead obtain more diversity out of a relatively limited number of genes through the process of alternative splicing (AS).

AS results in the cell type-, developmental stage-, sex-, or signal-regulated changes in composition of an mRNA produced from a given gene, brought about by changes in splice site choice (Black 2003; Matlin et al. 2005). There are many different types of AS events, ranging from the tissue-specific inclusion of a cassette exon to the Dscam gene in

Drosophila, which contains four clusters of exons containing 12, 48, 33, and 2 mutually exclusive variants, an extreme example of AS complexity (Bharadwaj and Kolodkin

2006). Additionally, it is clear from the relatively small number of AS events that have been extensively studied at a mechanistic level that regulation of AS takes many forms, as will be discussed in more detail later in this article. Lastly, the functional outcomes of

AS vary greatly, from effectively turning off a gene (the result of including an exon containing a premature stop codon, for example), to a subtle change in a protein’s function.

In spite of this complexity, the goal of understanding the changes in AS patterns in terms of changes in expression and regulation of factors that regulate AS across cell types appears achievable. This is in part because recent technical advances allow us, starting with an individual splicing factor, to determine its genome-wide role in AS regulation. However, the ambitious goal of determining a cellular code for AS will be

75 impossible to realize without a complete list of AS regulators, and currently there is no reason to believe we are anywhere near completing such a list in any metazoan.

Kuroyanagi and colleagues have recently developed a system in C. elegans which allows for the straightforward visualization of individual alternative splicing events in vivo, which in turn allows for the identification of mutants defective in regulating the event (Kuroyanagi et al. 2006). Using this approach, the Kuroyanagi group has identified four proteins that regulate two different AS events, a feat that would be laborious using in vitro approaches (Kuroyanagi et al. 2006; 2007; Ohno et al., this issue). Three of these proteins had not previously been shown to participate in regulation of AS, indicating that we may be far from a complete list of AS regulators. Their data also highlight the fact that regulators of AS are often highly conserved throughout metazoans, meaning that filling the list of splicing regulators in C. elegans will likely contribute to filling our own.

Known AS regulators

There are a large number of factors known to be involved in AS, which can be crudely lumped into two classes. One class of AS regulators consists of relatively widely expressed proteins, which seem to have wide-ranging roles in mRNA biogenesis. These come in two groups: SR proteins, which, when bound to exons, tend to promote exon inclusion, and hnRNP proteins, which usually have the opposite effect (Manley and

Tacke 1996; Graveley 2000; Smith and Valcarcel 2000). While these proteins are widely expressed and have roles in numerous processes, cell type-specific changes in their expression levels or post-translational modifications that alter their activities or localization provide a means of alternative splicing regulation. For example, hnRNP A1, which inhibits the inclusion of many alternative exons, becomes phosphorylated upon

76 osmotic shock resulting in its cytoplasmic accumulation, which in turn affects a number of AS events (Allemand et al. 2005). While we are very far from knowing all the activities of the hnRNP and SR proteins in AS regulation, it is safe to assume that all members of these widely expressed families have been identified, although some tissue- specific isoforms relevant to AS may yet be discovered.

No such confident statements can be made about the other broad class of AS regulators, proteins with a much narrower job description. These are factors with restricted expression patterns that are responsible for regulating tissue-specific AS events.

The idea that factors of restricted expression regulate specific AS events has been around since the elucidation of the Drosophila sex determination pathway. This pathway begins with a female-specific RNA binding protein, Sxl, which regulates the splice site choices of a small number of transcripts that are alternatively spliced in a sex-specific manner

(Lopez 1998). Additional tissue-specific factors (often small families of factors) that play a role in regulating AS of a relatively small number of transcripts have been identified and characterized to varying extents. In mammals, these include Nova-1/2

(Jensen et al. 2000), TIA1/TIAR (Del Gatto-Konczak et al. 2000; Forch et al. 2000), Fox-

1/2 (Jin et al. 2003), nPTB (Markovtsov et al. 2000), STAR/GSG family proteins

(Butcher and Wickens 2004), Tra2α/β (Tacke et al. 1998), CELF family proteins (Ladd et al. 2001), and Hu proteins (Zhu et al. 2006). These proteins have been identified by a wide variety of approaches, and each of the factors listed shares a salient feature: each contains an RNA binding domain of the KH or RRM type.

For many of the specific AS regulators we currently know, the initial implication in AS was based solely on the presence of an RNA binding motif. Using databases made

77 possible by complete genomic sequence available in mouse, Silver and colleagues sought to compile a comprehensive list of mouse RNA binding proteins based on known RNA binding motifs. The resulting set contained 380 proteins, most of which contained RRMs or KH domains (McKee et al. 2005). While this is a large number, it does not approach that of transcription factors, which, according to some estimates number up to 2500

(Tupler et al. 2001). McKee et al. showed that 221 of these putative RNA binding proteins exhibit regionally restricted expression in the brain. A large number of the proteins identified as putative RNA binding proteins were minimally characterized, or not characterized at all. So while the list of known specific AS regulators in mammals numbers under two dozen, this genome-wide analysis suggests that there is room for that number to expand. It should be pointed out that AS regulators do not need to contain an

RNA binding domain to control specific AS events; here the Drosophila sex determination pathway offers another lesson. The female-specific Tra protein, devoid of any known RNA-binding domain, is required for the cooperative assembly of RBP1 and other SR proteins, as well as the SR-related protein Tra2, on the dsx enhancer element

(Tian and Maniatis 1994). This type of regulation, analogous to that of transcriptional coactivators, has also been demonstrated in humans. SRm160, an RS-domain containing splicing factor without an apparent RNA binding domain participates in the regulation of

CD44 variable exon 5 splicing (Cheng and Sharp 2006).

Also of interest is the potential influence of transcription itself on AS, and the possibility that transcription factors might serve as AS regulators. A decade ago,

Kornblihtt and colleagues made the observation that the identity of the promoter driving a given transiently transfected minigene encoding an alternatively spliced transcript can

78 alter the outcome of the AS event (Cramer et al. 1997). This finding raised the intriguing possibility that transcription factors could serve as AS regulators, either by influencing the concentration of direct regulators of AS within the alternatively spliced gene’s transcriptional milieu, or by altering the rate of RNA polymerase II elongation, leading to indirect effects on AS (Kornblihtt 2005). An example of a transcription factor that can influence splicing was provided by the mammalian protein PGC-1, a transcriptional coactivator involved in adaptive thermogenesis. This protein, in addition to domains relevant to transcription, contains an RRM as well as an RS-rich domain, both characteristic of splicing factors, and these apparently influence the splicing of transcripts it regulates (Monsalve et al. 2000). Identifying instances in which tissue-specific changes in transcription factor expression alter AS patterns in a direct and physiologically relevant way will be of interest in the future.

Most of the factors listed above as tissue-specific AS regulators were of interest prior to their implication in AS. In fact, very few AS factors have been first characterized solely on the basis of their participation in AS. Identifying regulators of a particular AS event, starting from scratch, is difficult. A blueprint for this approach is provided by the work of Black and colleagues on the neuron-specific inclusion of the c-src N1 exon. The first step was the reconstitution of the AS pattern in cultured cells (Black 1992). Next, cis-elements required for the regulation of the event were mapped (Black 1992; Chan and

Black 1995; Modafferi and Black 1997). These sequences in hand, RNA affinity chromatography was used to identify factors that bind to these sequences in nuclear extracts, which led to the identification of two novel AS regulators, KSRP and nPTB

(Min et al. 1997; Markovtsov et al. 2000). While this approach was successful in

79 identifying novel AS regulators, it is too involved to be applied to a large number of AS events. Another way of identifying regulators of particular AS events is through RNA interference screens (Celotto and Graveley 2004). Genome-wide screens have already been applied to other mRNA processing events (Wagner et al. 2007), and this approach

will certainly prove useful in the analysis of alternative splicing regulation. One

shortcoming of this approach is that alternative splicing events must first be recapitulated

in cell culture, which may not always be possible.

A genetic approach to identifying AS regulators in C. elegans

Kuroyanagi and colleagues have developed a system that allows for the visualization of AS decisions in intact organisms (Kuroyanagi et al. 2006). The method involves the use of minigene reporters that express red fluorescent protein (RFP) or green fluorescent protein (GFP), dependent on the cell-specific outcome of the AS event being studied. In this issue of Genes and Development, Ohno et al. apply this method to the mutually exclusive splicing of let-2 exons 9/10. Let-2 encodes α2(IV) collagen in C. elegans, and expression undergoes a switch during development, from an mRNA isoform that contains exon 9 (E9) in embryos to an isoform that instead contains exon 10 (E10) during late larval stages and in the adult worm. This AS event was first reported 15 years ago, but until now nothing was known about the factors that regulate it (Sibley et al.

1993).

Ohno et al. constructed two reporter minigenes, each containing either GFP or

RFP downstream of a region of the let-2 gene spanning exon 8 to exon 11, and under the control of the body wall specific promoter myo3. To construct a minigene that produces

80 GFP when E9 is selected, the authors altered a single nucleotide in E10 to introduce a stop codon upstream of GFP, should it be included. In the other minigene, the stop codon was instead introduced in E9, upstream of RFP, resulting in RFP expression only when

E10 is selected. Worms containing these two minigenes showed a distinct switch from green in embryos to red in more mature organisms, in a way that precisely matched the switch from E9 to E10 usage in the endogenous let-2 transcript.

Armed with the ability to monitor visually the E9/E10 switching phenotype, the authors next screened for mutants defective in the regulation of this event. Of the several independent mutant alleles they identified, all resulted in worms that remained green throughout development. Remarkably, all the mutations were in the same uncharacterized gene, which the authors named alternative-splicing-defective-2 (asd-2).

Expressed sequence tag (EST) analysis indicated that this locus produced two mRNA isoforms containing alternate first exons, predicted to result in proteins differing at the N- terminus, ASD-2a and ASD-2b. Of these, ASD-2b was the only isoform detected in body wall muscles. Three null alleles identified were predicted to specifically disrupt the

2b isoform, while another five were missense mutations that fell within a conserved

GSG/STAR domain (see below). Overexpression of ASD-2b in body wall muscles led to increased E10 inclusion earlier in development than normally observed, while RNAi depletion of ASD-2b phenocopied the original asd-2 mutants.

The GSG/STAR domain is the signature of the STAR family of RNA binding proteins, a highly conserved group of multifunctional regulators of gene expression found throughout metazoans. These include the C. elegans protein GLD-1, known to mediate translational repression of some target mRNAs (Schumacher et al. 2005), and quaking

81 (QKI), which has been appears to regulate AS in mammals (Wu et al. 2002; Schumacher

et al. 2005) . A QKI high-affinity binding site has been determined in vitro and named

the quaking response element (QRE) (Galarneau and Richard 2005). Galarneau and

Richard also observed that previously identified GLD-1 binding sites fall within QREs,

demonstrating conservation of RNA binding specificities among the STAR family.

Previous studies had indicated that a short stretch of let-2 intron 10, conserved

between C. elegans and C. briggsae, is required for the proper regulation of exon 9/10

splicing (Kabat et al. 2006). Inspection of this region revealed a consensus QRE, to

which ASD-2b indeed bound in vitro. Disruption of the consensus QRE by two point mutations drastically reduced ASD-2b binding in vitro, phenocopied the splicing defect

of the asd-2 mutants isolated in the original screen, and failed to respond to ASD-2b

overexpression, making a persuasive case that ASD-2b functions through this element to

promote the switch from E9 to E10 inclusion during development. An additional

observation, however, makes it clear that we are not yet privy to the whole story: ASD-2b

is uniformly expressed throughout development. One potential explanation for the

inactivity of ASD-2b early in development is that it is subject to post-translational

modification that regulates its RNA binding activity, as is the case for the closely related

mammalian protein Sam68 (Matter et al. 2002). Another possibility is that ASD-2b

requires another factor to stabilize its RNA binding, while a third possibility is that ASD-

2b, once bound, cooperates with another factor to promote exon 10 inclusion (discussed

below).

Analysis of the splicing intermediates in the region spanning exons 8-11 provides

some insight into the mechanism by which ASD-2b functions in promoting the switch

82 from E9 to E10. In wild-type embryos, a species retaining intron 8 in which E9 and E11

are ligated together accounts for nearly all partially spliced RNA. In L4 stage worms,

this species is greatly reduced, and instead a form in which E10 and E11 are joined

predominates (Figure 1). In this intermediate, the E9 and E10 splice acceptors are in

competition. The E10 acceptor site is favored in this competition, apparently because the

region upstream of it contains a better binding site for the general splicing factor U2AF65

than the region upstream of E9. The E10/E11 intermediate is therefore exclusively

processed to remove E9. In asd-2 mutants, however, the predominant partially spliced

form throughout development remains the E9/E11 form. This suggests that the primary

function of ASD-2b in effecting the switch from E9 to E10 is mediated by binding to an

intronic site and promoting the joining of E10 with E11. This is not the end of the story

as the authors also note that there is an increase in the amount of an E8/E10 splicing

intermediate in adult worms that is not significantly decreased in the asd-2 mutants, indicating that additional factors are at play in promoting the E9 to E10 switch. The additional factor(s) regulating this event may be essential for early embryonic development in C. elegans, which would explain its failure to be detected in the mutant screen.

What is the mechanism by which ASD-2b mediates exon inclusion? The activity of ASD-2b in let-2 splicing is reminiscent of a number of other alternative splicing factors that bind downstream of alternative exons they positively regulate. One regulator,

TIA1, functions by directly interacting with U1 snRNP to recruit it to the upstream 5’ splice site (Figure 2) (Del Gatto-Konczak et al. 2000; Forch et al. 2000). This mechanism was originally predicted because TIA1 is homologous to the yeast U1 snRNP

83 protein Nam8p (Del Gatto-Konczak et al. 2000; Forch et al. 2000). It may be that ASD-

2b functions in a related way. However, given the diversity of other AS regulators that also function by binding to downstream introns, it is possible that not all function through

direct interactions with the splicing machinery. Instead, some intron-bound AS

regulators may act by intron definition. This could occur through an interaction between

proteins bound at either end of the intron downstream of the regulated exon, leading to

the looping out of intervening intronic sequence, bringing the regulated exon’s donor site

into proximity with the downstream acceptor site (Figure 2). This mechanism has been

suggested for some instances of splicing enhancement mediated by hnRNPs binding to

intronic splicing enhancers, as well as for the Nova proteins (Martinez-Contreras et al.

2006; Ule et al. 2006). If this is the case in let-2 regulation, other conserved regions of

intron 10 which do not bind ASD-2b may come into play, along with (an) additional

factor(s) not identified in the screen, offering another conceivable explanation for why

ASD-2b expression throughout development is insufficient to affect splicing early in

development.

Three conserved splicing regulators identified in another AS event

Kuroyanagi and colleagues recently used the same approach to study another AS

event: tissue-specific expression egl-15 E5A-containing isoform, rather than the alternative E5B isoform, in sex myoblasts, daughter cells of which are destined become body wall muscles, as well as specialized sex-specific muscles later in development.

Mutant screening revealed three regulators of this event, FOX-1, ASD-1, a previously uncharacterized member of the Fox-1 family, and SUP-12, an RRM-containing protein previously shown to regulate AS (Anyanful et al. 2004; Kuroyanagi et al. 2006;

84 Kuroyanagi et al. 2007). In another example of the conservation of AS regulators and the

sequences to which they bind, FOX-1 and ASD-1 were shown to function through a

conserved UGCAUG sequence located upstream of the mutually exclusive exons. This

sequence is identical to that bound by mammalian Fox-1/2 (Jin et al. 2003). This is

unsurprising since all residues shown to mediate the specific recognition of the

UGCAUG sequence in human Fox-1 are conserved in the C. elegans proteins (Jin et al.

2003; Auweter et al. 2006). FOX-1 and ASD-1 are more widely expressed than the E5A

isoform, and therefore their expression pattern does not explain the tissue specificity of

the egl-15 AS. Additional screening revealed another factor required for the expression

of the E5A isoform, SUP-12. SUP-12 is a muscle-specific protein which cooperates with

FOX-1 and ASD-1 to bind stably to the egl-15 pre-mRNA and promote the inclusion of

E5A (Kuroyanagi et al. 2007). SUP-12 and its poorly-characterized human homolog

RNPC1 share a higher degree of identity than even the Fox-1 family. Given the high

degree of conservation between other known AS regulators, RNPC1 is highly likely to be

an AS regulator in mammals.

From the identification of AS regulators to a cellular code of AS regulation

Thanks to some important technical developments, known AS regulators, as well

as potential ones such as RNPC1, can be evaluated for their transcriptome-wide RNA

binding. From this, their contribution to the AS of alternative exons can be predicted

genome-wide. This is due in part to the development of the crosslinking and

immunoprecipitation (CLIP), a procedure pioneered by Darnell and colleagues (Ule et al.

2003). Briefly, CLIP involves the UV irradiation of a tissue of interest to induce crosslinks between RNA and bound proteins. Following light RNase treatment to reduce

85 the average length of RNA fragments to ~50 nt, the protein of interest is

immunoprecipitated under stringent conditions and then digested with proteinase K,

leaving RNA that can be amplified and cloned (Ule et al. 2003), hybridized to a

microarray, or amplified and sequenced directly, now using high-throughput technology

(J. Sanford, personal communication). The latter two approaches have the potential to

identify the genome-wide distribution of a given factor. As has been demonstrated with

Nova, an RNA map of this kind makes it possible to predict the effect that a given AS

regulator has in regulating individual alternative splicing events based on binding and

splicing patterns observed in a much smaller subset of genes (Ule et al. 2006). In the

case of Nova, binding sites within alternate exons predicted its negative regulation of that

exon, whereas binding at specific sites within the flanking introns was predictive of

positive regulation, raising the possibility that AS regulators function through non-

specific mechanisms based solely on the position of their binding sites. It will be of great interest to create similar RNA-binding maps for additional factors that regulate AS, both

the ubiquitous (SR proteins and hnRNPs) and the tissue-specific proteins. In light of the

diversity of AS regulators that function through intronic binding sites to positively

regulate exon inclusion, the identification of shared patterns between different families of

factors may illuminate the mechanisms through which they work. Additional methods

such as RNP immunoprecipitation-microarray (RIP-Chip) will supplement CLIP to

provide complete maps of RNA binding factor distribution (Keene et al. 2006).

Conclusions and perspectives

Methods such as CLIP have made it possible to map RNA binding factors throughout the genome, while the advent of technologies such as AS microarrays have

86 also greatly increased the number of AS events of which we are aware (e.g., see Pan et al.

2004). The combination of the two approaches may prove useful in explaining the tissue

specificity of a large number of AS events, but if our catalog of regulators is not complete, it will not explain all AS events.

In contrast to previous approaches, the major advantage to the system developed by Kuroyanagi and colleagues is that it can be widely applied to additional AS events in

C. elegans, as well as other genetically tractable models such as Drosophila. Because so many of the AS regulators identified using this method in C. elegans have clear mammalian homologues, this approach will certainly prove useful in identifying candidate regulators in humans. Widespread use of this system is likely to unearth many additional regulators of AS that are currently on the long list of uncharacterized proteins containing an RNA binding domain. We may also be reminded that, as we learned long ago in a genetic screen for Drosophila sex determination genes, an RNA binding domain is not a requirement for membership in the club of AS regulators. While the complexity of AS is at first glance daunting, systematic approaches to identifying regulators of AS combined with novel methods that allow us to assess the genome-wide activity of individual splicing factors, mean that elucidation of an alternative splicing code may be feasible.

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93 Figure Legends

Figure 1. ASD-2b promotes E10 inclusion by altering 5’ splice site usage in the intron

downstream of the regulated exons. In embryos, E10 is removed in a fast splicing event,

leaving a product with E9 and E11 ligated together as the predominant partially spliced

isoform, which is processed to remove intron 8 in a slower reaction. In adults, ASD-2b,

through an unknown mechanism, promotes usage of the E10 donor site in the fast

reaction. This results in a competition between E9 and E10 acceptor sites. E9 is

removed because it lies downstream of a weaker 3’ splice site.

Figure 2. Potential mechanisms for exon inclusion stimulated by AS regulators binding

to the downstream intron. In the recruitment pathway, the intron-bound AS regulator

promotes exon recognition through specific contacts with the splicing machinery. TIA1

recruitment of U1 snRNP to 5’ splice sites of exons that it regulates serves as an example.

In the conformational pathway, regulators bound to an intronic site promote regulated exon inclusion by intron definition. This might involve an interaction between regulators bound at either end of the intron that would serve to loop out intronic sequences and

bring the regulated exon into proximity with the downstream constitutive exon, thereby

favoring its inclusion.

23 96

Chapter 3: HnRNP proteins controlled by c-Myc deregulate pyruvate kinase mRNA splicing in cancer.

HnRNP proteins controlled by c-Myc deregulate pyruvate kinase mRNA splicing in cancer

Charles J. David1,*, Mo Chen1,*, Marcela Assanah2, Peter Canoll2, and James L. Manley1

1Department of Biological Sciences, Columbia University, New York, NY 10027. 2Department of

Pathology and Cell Biology, Columbia University, New York, NY,10032. *These authors contributed equally to this work.

Correspondence to: James L. Manley1. Correspondence and requests for materials should be addressed to J.L.M. ([email protected])

98 When oxygen is abundant, quiescent cells efficiently extract energy from glucose

primarily by oxidative phosphorylation, while under the same conditions tumor cells consume glucose more avidly, converting it to lactate. This long-observed phenomenon is

known as aerobic glycolysis1, and is now understood to be important for cell growth2, 3.

Because aerobic glycolysis is only useful to growing cells, it is tightly regulated in a proliferation-linked manner4, in part through control of pyruvate kinase (PK) isoform

expression. The embryonic isoform, PKM2, is almost universally re-expressed in cancer2, and promotes aerobic glycolysis, while the adult isoform, PKM1, promotes oxidative phosphorylation2. These two isoforms result from mutually exclusive alternative splicing of the PKM pre-mRNA, reflecting inclusion of either exon 9 (PKM1) or exon 10 (PKM2).

Here we show that three hnRNP proteins, PTB, hnRNPA1 and hnRNPA2, bind repressively to sequences flanking exon 9, resulting in exon 10 inclusion. We also demonstrate that the oncogenic transcription factor c-Myc upregulates transcription of

PTB/A1/A2, ensuring a high PKM2:PKM1 ratio. Establishing a relevance to cancer, we show that gliomas overexpress c-Myc and PTB/A1/A2 in a manner that correlates with

PKM2 expression. Our results thus define a pathway that regulates an alternative splicing event required for tumor cell proliferation.

Alternative splicing of PKM plays an important role in determining the metabolic phenotype of mammalian cells. The single exon difference imparts the enzymes produced with important functional distinctions. For example, PKM2, but not PKM1 is regulated by the binding of tyrosine phosphorylated peptides, which results in release of the allosteric activator fructose 1-6 bisphosphate and inhibition of PK activity5, a property that might allow growth-

factor initiated signaling cascades to channel glycolytic intermediates into biosynthetic processes.

1 99 The importance of tumor reversion to PKM2 was underscored by experiments in which

replacement of PKM2 with PKM1 in tumor cells resulted in markedly reduced growth2.

Consistent with a critical role in proliferation, re-expression of PKM2 in tumors is robust2, although little is known about the regulation of this process.

We set out to identify RNA binding proteins that might regulate PKM alternative splicing.

To this end, we prepared an [α-32P]-UTP labeled 250 nucleotide (nt) RNA spanning the E9 5’

splice site (EI9), previously identified as inhibitory to E9 inclusion6, as well as a labeled RNA

from a corresponding region of E10 (EI10) (Fig. 1b), and performed ultraviolet (UV)

crosslinking assays with HeLa nuclear extracts (NE)7. After separation by SDS-PAGE, multiple

proteins from 35-40 kDa appeared using the EI9 substrate, while little binding was observed

using the EI10 substrate (Fig. 1b). Strong binding was mapped to a 19 nt region we named

EI9(50-68) that spans the E9 5’ splice site (Supplementary Fig. 1). To identify the bound

proteins, we performed RNA affinity chromatography using a 5’ biotin-labeled RNA

corresponding to EI9(50-68). After SDS-PAGE and Coomassie staining, the pattern of

specifically bound proteins closely matched that observed after UV crosslinking (Fig. 1c). The

four indicated proteins between 35-40 kDa were excised, and identified by mass spectrometry as

isoforms of hnRNPA1 and hnRNPA2, RNA binding proteins with well established roles as

sequence-specific repressors of splicing (e.g., refs. 7, 8). This result was confirmed by

immunoblotting with antibodies against hnRNPA1 (Supplementary Fig. 2).

The sequence immediately downstream of the E9 5’ splice site contains a UAGGGC

sequence that is highly related to the consensus hnRNPA1 high affinity binding site identified by

9 SELEX, UAGGG(A/U) (Fig. 1d). Consistent with previous mutational studies of an identical

2 100 A1 binding site8, mutation of the G3 nucleotide of this motif to C led to a large decrease in

hnRNPA1/A2 binding (Fig. 1d; Supplementary Fig. 3). The G3C mutation resulted in increased

splicing in vitro when introduced into a splicing substrate containing E9 (Supplementary Fig. 4),

and led to increased E9 inclusion in a minigene construct in vivo (Supplementary Fig. 5). These

data confirm the presence of an inhibitory hnRNPA1/A2 binding site immediately downstream

of the E9 5’ splice site.

To explore the possibility that other splicing regulators bind upstream of E9 or E10, we

constructed crosslinking substrates (48 nt) that span the region upstream of each exon. Using

these RNAs for UV crosslinking showed strong binding of a 55 kDa protein to the I8 RNA probe,

but not to the I9 probe (Fig. 1e). Inspection of the polypyrimidine tract upstream of E9 revealed

two potential PTB (polypyrimidine tract binding protein, or hnRNPI) binding sequences

10 (UCUUC) within 35 nucleotides of the intron/exon boundary, while no such sequence exists in

the E10 polypyrimidine tract. PTB frequently functions as a splicing repressor10, often by

binding repressively to the polypyrimidine tract11. Immunoprecipitation confirmed that the 55

kDa crosslink observed using I8 RNA is PTB (Fig. 1e), and we observed strong binding of PTB

to a biotinylated version of I8 (Supplementary Fig. 6). In addition, mutation of the two putative

PTB binding sites from UCUUC to UGUUC significantly diminished binding (Fig. 1f;

Supplementary Fig. 7). Our data indicate that the splicing repressor PTB binds specifically to the

polypyrimidine tract of E9.

Because the locations of hnRNPA1/A2 and PTB binding sites flanking E9 overlap

elements critical to exon inclusion (the polypyrimidine tract for PTB11, the site of U1snRNA-pre-

mRNA base-pairing for A1/A212), we speculated that these proteins are inhibitors of E9

3 101 inclusion. To examine this possibility, we used siRNA to deplete hnRNPA1, hnRNPA2 and/or

PTB from HeLa cells. We assayed the PKM mRNA isoform ratio using RT-PCR followed by

exon-specific restriction digestion (Fig. 2a). Knockdown of hnRNPA1 or hnRNPA2 in HeLa

cells resulted in little change in splicing pattern (Supplementary Fig. 8). Because we have previously observed functional redundancy of hnRNPA1/A27, we next simultaneously depleted both proteins (Fig. 2b). This resulted in an increase in PKM1 mRNA, from 2% to 29%, and a concomitant decrease in PKM2 mRNA (Fig. 2c; Supplementary Fig. 8). PTB knockdown also increased the PKM1 isoform, to 16% (Fig. 2c; Supplementary Fig. 8), consistent with earlier

observations13. Next, we simultaneously depleted all three factors, which further increased

PKM1 levels, to about 48% (Fig. 2c). Similar results were obtained using 293 cells, with the

triple knockdown resulting in an increase from 5% to 67% PKM1 (Fig. 2d). Increases in PKM1

mRNA upon A1/A2/PTB knockdown were observed in all cell lines tested, including the breast

cancer cell line MCF-7 and the glioblastoma cell line U87 (Supplementary Fig. 9). Knockdown of two other cancer-associated splicing factors in HeLa cells, the SR proteins ASF/SF2 and

SRp20, while also resulting in slowed growth, failed to significantly affect PKM1/2 ratios

(Supplementary Fig. 8 and 10), indicating that the effects seen in PTB/A1/A2 depleted cells on

PKM splicing are specific and not the result of pleiotropic effects due to changes in cell growth.

Together, our results indicate that PTB/A1/A2 expression is the critical determinant of PKM isoform in transformed cells.

We next wished to determine whether PTB/A1/A2 expression levels and PKM1/2 alternative splicing are correlated. We first examined whether changes in PTB/A1/A2 levels correlate with changes in PKM splicing during switching from growth to quiescence. To this end, we used the mouse myoblast cell line C2C12, which, when grown to confluence and then

4 102 switched to low-serum medium, undergoes myogenic differentiation, a process that includes

PKM2 to PKM1 switching14. We differentiated C2C12 cells for 6 days, and used RT-PCR

followed by restriction digestion to assess the PKM1/2 ratio each day. We observed a large

increase in PKM1 and a corresponding decrease in PKM2 mRNA during differentiation (Fig. 3a).

We then prepared lysates of C2C12 cells at time points throughout differentiation and examined

protein levels by immunoblotting (Fig. 3b). PTB expression dropped over 70% by day 3 of

differentiation, after which it remained stable, consistent with previous studies15. We also

observed an approximately 50% decrease in hnRNPA1 levels by day 3 of differentiation, though no significant changes were observed in the level of hnRNPA2. This result is consistent with a

role for PTB/A1 in maintaining high PKM2 levels in proliferating C2C12 cells.

Because of the importance of the PKM2 isoform to the growth of cancer cells, we next

examined human glioma tumor samples for a correlation between PTB/A1/A2 expression and

PKM splicing. We first assayed PKM1/2 mRNA levels as described earlier. Normal brain tissue

ranged from 4-13% PKM2, pilocytic astrocytomas (PA) samples expressed approximately 66-

77% PKM2, low grade astrocytomas (LGA) ranged from 7-73%, and glioblastoma multiforme

(GBM) samples expressed 72-86% PKM2 (Fig. 3c). To explore a potential correlation between

elevated PKM2 mRNA levels and expression of the regulatory proteins we identified, we

performed immunoblots for PTB, hnRNPA1 and hnRNPA2. Significantly, all high-PKM2

tumors expressed elevated levels of PTB/A1/A2, with the most striking overexpression in GBMs

(Fig. 3d). Consistent with their uniformly high PKM2 expression, all four PA samples also showed overexpression of the PTB/A1/A2. In LGAs the two high PKM2 tumors showed elevated expression of the three proteins, while the two low PKM2 tumors showed expression levels similar to normal brain. Immunoblotting for four other splicing factors (ASF/SF2, Tra2β,

5 103 TLS/FUS, and hnRNPK) revealed no correlation with PKM2 expression (Supplementary Fig.

11), indicating that the correlation between an elevated PKM2/1 mRNA ratio and overexpression of PTB/ A1/ A2 is specific and not reflective of a general property of splicing factors.

The tight coupling of PKM2 expression to proliferation suggests that the expression of the PKM splicing regulatory proteins we identified might be under the control of a proliferation- associated regulatory mechanism. A strong candidate to control this is the oncogenic transcription factor c-Myc, which, like PTB/A1/A2, is upregulated in GBMs16, and has been shown to bind the PTB/A1/A2 promoters17, 18 and upregulate the expression of all three19, 20.

Consistent with a role for c-Myc in PTB/A1/A2 regulation, we observed a near perfect correlation between the levels of c-Myc and PTB/A1/A2 in gliomas and differentiating C2C12 cells (Fig. 3b and 3d). In addition, the transcription factor N-myc, which is closely related to c-

Myc21, was upregulated in PAs and to a lesser extent in GBMs (Supplementary Fig. 11),

indicating that this protein may in some cases contribute to PTB/A1/A2 upregulation.

We next examined directly c-Myc’s involvement in PTB/A1/A2 expression and PKM

splicing regulation. We first asked whether decreasing c-Myc levels can affect PTB/A1/A2

levels and the PKM1/PKM2 mRNA ratio. To this end, we transfected NIH-3T3 cells with

vectors bearing a puromycin resistance marker that express either a c-Myc-targeting shRNA or a

control shRNA. Immunoblotting showed a reduction in c-Myc levels in cells stably transfected

with c-Myc shRNA, compared to control cells (Fig. 4a). PTB/A1/A2 protein levels were also

significantly reduced after depletion of c-Myc, in contrast with two other RNA processing

factors not implicated in PKM splicing regulation, ASF/SF2 and CPSF73 (Fig. 4a). PTB/A1/A2

mRNA levels were also significantly reduced in the knockdown cells (Fig. 4b), supporting the

6 104 idea that c-Myc regulates transcription of these genes. Importantly, the cells stably expressing the c-Myc shRNA showed a pronounced increase in the PKM1/2 ratio, expressing 33% PKM1 mRNA compared to 7% in the control (Fig. 4c). A separate line stably expressing a second c-

Myc shRNA revealed a similarly elevated PKM1/2 ratio, as well as reduced levels of

PTB/A1/A2, showing that the observed effects were not due to off-target effects of the c-Myc shRNA (Supplementary Fig. 12). Additionally, we co-transfected an hnRNPA1 promoter- luciferase construct with a c-Myc expression vector22, which resulted in a dose- and c-Myc binding site- dependent increase in promoter activity (Supplementary Fig. 13).

The above results demonstrate a direct role for c-Myc in maintaining high PTB/A1/A2 levels in NIH-3T3 cells. In contrast, c-Myc knockdown in HeLa cells revealed only a small decrease in PTB/A1/A2 levels, and no change in the PKM1/2 ratio (Supplementary Fig. 14), suggesting that factors other than c-Myc might promote PTB/A1/A2 expression in these cells.

One possibility is the transcription factor E2F1, which like c-Myc binds upstream of all three genes18. However, knockdown of E2F1, or of Rb, a negative regulator of E2F family transcription factors23, resulted in little change in PTB/A1/A2 levels (unpublished data).

However, since the E2F and Rb families exhibit redundancy, this result does not rule out involvement of the E2F/Rb pathway in PTB/A1/A2 regulation. Indeed, because of their importance to proliferating cells, it is likely that PTB/A1/A2 can be upregulated by proliferation- associated factors in addition to c-Myc.

The fact that PTB/A1/A2 depletion results in switching to the PKM1 isoform suggests that RNA binding proteins can control the outcome of a mutually exclusive splicing event by simultaneously acting as repressors of one exon (E9) and activators of the other (E10)

7 105 (Fig. 4d). While it is easy to envision how these proteins exclude E9, how might PTB/A1/A2

promote E10 inclusion? A variety of RNA binding proteins, including hnRNPA1/A2, have been

shown to stimulate splicing of an adjacent exon through intronic binding sites24. One proposed

mechanism for this is intron definition, in which intron-binding proteins induce intronic

structures conducive to inclusion of the neighboring exon24. We propose that, like many

alternatively spliced exons, PKM E10 is poorly recognized by the splicing machinery in the

absence of adjacent intron definition, and such a structure is promoted by PTB/A1/A2 binding

(Fig. 4d).

We have demonstrated a critical functional consequence for observations connecting

PTB/A1/A2 upregulation with cell proliferation25, 26, transformation27, 28, and a wide variety of

cancers (e.g., refs. 26, 27, 29, 30). Given the critical role of these proteins in promoting PKM2

production in tumors, overexpression of some combination of them is, like PKM2 expression,

likely to be a general phenomenon in cancer. The fact that the proteins show some redundancy in

promoting PKM2 splicing may ensure robust re-expression of PKM2 in tumors.

Methods summary

UV crosslinking substrates were cloned into pcDNA3 vector (Invitrogen) and UV

crosslinking was performed as previously described7. Mutations were introduced in EI9 by PCR-

based site-directed mutagenesis. Biotinylated RNAs for affinity purification were purchased

from Dharmacon, and RNA affinity chromatography was carried out as described7.

Immunoprecipitations were carried out using protein A-agarose beads (Roche). RNAi was performed as described7. We transfected 50 pmol of hnRNPA1 siRNA and 25 pmol of other

siRNA duplex in a 24-well plate. After 72 hours, we collected cells for RNA isolation and

8 106 immunoblotting. C2C12 cells were grown in DMEM (Invitrogen) supplemented with 20% fetal bovine serum (FBS) (Hyclone) at 37°C in 5% CO2. For differentiation treatment, C2C12 were plated on gelatin coated plates, allowed to reach confluence, and then switched to DMEM 2% donor equine serum (Hyclone). Human brain and glioma samples were obtained from the Bartoli

Brain Tumor Bank at the Columbia University Medical Center. Samples were homogenized and used for Trizol RNA extraction and western blotting as described30. In all cases, immunoblots were scanned and quantified using the LI-COR Odyssey system. c-Myc shRNA DNA sequences were purchased from Invitrogen and cloned into the pRS vector (Origene). shRNA constructs were transfected into NIH3T3 cells and stable cell lines were selected with puromycin for RNA isolation and immunoblotting. PKM1/PKM2 ratio was analyzed by extracting total RNA from cells and tissue samples and preforming by RT-PCR followed by PstI, Tth111I, or EcoNI digestion. qPCR for PTB/A1/A2 in control and c-Myc knockdown cells was performed with

SYBR green from Fermentas using the Applied Biosystems 7300 real-time PCR system. hnRNPA1 promoter sequence for dual luciferease reporter (DLR) assay was cloned into PGL3- enhancer vector (Promega) and DLR assays were performed using Dual Luciferase Reporter

Assay System (Promega)

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Acknowledgements We thank D. Black for BB7 antibody, T. Kashima for hnRNP A1/A2

siRNA, R. Prywes and E. Henckles for C2C12 cells, M. Sheetz and X. Zhang for NIH-3T3 cells,

12 110 C. Prives, T. Barsotti, and L. Biderman for MCF-7 cells, Rb, and E2F-1 siRNA, R. Dalla-Favera and Q. Shen for anti-c-Myc antibodies and c-Myc expression vector, R. Eisenmann for N-myc antibody, and members of the Manley laboratory for discussions. This work was supported by grants from the Avon Foundation and the NIH.

Author contributions C.J.D., M.C., and J.L.M. conceived the project and designed experiments,

C.J.D. and M.C. carried out experiments, P.C. and M.A. provided tumor samples, C.J.D., M.C. and J.L.M. interpreted data and wrote the paper.

Figure legends

Figure 1: hnRNP proteins bind specifically to sequences flanking E9. a, Schematic diagram of PKM splicing. b, Position of probes spanning the E9 or E10 5’ splice sites (top). After UV crosslinking, proteins were detected by autoradiography (bottom). c, Affinity chromatography using EI9(50-68). Bound proteins were separated by SDS-PAGE and Coomassie stained. Bands excised for mass spectrometry are indicated. d, Sequence of EI9(50-68), putative hnRNPA1/A2 binding site indicated in bold italics (top). UV crosslinking with wild-type RNA, or RNA with a mutation in the putative hnRNPA1/A2 binding site (bottom). e, Position of I8 and I9 (top). UV crosslinking using I8 or I9 substrates (bottom left). UV crosslinking reactions were IPed with either α-PTB (BB7) or α-HA antibodies (bottom right). f, UV crosslinking with I8 and mutant derivative I8mu, sequences indicated above. Putative PTB binding sites in I8 are underlined.

Figure 2: PTB, hnRNPA1, and hnRNPA2 are required for high PKM2:PKM1 mRNA ratios. a, Scheme for assaying PKM1/PKM2 ratios in human cells. b, Immunoblots showing protein levels after the indicated siRNA treatment. Protein bands were quantified after LI-COR

13 111 Odyssey scanning and normalized to GAPDH. c, The indicated splicing factors were depleted by siRNA, followed by PKM splicing assay outlined in (a). Products corresponding to M1 and M2 are indicated with arrows. The PKM1 percentage is indicated below. d, PKM1/2 levels assayed after the indicated siRNA treatment in 293 cells.

Figure 3: Expression of PTB/A1/A2 and c-Myc correlates with PKM2 expression in C2C12 cells and tumors. a, PKM splicing assay after the indicated number of days of C2C12

differentiation. b, Immunoblots for the indicated proteins were performed throughout

differentiation, and normalized to GAPDH (Day 0 = 1). c, RNA was extracted from brain tissue or tumor samples and assayed for PKM mRNA isoforms. d, Lysates were immunoblotted for

PTB, hnRNPA1, hnRNPA2 or c-Myc and normalized to Actin. Sample order is the same for

RT-PCR and immunoblotting.

Figure 4: c-Myc upregulates PTB/A1/A2 alters PKM splicing. a, Immunoblotting using

NIH-3T3 cells stably expressing control or c-Myc-targeting shRNAs. Signals were quantified and normalized to actin. b, RT-PCR using the same cell lines as in (a). Realtime RT-PCR was performed separately to quantify the relative levels of PTB/A1/A2 mRNAs in control and c-Myc

knockdown cells, using RPL13A as a reference gene. Relative levels of each are shown below

each panel, with s.d. indicated (n=3). c, PKM1/2 ratios in control and c-Myc knockdown cells

determined as in Fig. 2a., d, A model for PKM splicing regulation. Top, in adult tissues, low

expression of PTB/A1/A2 allows for recognition of E9 by the splicing machinery and disrupts intronic structures favorable for E10 inclusion. Bottom, in embryonic and cancer cells,

PTB/A1/A2 are upregulated, bind to splicing signals flanking E9 and repress its inclusion.

14 112 Binding of these proteins around E9 and possibly to other sites creates an intronic structure

favorable to E10 inclusion.

Methods

Plasmid constructs. Long UV crosslinking substrates (EI9, EI10) were prepared by amplifying

fragments from HeLa genomic DNA using Pfu turbo (Stratagene), and cloning the products into

pcDNA3 (Invitrogen). EI9(1-20), EI9(21-49), and EI9(50-68, I8, I8mu, and I9 DNA sequences

were ordered from Invitrogen and cloned into pcDNA3. Primers used to amplify genomic DNA fragments were: EI9 forward, CGC GGA TCC TTC TTA TAA GTG TTT AGC AGC AGC T , reverse, CGG AAT TCA CTG AGC CAC AGG ACC CTT TG ; EI10 forward, CGC GGA TCC

CTC CTT CAA GTG CTG CAG TG , reverse, CGG AAT CCT GGG CCC AGG GAA GGG G;

I8E9 forward, CCC AAG CTT AAA TTC CCC ATT CTG TCT TCC CAT G , reverse, CGG

GAT CCC TGC CAG ACT CCG TCA GAA CT; I9E10 forward, CCC AAG CTT CTG TCC

GGT GAC TCT TCC CC , reverse, CGG GAT CCC TGC CAG ACT TGG TGA GGA CG.

Mutations were introduced in EI9 by PCR-based site-directed mutagenesis. Mouse c-Myc and control shRNA DNA sequences were ordered from Invitrogen and cloned into pRS vector

(Origene) with BamH I and Hind III. The hnRNPA1 promoter region, either wild-type or the E box mutant, was cloned into PGL3-enhancer vector (Promega).

Antibodies. The following antibodies were used in this study: BB7 for human PTB IP (gift from

Dr. Douglas Black, UCLA), 3H8 for mouse/human PTB immunoblots (Sigma), MC3 for

15 113 U2AF65 (Sigma), α-HA (Covance) DP3D3 for hnRNPA2 (Abcam), N-262 for c-Myc (Santa

Cruz), α-Actin (Sigma), α-GAPDH (Sigma), 9H10 for hnRNPA1 (Sigma), mAb104 for SRp20.

UV crosslinking, RNA affinity purification, and immunoprecipitation assays. We carried out

ultraviolet crosslinking as previously described7. Briefly, we linearized the UV crosslinking

plasmids with an appropriate restriction enzyme and synthesized the RNAs with [32p]-UTP or

[32p]-CTP. We incubated 1X105 c.p.m. RNAs with 10 μg HeLa or C2C12 NE in buffer D in a 20

μl reaction at 30°C for 15 minutes, then irradiated the samples with ultraviolet light in a

Stratalinker 1800 (Stratagene), digested them with RNase A(10µg/ml) and resolved them by

SDS-PAGE. The RNA affinity pull-down experiment and immunoprecipitation was preformed as described7. The 5’biotinylated EI9(50-68) and I8 RNA oligonucleotides were purchased from

Dharmacon. Antibodies were bound to protein A-agarose beads prior to IP. We used the following antibodies for IP: BB7 for PTB, and MC3 for U2AF65.

In vitro and in vivo splicing assays. Minigene containing PKM gene exon 8, exon 9, exon 10 , exon 11 and flanking regions was cloned into pcDNA3 vector (Invitrogen). G to C mutation was introduced in Minigene by PCR-based site-directed mutagenesis11. Wildtype and mutated

minigene vectors were transfected into HeLa cells. 24 hours after transfection, cells were

collected and PKM1/M2 ratio was analyzed using RT-PCR followed by Pst I digestion. In vitro

splicing substrates were constructed by replacing the first exon and downstream intronic

sequence of AdML pre-mRNA with PKM exon 9 and downstream intron 9 sequences or

sequence with mutated hnRNP A1 binding site. pre-mRNA substrates were synthesized by in

vitro transcription using T7 RNA polymerase (Promega) following product protocol. In vitro

16 114 splicing of the wildtype and mutated pre-mRNA was carried out using HeLa nuclear extract as

described31.

RNA interference. We carried out RNA interference of PTB and hnRNPA1/A2 as described7.

Briefly, we plated HeLa, 293, MCF-7, or U87 cells at 2.5-3x104 cells per well in 24-well plates.

The next day, we mixed 50 pmole of hnRNPA1 duplex RNA and 25 pmole of the other duplex

RNAs with 1.5 μl lipofectamine 2000 transfection reagent (Invitrogen) plus 100 μl of Opt-MEM

medium and added this to cells after RNA duplex-lipid complex formation. For double and triple

knockdowns in HeLa and 293 cells, RNA duplexes were transfected simultaneously. The control

RNA duplex was used to ensure that parallel experiments had equal amounts of RNA. In MCF-7

and U87 cells, the second and third RNA duplexes were transfected 6 hours after the previous

transfection. 72 hours after transfection, we collected cells for RNA isolation and

immunoblotting. We used the following siRNAs (Dharmacon; the sense strand sequences are

given): human hnRNPA1, CAGCUGAGGAAGCUCUUCA; human hnRNPA2,

GGAACAGUUCCGUAAGCUC; human PTB, GCCUCAACGUCAAGUACAA. ASF/SF2

depletion was performed as previously described7.

c-Myc shRNA stable cell lines. Stable cell lines expressing c-Myc shRNAs or control shRNA

were obatined by transfecting pRS-shRNA vectors into NIH3T3 cells followed by drug selection.

Cells were plated in 10cm plates. The next day, transfected cells were diluted and medium was

replaced with medium containing a final concentration 3 μg/ml puromycin. After 7-10 days, a

mixture of fast- and slow-growing colonies appeared in cells transfected with c-Myc shRNA,

while only fast-growing colonies appeared in cells transfected with control shRNA. Single slow-

growing colonies were isolated and cultured for c-Myc expressing cells. c-Myc expression was

17 115 examined by immunoblotting. Positive colonies were collected for RT-PCR and western blotting.

The following sense shRNA sequences were used:

control, gat ccG AGG CTT CTT ATA AGT GTT TAC TCG AGT AAA CAC TTA TAA GAA

GCC TCT TTT Ta ; Mouse c-Myc shRNA1, gat ccC ATC CTA TGT TGC GGT CGC TAC

TCG AGT AGC GAC CGC AAC ATA GGA TGT TTT Ta; Mouse c-Myc shRNA2, gat ccC

GGA CAC ACA ACG TCT TGG AAC TCG AGT TCC AAG ACG TTG TGT GTC CGT TTT

Ta; human c-Myc shRNA, gat ccC CAT AAT GTA AAC TGC CTC AAC TCG AGT TGA

GGC AGT TTA CAT TAT GGT TTT Ta.

RT-PCR. Total RNA was extracted from tissue culture and human brain tumor samples using

Trizol (Invitrogen) according to the manufacturer’s instructions. Total RNA (2.5–5 μg) was used

for each sample in a 20 μl reaction with 0.5μL of SuperScript III RT (Invitrogen). 1μl of the

cDNA library was used in a 50 μl PCR reaction containing 3 μCi [32p]-dCTP. 10μl of the PCR

products were digested by Pst I and Tth111 I (human PKM) or EcoN I (mouse PKM) and the

products were resolved by 6% non-denaturing PAGE. Primers used in the PCR reactions were:

human PKM exon8 forward, CTG AAG GCA GTG ATG TGG CC; human PKM exon11 reverse, ACC CGG AGG TCC ACG TCC TC; mouse PKM exon 8 forward, CAA GGG GAC

TAC CCT CTG G; mouse PKM exon11 reverse, ACA CGA AGG TCG ACA TCC TC, human

µglobulin: forward, GGC TAT CCA GCG TAC TCC AAA, reverse, CGG CAG GCA TAC

TCA TCT TTT T; mouse µglobulin: forward, TTC TGG TGC TTG TCT CAC TGA, reverse,

CAG TAT GTT CGG CTT CCC ATT C. qRT-PCR was performed using the following primers:

mouse hnRNPA1: forward, TGG AAG CAA TTT TGG AGG TGG, reverse, GGT TCC GTG

GTT TAG CAA AGT; mouse hnRNPA2: forward, AAG AAA TGC AGG AAG TCC AAA GT,

18 116 reverse, CTC CTC CAT AAC CAG GGC TAC; mouse PTB: forward, AGC AGA GAC TAC

ACT CGA CCT, reverse, GCT CCT GCA TAC GGA GAG G; mouse RPL13A forward, GGG

CAG GTT CTG GTA TTG GAT, reverse, GGC TCG GAA ATG GTA GGG G. Relative

amounts of mRNA were calculated using the comparative Ct method.

Cell culture and differentiation. C2C12 cells were grown in DMEM (Invitrogen) supplemented

with 20% fetal bovine serum (FBS) (Hyclone) at 37°C in 5% CO2. For differentiation treatment,

C2C12 were plated on gelatin coated plates, allowed to reach confluence, and then switched to

DMEM 2% donor equine serum (Hyclone). HeLa and 293 cells were grown in DMEM, 10%

FBS. NIH3T3 cells were grown in DMEM, 10% bovine calf serum (BCS) (Hyclone).

Human brain tumor samples. De-identified brain and glioma samples were obtained from the

Bartoli Brain Tumor Bank at the Columbia University Medical Center. Non-cancerous samples removed from epileptic patients were used for normal brain. Approximately 25-200 mg of each sample was obtained. Half of the homogenate was used for Trizol RNA extraction, the other half of each sample was processed for immunoblotting as described30.

Dual Luciferase Reporter (DLR) Assay. c-Myc expression vector and hnRNP A1 promoter vector were co-transfected into HeLa cells. 24 hours after transfection, cells were collected and

DLR assays were preformed using Dual Luciferase Reporter Assay System (Promega) following product protocol.

31 Krainer, A.R., Maniatis, T., Ruskin, B., & Green, M.R., Normal and mutant human beta- globin pre-mRNAs are faithfully and efficiently spliced in vitro. Cell 36, 993-1005 (1984)

19 117 118 119 120 121

Chapter 4: RNA polymerase II and pre-mRNA splicing.

122 Chapter 4: RNA polymerase II and pre-mRNA splicing.

Of the major pre-mRNA processing reactions, which include capping, splicing, and cleavage/polyadenylation, splicing is certainly the most complex. Splicing is carried out by a

large complex known as the spliceosome, which consists of at least 150 protein components, as

well as 5 snRNAs (Smith et al. 2008; Wahl et al. 2009; Valadkhan and Jaladat 2010). The

spliceosome is a highly dynamic entity that undergoes sequential compositional/conformational

changes between the initial recognition of pre-mRNA splice sites and the eventual excision of

the intron. In vitro, this process can occur faithfully in the absence of transcription. In contrast, in

vivo, transcription by RNA polymerase II is required for the efficient intron removal (Sisodia et

al. 1987). This observation implies that there is an indispensable functional communication

between the transcription and splicing machineries, a phenomenon known as coupling. After a

brief discussion of spliceosome assembly, this chapter will discuss our current knowledge of the

molecular connections between splicing and transcription in mammalian cells.

The spliceosome assembly pathway

In vitro systems have been useful in elucidating the highly complex sequence of events

that define the process of mammalian spliceosome assembly (Smith et al. 2008; Wahl et al.

2009). During in vitro splicing, a series of stable intermediate complexes form on the pre-mRNA

substrate. These complexes are amenable to biochemical analysis, and thus provide “snapshots”

of the assembly pathway. The spliceosomal complexes, termed E, A, B, and C (in order of

formation) can be separated by native gel electrophoresis, a technique that allowed the initial characterization of this process (Konarska and Sharp 1986). More recently, affinity purification approaches coupled with proteomic analysis has provided significant insight into the 123 compositional remodeling that occurs along the spliceosome assembly pathway (Makarov et al.

2002; Behzadnia et al. 2007).

Spliceosome assembly begins with the formation of E complex, which can form in the

absence of ATP. The initial recognition of splice sites on mammalian pre-mRNAs is a highly

cooperative process, usually involving the physical association of splicing factors whose

individual affinities for pre-mRNA is low (Wahl et al. 2009). In E complex, the 5’ splice site is recognized by U1 snRNP, the binding of which requires the assistance of exon-bound SR proteins (Michaud and Reed 1991; Michaud and Reed 1993; Kohtz et al. 1994). SR proteins are a class of essential splicing factor characterized by an N-terminal RNA binding domain, and a C- terminal arginine-serine rich domain (Shepard and Hertel 2009). SR proteins also play a role in the early recognition of the 3’ splice site, by binding to and recruiting the dimeric splicing factor

U2AF (Wu and Maniatis 1993). An additional important feature of early stages in splice site recognition is crosstalk between splice sites flanking an individual exon. This phenomenon, known as exon definition, involves communication between factors that recognize the 5’ and 3’ splice sites, resulting in the cooperative recognition of the exon (Berget 1995). In mammals, exons are usually dwarfed by the introns that surround them, so it appears likely that splice sites are first recognized cooperatively through cross-exon interactions, although recognition in this manner by no means guarantees the eventual inclusion of the exon in the spliced mRNA (House and Lynch 2006).

Formation of the next intermediate in spliceosome assembly, A complex, requires ATP hydrolysis, and results in the stable recruitment of additional factors to the pre-mRNA, most notably the U2 snRNP. The U2 snRNA basepairs with the branchpoint sequence at the 3’ splice site, assisted by the proteins of the associated SF3a/b complexes as well as U2AF65 (Gozani et 124 al. 1996; Valcarcel et al. 1996). Proteomic analysis has shown that the Prp19 complex (PRP19C) is also associated with the spliceosome in A complex. Following A complex formation, the

U4/U6 and U5 snRNPs become associated with the spliceosome as part of a pre-assembled unit known as the tri-snRNP, forming the intermediate known as B complex. Additional rearrangements, notably ejection of two snRNA components, U1 and U4, precedes the formation of the activated B complex (B*). This is followed by the execution of the first catalytic step, which generates C complex. The transition from the B to C complex involves the destabilization of the SF3a/b proteins, while PRP19C becomes more stably integrated in the catalytic core of the spliceosome at this stage (Bessonov et al. 2008). Further rearrangements are necessary for the second catalytic step, and spliceosome disassembly (Wahl et al. 2009).

Evidence for coupling of transcription and splicing

Early electron microscopy studies showed strikingly that spliceosome assembly and intron removal occurs rapidly after the transcription of introns in vivo (Beyer and Osheim 1988).

This on its own does not necessarily imply that transcription and splicing are functionally coupled. However, around the same time, it was shown that transcription by RNAPII is required for proper processing of pre-mRNA (Sisodia et al. 1987), indicating that RNAPII itself plays a pivotal role in ensuring the rapid processing of its transcripts. A consideration of some important differences between in vitro splicing and the removal of introns in vivo underscores the importance of such coupling. For one, introns in mammalian cells can be extremely long, often exceeding 100,000 nucleotides (nt). A recent study by Singh et al., showed that there is no correlation between the length of an intron and the time required for its excision (Singh and

Padgett 2009). These investigators showed that introns spanning 1,500 to well over 100,000 nt were all spliced within 5-10 minutes of the transcription of the downstream exon. In contrast, in 125 vitro splicing rarely occurs efficiently on substrates longer than 500 nt, with a lag time of 20-30 minutes.

How does a cell ensure the accurate and swift pairing of splice sites of a 100,000 nt intron?

One appealing possibility is that exons are recognized co-transcriptionally (through exon definition, discussed above), and through multiple interactions between RNA-bound splicing factors and the RNAPII elongation complex, the exon becomes “tethered” to the elongating polymerase, which then scans the nascent RNA for an appropriate acceptor site. Direct evidence for the tethering hypothesis was provided by Proudfoot and colleagues, who showed that the presence of a self-cleaving hammerhead ribozyme placed within an intron was insufficient to disrupt splicing in a β-globin derived minigene (Dye et al. 2006). It should be noted that the validity of the conclusions drawn from the work of Dye et al. have been more recently called into question by Bentley and colleagues, who showed that use of the faster-cleaving hepatitis delta ribozyme in the same position was in fact sufficient to disrupt splicing (Fong et al. 2009).

To resolve the controversy, it will be important to test the hypothesis using ribozymes situated in longer introns in constructs stably integrated in the genome.

Other observations also seem to support functional links between transcription and splicing. Kornblihtt and colleagues have provided a wealth of data showing that alternate promoter usage can result in dramatic changes in the regulation of alternative splicing

(Kornblihtt 2005). This phenomenon may in some cases be explained in ways that do not require physical interactions between the splicing and transcription machineries. For example, it has

been shown that alterations in the elongation rate of RNAPII, which can be affected by promoter choice, can affect the inclusion of alternatively spliced exons (de la Mata et al. 2003). While some of the promoter effects on splicing may be explained through alterations in RNAPII 126 kinetics, additional work has shown that promoter-bound transcription factors can affect pre- mRNA splicing by directly altering the composition of the transcription elongation complex

(TEC). In one example of this, it was shown that a strong transcriptional activator, VP16 can recruit splicing factors to the promoter (Rosonina et al. 2005). These splicing factors then join the TEC through interactions that will be discussed below, and enhance the efficiency of splicing of the transcribed pre-mRNA. In addition, some of the effects of promoters on pre-mRNA splicing are mediated by specialized proteins that serve as dual transcription/splicing factors. For example, the CAPERα/β proteins, both of which are highly homologous to the essential splicing factor U2AF65, have been shown capable of both transactivating transcription and regulating the splicing of the resulting transcript (Dowhan et al. 2005). In conclusion, the data described here point to extensive connections between the transcriptional machinery and splicing factors being essential to the execution of constitutive splicing, as well as for the regulation of alternative splicing.

The RNAPII CTD: a critical mediator of transcription/splicing coupling

Our understanding of transcription/splicing coupling reached another milestone when it was shown that truncation of the RNAPII C-terminal domain (CTD) severely impaired the splicing of a transiently transfected minigene transcript in vivo (McCracken et al. 1997). CTD truncation also impaired the other two principal RNA processing events, capping and polyadenylation, both of which have been shown to influence splicing efficiency (Konarska et al.

1984; Niwa et al. 1990). However, Bentley and colleagues went on to show that each event was independently impaired by CTD truncation, providing strong evidence that the CTD can directly stimulate pre-mRNA splicing (Fong and Bentley 2001). Direct evidence was soon provided for this in vitro, when Hirose et al. showed that RNAPII with a phosphorylated CTD was capable of 127 increasing the kinetics of in vitro splicing (Hirose et al. 1999). This observation led to

speculation that the phosphorylated RNAPII CTD directly enhances spliceosome assembly by

acting as a scaffold that binds multiple splicing factors (see below).

CTD structure and modification

The CTD of RNAPII in mammals is composed of 52 heptad repeats of the consensus

sequence YSPTSPS. Heptads in the N-terminal half of the CTD mostly conform to the consensus

sequence, while heptads in the C-terminal half are much more degenerate. The degenerate and

consensus ends of the CTD are functionally indistinguishable in assays for splicing and

polyadenylation (Rosonina and Blencowe 2004). To date, the only function identified that is

unique to either half is RNA binding by the C-terminal portion, a function that may have

implications in the regulation of poly(A) site choice (Kaneko and Manley 2005).

The CTD is subject to extensive phosphorylation, which has major impacts on its

function. RNAPII recruited to promoters is hypophosphorylated on its CTD. Upon

transcriptional initiation, the CTD is phosphorylated by the cdk7 subunit of TFIIH at serine 5 of

the heptad repeat (S5). This phosphorylation event is necessary for promoter clearance (Sogaard and Svejstrup 2007). S5 phosphorylation levels peak near the 5’ ends of genes, then decline as

RNAPII moves towards the 3’ end (Komarnitsky et al. 2000; Schroeder et al. 2000). The other major phosphorylation event occurs on serine 2 (S2), which in contrast to S5, increases towards the 3’ ends of genes (Komarnitsky et al. 2000). A complex interplay of S5 or S2 specific kinases and phosphatases determines the pattern of CTD phosphorylation along each gene (Phatnani and

Greenleaf 2006; Buratowski 2009). 128 CTD phosphorylation appears to play an important role in coupling splicing and

transcription. In vitro, CTD phosphorylation has been shown to be essential for the stimulatory effect of the CTD on splicing (Hirose et al. 1999; Millhouse and Manley 2005). Interestingly, the

coupling of transcription and splicing through CTD phosphorylation has emerged as regulatory

point in the control of gene expression. Medzhitov and colleagues recently described a set of

inducible genes, which, when in the “off” state, are actively transcribed by RNAPII

phosphorylated on S5 but not S2 (Hargreaves et al. 2009). The transcripts that result from

transcription by the S5-phosphorylated RNAPII, while full length, remain unspliced. When the

gene is induced, PTEF-b, a S2-specific CTD kinase is recruited to the gene, resulting in proper

splicing of the transcribed RNA. In addition to its repercussions for the gene regulation, this

important finding directly implicates S2 phosphorylation as essential to the integration of

splicing and transcription.

Physical links between the CTD and splicing

What are the physical links between the phosphorylated RNAPII CTD and the splicing

apparatus? A number of splicing factors have been shown to bind to the CTD. The S. cerevisiae

protein Prp40, which is associated with U1 snRNP, binds to phosphorylated CTD repeats

through its WW domain (Hargreaves et al. 2009). Phospho-CTD binding appears to be a frequent

function of WW domains, and also of the related FF domain, which are present on a number of

splicing factors. These domains are found on multiple splicing factors, and in addition to

interactions with the CTD also mediate interactions with other splicing factors. For example, the

splicing factor CA150 interacts with the phosphorylated CTD through FF domains (Carty et al.

2000), while interacting with other splicing factors, such as SF1 through its WW domains

(Goldstrohm et al. 2001). Based on these observations, an appealing hypothesis is that proteins 129 such as CA150 provide a link between the CTD and factors that play a central role in

spliceosome assembly, although direct evidence to support this idea is still lacking. In addition,

the splicing factors PSF and p54nrb, which interact to form a stable heterodimer, were shown to

bind directly to the CTD (Emili et al. 2002). Interestingly, these proteins bound to the CTD

regardless of its phosphorylation status. Binding of PSF to the CTD has provided the only

demonstration to date of a functional significance for an interaction between a splicing factor and

the CTD. Rosonina et al., showed that PSF, which is recruited to promoters by the strong

transcriptional activator VP16, can promote enhanced splicing of transcripts from promoters

harboring binding sites for the VP16 (Rosonina et al. 2005). This effect is abolished when

transcription is carried out by RNAPII bearing a truncated CTD, indicating that PSF is recruited

to promoters then travels along the gene with RNAPII through interactions with the CTD.

While a few splicing factors have been shown to bind to the CTD, demonstrated

functional connections between the RNAPII CTD and splicing factors remain few. In Chapter 5,

I describe an approach, initiated by Scott Millhouse (Millhouse and Manley 2005), that has

allowed for the identification of a splicing factor that interacts functionally with the CTD of

RNAPII. The study described in Chapter 5 also resulted in the identification of an interaction between two central players in spliceosome assembly, providing insight into that process, and its connections to transcription through the CTD.

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133

Chapter 5: The RNA polymerase II C-terminal domain promotes

splicing activation through recruitment of a U2AF65-Prp19 complex

134

The RNA polymerase II C-terminal domain promotes splicing activation

through recruitment of a U2AF65-Prp19 complex

Charles J. David, Alex R. Boyne, Scott R. Millhouse

and James L. Manley

Columbia University

Department of Biological Sciences

New York, NY 10027 135 ABSTRACT

Pre-mRNA splicing is frequently coupled to transcription by RNA polymerase II

(RNAPII). This coupling requires the C-terminal domain of the RNAPII largest subunit (CTD), although the underlying mechanism is poorly understood. Using a biochemical complementation assay, we previously identified an activity that stimulates CTD-dependent splicing in vitro. We have now purified this activity, and find that it consists of a complex of two well-known splicing factors, U2AF65 and the Prp19 complex (PRP19C). We provide evidence that both U2AF65 and

PRP19C are required for CTD-dependent splicing activation, that U2AF65 and PRP19C interact both in vitro and in vivo, and that this interaction is required for activation of splicing. Providing the link to the CTD, we show that U2AF65 binds directly to the phosphorylated CTD, and that this interaction results in increased recruitment of U2AF65 and PRP19C to the pre-mRNA. Our results not only provide a mechanism by which the CTD enhances splicing, but also describe unexpected interactions important for splicing and its coupling to transcription.

1 136 INTRODUCTION

Almost all mammalian RNAPII transcripts undergo three principal processing events

before their export from the nucleus: capping, splicing and polyadenylation. These events are frequently coupled to transcription by RNAPII in a manner that ensures the faithful and efficient execution of each step (Hirose and Manley 2000; Pandit et al. 2008; Perales and Bentley 2009).

The repetitive C-terminal domain of the large subunit of RNAPII (CTD) has been shown to play a central role in coupling transcription to all three of the main processing reactions (Phatnani and

Greenleaf 2006; Perales and Bentley 2009; Munoz et al. 2010). Deletion of most of the CTD can result in inefficient capping, splicing and polyadenylation in vivo (McCracken et al. 1997a; Fong and Bentley 2001). Consistent with this, the stimulatory effect of the CTD on 3’ cleavage and splicing can be recapitulated in vitro (Hirose and Manley 1998; Hirose et al. 1999; Zeng and

Berget 2000). In mammals, the CTD consists of 52 heptad repeats of the consensus sequence

YSPTSPS. Multiple residues within the CTD heptad are phosphorylated throughout the transcription cycle (Egloff and Murphy 2008; Buratowski 2009). Phosphorylation of serine 5 (S5) of the heptad is most prominent at the 5’ end of genes (Komarnitsky et al. 2000; Schroeder et al.

2000)，while serine 2 (S2) phosphorylation increases towards the 3’ end (Komarnitsky et al.

2000). CTD phosphorylation plays an important role in generating elongation-competent

RNAPII (Sims et al. 2004), and is required for the stimulatory effect of the CTD on splicing

(Hirose et al. 1999; Millhouse and Manley 2005).

A number of interactions linking pre-mRNA processing and the CTD have been documented. For capping, the functional connection with the CTD is straightforward; the guanylytransferase and methyltransferase enzymes necessary for capping both bind to the S5- phosphorylated CTD, which allosterically activates guanylytransferase activity (McCracken et al.

1997b; Yue et al. 1997; Ho and Shuman 1999). Connections between the polyadenylation machinery and the CTD have also been demonstrated. Human CstF50 was shown to physically interact with both the phosphorylated and unphosphorylated CTD, an interaction that appears

2 137 important for efficient cleavage/polyadenylation in vivo (Fong and Bentley 2001). The yeast CFI

subunit Pcf11 also interacts with the S2-phosphorylated CTD, the functional importance of which was suggested by a genetic interaction between a Pcf11 allele and an RBP1 CTD truncation allele (Licatalosi et al. 2002). Also, a CTD phosphatase, Ssu72, was recently shown to be important for transcription-coupled 3’ processing in vitro (Xiang et al. 2010).

The machinery that carries out pre-mRNA splicing is considerably more complex than those responsible for capping and polyadenlyation. The spliceosome, the protein-RNA assembly that catalyzes intron removal, contains at least 150 proteins, and undergoes dynamic changes in conformation and protein composition during the series of events that begin with splice site recognition and end after the execution of the two catalytic steps (Jurica and Moore 2003; Smith et al. 2008; Wahl et al. 2009; Valadkhan and Jaladat 2010). In vitro, spliceosome assembly proceeds through the formation of a series of stable intermediate complexes, which are biochemically separable and amenable to proteomic analysis (Wahl et al. 2009). Among the earliest steps in spliceosome assembly is recognition of the 5’ and 3’ splice sites by the U1 snRNP and U2AF, respectively. U2AF is a dimer comprised of U2AF65 and U2AF35 (Zamore and Green 1989). U2AF65 binds to polypyrimidine-rich sequences found near the 3’ end of most introns, and promotes stable U2 snRNP association with the pre-mRNA, an activity that requires its N-terminal arginine-serine rich (RS) domain (Valcarcel et al. 1996). U2AF35 contacts a well- conserved AG dinucleotide at the 3’ end of the intron (e.g. Wu et al. 1999), and can interact with exon-bound SR proteins; both interactions can stabilize U2AF binding to suboptimal polypyrimidine tracts (Zuo and Maniatis 1996). Later steps in spliceosome assembly involve the activity of numerous additional factors, including the U4/U6.U5 tri-snRNP and the Prp19 complex, or PRP19C (Wahl et al. 2009). PRP19C, which consists of four polypeptides that form a salt-stable core (CDC5L, PRLG1, Prp19, SPF27), and three more loosely associated polypeptides (HSP73, CTNNBL1, and AD002) (Grote et al. 2010), is found at the core of catalytically activated spliceosomes, and plays a critical but poorly understood role in activation

3 138 of the spliceosome (Chan et al. 2003; Bessonov et al. 2008; Song et al. 2010). Because PRP19C does not contain any proteins known to bind RNA, it is likely that PRP19C recruitment to the spliceosome occurs through protein-protein interactions with RNA-bound factors, although no such interaction has yet been described.

Most of what is known about the process of spliceosome assembly has come from the use of in vitro systems that are uncoupled from transcription, leaving the role of the transcriptional machinery in the process relatively poorly understood. However, a few physical interactions between splicing factors and the CTD have been documented. The yeast U1 snRNP component

Prp40 was shown to bind to the phosphorylated CTD through multiple WW domains (Morris and

Greenleaf 2000; Gasch et al. 2006). In humans, splicing factors that have been shown to bind directly to the CTD include CA150 (Carty et al. 2000), PSF and p54/NRB (Emili et al. 2002). Of these, support for a functional significance to the CTD interaction has only been provided for

PSF, which can be recruited to promoters by strong transcriptional activators to promote splicing in a CTD-dependent manner in vivo (Rosonina et al. 2005).

In order to study the functional connections between the CTD and pre-mRNA splicing, we previously constructed a fusion between the CTD and the SR protein SRSF1 (formerly

ASF/SF2). This allowed recruitment of the CTD to splicing substrates harboring SRSF1 binding sites independent of transcription. Using this fusion protein, which we now call SRSF1-CTD, in in vitro splicing assays, we observed an increase in splicing kinetics in its presence when compared to SRSF1 alone (Millhouse and Manley 2005). In addition, we found that a HeLa nuclear fraction (NF20-40) was capable of activating splicing in HeLa nuclear extract (NE) of one substrate, IgMA3, in the presence of SRSF1-CTD but not SRSF1, suggesting that NF20-40 contains a factor capable of functionally interacting with the CTD. We have now purified and characterized the factor responsible for this activity, and found that it consists of a complex containing both U2AF65 and PRP19C. U2AF65 and PRP19C interact directly in vitro, and in an

RNA-independent manner in vivo. Additionally, U2AF65 binds directly to the phosphorylated

4 139 CTD, increasing U2AF association with the pre-mRNA and recruitment of PRP19C. U2AF65

thus bridges the transcriptional machinery and later stages of spliceosomal assembly, through

novel interactions with the RNAPII CTD and the PRP19C.

RESULTS

CTD-specific splicing activity co-purifies with U2AF65 and PRP19C

We previously observed that a splicing substrate called IgMA3 (see Fig. 1A) is not

spliced in S-100 complementation assays in the presence of 100 nM SRSF1 or SRSF1-CTD

(structure of the fusion protein is shown in Figure 1A). However, addition of an ammonium

sulfate (AS) fraction derived from HeLa NE (NF20-40) specifically activated splicing in the

presence of SRSF1-CTD, but not SRSF1 alone (Millhouse and Manley 2005). To identify the

factor(s) involved in the CTD-dependent splicing activity, we chromatographically purified the

activity (purification scheme shown in Supplementary Fig. 1). We found that the NF20-40

activity bound to a butyl-FF column, and eluted during a gradient between 400-0 mM AS (data

not shown). The active fractions were applied to a Mono Q column, with the activity eluting

early in the gradient, between 40-60 mM AS. The active Mono Q fractions were then loaded on a

Mono S column, and the activity bound tightly, with the strongest peak of activity eluting at 800

mM AS (Figure 1B, fractions 17 and 18). The Mono S fractions were separated by SDS-PAGE

and stained with commassie blue (Figure 1C). Protein bands in the most active fraction, 18, were

excised and proteins identified by mass spectrometry (Figure 1C). Unexpectedly, we identified not only all four of the core components of PRP19C, as well as HSP73, but also the early-acting splicing factor U2AF65. The only species identified not associated with PRP19C or U2AF65 were multiple isoforms of the transcription factor TFII-I. The identities of U2AF65, Prp19 and

TFII-I was confirmed by immunoblotting (Figure 1D). Importantly, U2AF65 and Prp19 were present in all active Mono S fractions. TFII-I eluted later in the gradient than U2AF65 and Prp19

(Figure 1D, fractions 15-18), and was barely detectable in the first active fraction (data not

5 140 shown). Because U2AF65 usually exists as a dimer with U2AF35, we tested the Mono S

fractions for the presence of U2AF35 by immunoblotting (Figure 1D). U2AF35 was weakly detectable in fractions 17 and 18, but undetectable in all earlier fractions, indicating that U2AF35 is dispensable for the CTD-dependent splicing activity.

We also examined NF20-40 Mono Q fractions for a potential correlation between the presence of PRP19C and U2AF65 and CTD-dependent splicing activity. Immunoblotting

showed that PRP19C, specifically CDC5L, Prp19 and SPF27, eluted early in the Mono Q

gradient, and coincided with U2AF65 in fractions 2-5 (Figure 1E). U2AF65 eluted in a broader

peak, beginning in the earliest gradient fraction and continuing through gradient fraction 7.

Notably, splicing activity was restricted to fractions that contained both U2AF65 and PRP19C

(Figure 1F, fractions 2-5). Later fractions, which were also enriched in U2AF65, were inactive in the splicing assay.

U2AF65 interacts with the PRP19C in vitro and in vivo

Given that U2AF65 and PRP19C co-purify in active fractions, we next wished to test the possibility that U2AF65 and PRP19C physically interact. To this end, we expressed and purified

GST-tagged U2AF65 in E. coli (Figure 2A). Using GST-U2AF65 in a pulldown assay with

NF20-40, a strong interaction with Prp19 was detected by Western blotting with anti-Prp19 antibodies (Figure 2B, lane 6), while a somewhat weaker interaction was observed with S-100

(Figure 2B, lane 3). The Prp19 antibody detected additional bands (most prominently in S-100;

Figure 2B, lanes 1 and 2) that appear to correspond to previously described ubiquitinated forms of Prp19 (Lu and Legerski 2007). Consistent with that report, which showed that Prp19 ubiquitination disrupts its association with other complex members, these modified forms failed to co-fractionate with other PRP19C subunits on Mono Q (data not shown), suggesting that the modification disrupts interactions between Prp19 and other complex subunits. The modified forms of Prp19 also failed to interact with U2AF65 (Figure 2B), suggesting that only unmodified,

complex-associated Prp19 interacts with GST-U2AF65. To identify the region(s) of U2AF65

6 141 required to interact with PRP19C, we constructed deletion mutants lacking the N-terminal RS domain or the C-terminal U2AF homology (UHM) domain, regions implicated in protein-protein interactions (Kielkopf et al. 2004; Shepard and Hertel 2009). The mutant protein lacking the

UHM domain bound Prp19 with efficiency comparable to full-length U2AF65 (Figure 2B, lanes

5 and 8). In contrast, truncation of the RS domain completely eliminated the interaction with

Prp19 (Figure 2B, lanes 4 and 7).

We next wished to determine if U2AF65 and PRP19C interact in vivo. As a first experiment, we transiently transfected a plasmid expressing flag-tagged SPF27 into 293T cells, prepared NE, and incubated the NE with anti-flag M2 agarose beads. After incubation, aliquots of beads were washed with buffer containing 100 mM or 400 mM KCl. Bound proteins were eluted using flag peptide and analyzed by coomassie staining and immunoblotting. The resulting complexes contained polypeptides that correspond to all four core PRP19C subunits, CDC5L,

PLRG1, Prp19 and SPF27 (see Figure 2C, lanes 1 and 2; data not shown). The intensity of the

Prp19 band was considerably stronger than the SPF27-flag band, consistent with a 4:1

Prp19:SPF27 stoichiometry in PRP19C (Grote et al. 2010), and indicating that SPF27-flag was efficiently incorporated into the endogenous PRP19C. Importantly, immunoblotting with anti

U2AF65 antibodies indicated that washing with low salt buffer resulted in co-purification of

U2AF65, while higher salt, which did not affect the amount of Prp19 that co-purified with

SPF27-flag, eliminated the co-purification of U2AF65 (Figure 2C, lanes 4 and 5). Neither protein was detected in the eluate from beads incubated with NE from non-transfected cells

(Figure 2D, lanes 1-3). These data indicate that U2AF65 and PRP19C form a salt-labile complex in NE.

To examine whether endogenous U2AF65 and PRP19C interact, we performed immunoprecipitations (IPs) with NF20-40. Using anti-Prp19 antibodies for IP, we indeed detected co-IP of U2AF65 (Figure 2D, lane 3). The same interaction was observed using an anti-

CDC5L antibody, as well as antibodies against two additional PRP19C subunits, PLRG1 and

7 142 SPF27 (Figure 2E, lanes 2 and 4; data not shown). Importantly, co-IP of U2AF65 was

maintained after extensive treatment of the extract with RNase A (Figure 2F, compare lanes 2 and 4), indicating that the interaction was not bridged by RNA, as might occur for example in spliceosomes.

U2AF and PRP19C are required for CTD-dependent splicing activity

We next wished to determine whether U2AF65 and PRP19C are in fact required for the

CTD-dependent splicing activity. We first examined the effect of depleting U2AF from NF20-40.

To do this, we passed the NF20-40 through a Poly-U sepharose column at a salt concentration of

1M KCl, while a mock sample was prepared using an equal volume of glutathione sepharose

beads. As expected (Zamore and Green 1991), passage through a Poly-U column resulted in

efficient depletion of U2AF65 from the NF20-40 (Figure 3A, top panel, lane 2). Because under

high salt-conditions U2AF65 and PRP19C do not interact (see above), depletion of U2AF65 in

this manner resulted in minimal codepletion of PRP19C, as judged by CDC5L levels (Figure 3A,

bottom panel). Importantly, depletion of U2AF resulted in an approximately 5-fold reduction in

CTD-dependent splicing activity compared to the mock-depleted NF20-40 (Figure 3B, compare

lanes 2 and 3). We next tested purified U2AF65 for its ability to restore activity to the depleted

NF20-40. While neither baculovirus-expressed and purified U2AF65, nor the U2AF heterodimer,

was capable of activating CTD-dependent splicing alone (Figure 3B, lanes 6 and 7), we found

that both were able to partially restore activity to depleted NF20-40 (Figure 3B, lanes 4 and 5).

To confirm these results, we also used an anti-U2AF65 antibody to immunodeplete U2AF from

NF20-40, at 500 mM NaCl, which resulted in an approximately 85% depletion of U2AF65, but

no change in PRP19C levels, as judged by CDC5L (Supplementary Figure 2A). This treatment

resulted in over a 60% decrease in activity, compared to mock-depleted NF20-40

(Supplementary Figure 2B). On the basis of these experiments, we conclude that U2AF65 is

necessary but not sufficient for the CTD-dependent splicing activity.

8 143 We next wished to determine whether the PRP19C is necessary for CTD-dependent splicing activity. To do this, we used anti-CDC5L antibodies to immunodeplete PRP19C, in this case from an active Mono Q fraction. This resulted in an approximately 60% depletion of

PRP19C, as judged by CDC5L levels (Figure 3C, top panel, compare lanes 1 and 2). Importantly, this caused a greater than 50% reduction in activity when compared to a mock-depleted control

(Figure 3D, compare lanes 1 and 2). As PRP19C depletion was performed at 500 mM NaCl, levels of U2AF65 were not affected (Figure 3C, bottom panel, lanes 1 and 2). Although additional rounds of incubation with antibodies led to near complete depletion of PRP19C, this treatment resulted in loss of activity in both the mock and depleted fractions (data not shown).

Nonetheless, the reduction of activity we observed upon partial depletion of PRP19C indicates that PRP19C contributes to the CTD-dependent splicing activity of the NF20-40.

U2AF65 and PRP19C association is required for CTD-dependent splicing

U2AF65 and PRP19C are both present in S-100, which nonetheless is unable to substitute for NF20-40 to activate CTD-dependent splicing. To investigate the basis for this, we next examined the behavior of S-100-derived U2AF65 and PRP19C on a Mono Q column. The elution profile of each factor was strikingly different than observed with NF20-40, with PRP19C

(represented by CDC5L) eluting in a narrow peak early in the Mono Q gradient (Figure 4A, fraction 6) and U2AF65 eluting much later (Figure 4A, fractions 10-14). Consistent with the inability of S-100 to activate CTD-dependent splicing, none of the S-100 derived Mono Q fractions were active, even when concentrated PRP19C and U2AF65-containing fractions were mixed together (data not shown).

The failure of U2AF65 and PRP19C to co-fractionate when derived from S-100 suggests that the two complexes are not physically associated when present in the cytoplasmic extract. To test this, we performed parallel IPs in NE and S-100 using the anti-CDC5L antibody. In NE, as with the NF20-40, CDC5L co-IPed U2AF65, while in S-100 co-IP did not occur (Figure 4B, compare lanes 2 and 4). On the basis of co-IP and chromatographic data, we conclude that

9 144 endogenous U2AF65 and PRP19C are physically associated in NE but not in S-100. Possible

explanations for this behavior are discussed below. In any event, this data supports the idea that a physical interaction between U2AF65 and PRP19C in the NF20-40 is required for the CTD- dependent splicing of IgMA3.

The above data suggest that CTD-dependent splicing activity should reside in a complex containing U2AF65 and PRP19C. To estimate the molecular weight of the CTD-dependent splicing activity, we separated the NF20-40 on a Superdex 200 gel filtration column, then performed splicing assays using the resulting fractions. Importantly, the activity eluted at an apparent mass of over 670 kDa (Figure 4C, fractions 3-9), consistent with the expected molecular mass of a putative U2AF65-PRP19C complex. The elution of the activity in fractions

3-9 coincided perfectly with fractions in which PRP19C and U2AF65 were both present, as judged by immunoblotting (Figure 4D), while a number of lower molecular weight fractions containing U2AF65 alone (Figure 4D, fractions 10-14) exhibited no splicing activity (Figure 4C, fractions 10-14). Taken together, these results strongly suggest that the CTD-dependent splicing activity resides in a complex containing U2AF65 and PRP19C.

CTD-dependent splicing activity can be reconstituted by U2AF65/PRP19C purified from mammalian cells

We next tested the ability of purified U2AF/PRP19C to reconstitute CTD-dependent splicing activity. To isolate a PRP19C-U2AF complex, we transfected a plasmid encoding flag- tagged SPF27 into 293T cells, then used anti-flag beads to purify PRP19C from NE made from these cells by washing the beads with buffer containing either 100 mM or 400 mM KCl. As shown above (Figure 2C), washing the beads with 100 mM KCl resulted in co-purification of

U2AF65, while washing with 400 mM KCl removed U2AF65. Importantly, when used in CTD- dependent splicing assays, the low-salt PRP19C activated splicing in a concentration-dependent manner, while the stringently purified PRP19C exhibited no more activity than the mock eluate

10 145 (Figure 5A, compare lanes 3-5 and 6-8). As a complementary approach, we prepared the

U2AF65/PRP19C complex from 293T cells transfected with a plasmid encoding U2AF65-flag, using 100 mM KCl to wash the beads (Figure 5B). U2AF65 purified under these conditions was associated with PRP19C, as Western blotting revealed that both CDC5L and Prp19 were present in the U2AF65 preparation (Figure 5B, bottom panel). In contrast to the more stringently purified baculovirus preparations of U2AF/U2AF65 analyzed above (see Figure 3B), the PRP19C- containing U2AF65 preparation was able to activate splicing when used in the CTD-dependent splicing assay (Figure 5C, lanes 3). Together, these results indicate that a U2AF65-PRP19C complex can activate CTD-dependent splicing.

The RNAPII CTD binds U2AF65 directly

We next wished to elucidate the role of the CTD in the U2AF65-PRP19C splicing stimulatory activity. Based on its function in other processes, a likely possibility is that one or more U2AF65-PRP19C subunits interact directly with the CTD, and this interaction helps recruit the complex to the RNA substrate. Given that U2AF65 has been shown to associate tightly with elongating RNAPII in vitro (Ujvari and Luse 2004), it was a logical candidate to participate in such an interaction. We therefore tested whether U2AF65 can interact with the CTD, first by performing pulldown assays with GST-CTD that was either unphosphorylated or phosphorylated in and repurified from NE (p-CTD; see Hirose and Manley, 1998; Figure 6A). When we used

NF20-40 as input for the pulldown, we found that U2AF65 interacted with both CTD and p-CTD

(Figure 6A, top panel). However, after partial purification by Mono Q chromatography, U2AF65 only bound to p-CTD, suggesting that the interaction with the unphosphorylated CTD observed with NF20-40 was mediated by other factors present in the fraction (Figure 6A, middle panel).

Interestingly, we also detected pulldown of PRP19C (Prp19) from NF20-40 by p-CTD

(Supplementary Figure 3A). This interaction may be bridged by U2AF65, as it was not observed

11 146 with S-100, where the two complexes fail to interact, but in which U2AF65 bound strongly to p-

CTD (Supplementary Figure 3B).

We next wished to provide evidence that the CTD-U2AF65 interaction was direct. To

this end, we used purified his-tagged U2AF65 produced in baculovirus-infected cells in the GST- pulldown assay. Importantly, a strong, p-CTD-specific interaction was detected (Figure 6A,

lower panel), indicating that U2AF65 directly interacts with the phosphorylated CTD. In addition,

the U2AF65-CTD interaction was robust enough to persist after washing with up to 500 mM

NaCl (Supplementary Figure 4A), and was resistant to RNase A (Supplementary Figure 4B). In

order to identify the region(s) of U2AF65 that contribute to p-CTD binding, we used GST-

U2AF65 and the RS and UHM domain truncations described above in pulldown assays with

purified p-CTD that had been cleaved from GST using thrombin. We found that only full-length

U2AF65 bound strongly to p-CTD, while each of the truncation mutants displayed significantly

reduced p-CTD binding (Figure 6C, lanes 2-4). This suggests that U2AF65 interacts with the p-

CTD in a manner that requires both ends of the protein for optimal binding.

The RNAPII CTD-U2AF65 recruits U2AF and PRP19C to pre-mRNA

The interaction of U2AF65 with the phosphorylated CTD has the potential to underlie the

CTD-dependence of the U2AF-PRP19C stimulatory activity. The IgMA3 splicing substrate contains a weak polypyrimidine tract, to which U2AF65 is recruited inefficiently in the absence of an exonic splicing enhancer in the second exon (Graveley et al. 2001). A model for how

SRSF1-CTD stimulates IgMA3 splicing is that substrate-bound SRSF1-CTD recruits U2AF65 to the pre-mRNA more efficiently than SRSF1 alone, as a result of the p-CTD-U2AF65 interaction.

To test this possibility, we performed UV crosslinking with 32P-IgMA3 RNA and S-100 plus

NF20-40 in the presence of SRSF1 or SRSF1-CTD, followed by IP with anti-U2AF65 antibodies.

Significantly, we detected an increase in U2AF65 crosslinking in the presence of SRSF1-CTD

compared to SRSF1 (Figure 6D, compare lanes 2 and 3). To extend this result, we performed the

12 147 crosslinking assay with purified baculovirus U2AF65 together with SRSF1 derivatives lacking the SRSF1 RS domain (SRSF1ΔRS and SRSF1ΔRS-CTD; Millhouse and Manley, 2005). As above, enhanced crosslinking was observed with SRSF1ΔRS-CTD relative to SRSF1ΔRS

(Figure 6E, lanes 2 and 3). Together, these results provide strong evidence that the CTD-

U2AF65 interaction is sufficient to facilitate U2AF recruitment to the pre-mRNA.

We next asked whether the increased recruitment of U2AF65 to the IgMA3 pre-mRNA by SRSF1-CTD leads to increased recruitment of PRP19C. To test this, we again set up splicing reactions with 32P-IgMA3 RNA in the presence of NF20-40 and S-100, as well as SRSF1 or

SRSF1-CTD. After a 30 minute incubation, reaction mixtures were IPed using a mock antibody

(anti-GST) or anti-PLRG1 antibodies, and IPed RNA purified and resolved by denaturing PAGE.

Strikingly, we found that PLRG1 strongly associated with pre-mRNA in the presence of SRSF1-

CTD but not SRSF1, indicating that Prp19C recruitment to the pre-mRNA was significantly enhanced by the CTD (Figure 6F, compare lanes 3 and 4). We conclude that an interaction between U2AF65 and p-CTD promotes U2AF65 binding to the IgMA3 substrate, and this leads to enhanced Prp19C recruitment, reflecting the interaction between U2AF65 and Prp19C.

DISCUSSION

Here we have used a biochemical complementation assay described previously to purify and characterize an activity capable of activating CTD-dependent splicing in vitro. This resulted in the unexpected discovery that two well-studied splicing factors not previously known to associate, U2AF65 and PRP19C, functionally and physically interact to activate splicing. This was first indicated by our observation that both factors co-purify in active fractions after extensive chromatography. We also showed that U2AF65 and PRP19C interact in an RNA-

13 148 independent manner, and that the two indeed exist in a salt-sensitive complex in vivo.

Importantly, several experimental approaches confirmed that the U2AF65/PRP19C complex is

indeed responsible for the CTD-dependent splicing activity. The role played by the CTD was

suggested by our discovery that U2AF65 binds directly to the phosphorylated CTD, and we

showed that this enhanced both U2AF65 binding and PRP19C recruitment to the pre-mRNA.

Below we discuss the implications of these findings, with respect both to the link between splicing and transcription and to splicing per se.

How might U2AF65 and PRP19C cooperate to activate CTD-dependent splicing? Our data suggest that the association of the two factors in a complex, and not simply their concentration, is important. For example, the concentration of each is considerably higher in S-

100 than in some active fractions (unpublished data), but S-100 is incapable of activating CTD- dependent splicing. Also, addition of purified U2AF65 and PRP19C in combination did not result in a significant increase in activity. U2AF65/PRP19C-contining fractions activated CTD- dependent splicing only when the two factors were associated, and gel filtration indicated that

CTD-dependent splicing activity resides in an HMW complex of a size consistent with a

PRP19C-U2AF65 complex. What determines whether the two splicing factors interact is not clear, but may reflect the presence or absence of a bridging factor and/or differential protein modifications.

Earlier work on the IgM substrate may provide some clues as to why a pre-formed complex containing U2AF65 and PRP19C is necessary to activate IgMA3 splicing. Green and colleagues showed that the second exon of the IgM substrate contains an inhibitory element that is bound by the splicing inhibitory protein polypyrimidine tract binding protein (PTB) (Kan and

Green 1999; Shen et al. 2004). These investigators showed that splicing inhibition by PTB does

14 149 not prevent early steps in spliceosome assembly, but rather results in an ATP-dependent complex that is similar in electrophoretic mobility and composition to spliceosomal A complex, but which is unable to progress to later stages in spliceosome assembly (Kan and Green 1999). A similar stalled A-like complex also forms on an in vitro splicing substrate containing the PTB-inhibited c-src N1 exon (Sharma et al. 2008). Proteomic analysis comparing complexes formed in the presence or absence of PTB inhibition showed that PRP19C recruitment is impaired when PTB functions to inhibit splicing (Sharma et al. 2008). PTB inhibition is likely relevant to our S-100 complementation assay, as the PTB binding sites described by Green and colleagues are intact in the IgMA3 substrate, and UV crosslinking reveals PTB as the major crosslink in splicing reactions using S-100 (unpublished data).

Based on the above, we propose the following model for CTD-dependent activation of

IgMA3 splicing. This substrate is unable to recruit U2AF efficiently, due to its weak polypyrimidine (py) tract (Graveley et al. 2001). Instead, a U2AF65-PRP19C complex is recruited to the pre-mRNA by RNA-bound SRSF1-CTD, through the direct interaction between the CTD and U2AF65. U2AF35, which is normally required to mediate interactions between enhancer-bound SR proteins and U2AF65 (Graveley et al. 2001), appears to be dispensable, as

U2AF35 was not detectable in some active fractions, and was not required to reconstitute activity.

It is likely that the CTD-U2AF65 interaction makes U2AF35 unnecessary for recruitment of

U2AF65 to the IgMA3 py tract. The p-CTD-U2AF65 interaction also results in recruitment of

PRP19C, most likely through its interaction with U2AF65. The combination of SRSF1-CTD and the U2AF65-PRP19C complex thus overcomes two important barriers to the assembly of active

IgMA3 spliceosomes: First, the recruitment of U2AF65 to the suboptimal IgM py-tract, and second, recruitment of PRP19C to PTB-inhibited early spliceosomes.

15 150 Aside from a possible role in IgMA3 splicing, the U2AF65-PRP19C interaction likely

plays a broader role in spliceosome assembly. As mentioned in the Introduction, PRP19C is a

stable component of catalytically activated spliceosomes (Bessonov et al. 2008), but the

interactions that underlie its recruitment to the spliceosome are currently poorly understood.

Studies in yeast originally suggested that PRP19C associates with the spliceosome at a late stage

in its assembly, after the recruitment of the U4/U6.U5 tri-snRNP (Tarn et al. 1993). However, more recent data have shown that human PRP19C is a part of the first ATP-dependent spliceosomal complex, A complex, and its recruitment is independent of the tri-snRNP

(Behzadnia et al. 2007). PRP19C association may occur even earlier than A complex, as it can associate with pre-mRNA in the absence of ATP (Jurica et al. 2002). These data indicate that

PRP19C interacts with a factor that associates with early spliceosomes, resulting in its initial

recruitment to the pre-mRNA. The interaction with U2AF65 that we documented here is a strong

candidate for such an interaction. Additionally, the fact that the U2AF65 RS domain is required

to interact with PRP19C raises the possibility that RS domains on other spliceosomal proteins

may be able to perform a similar function. Interestingly, a U2AF-PRP19C connection appears to

be conserved in yeast, as the yeast U2AF65 homolog Mud2 was shown to interact physically

with a subunit of yeast PRP19C, Clf1 (Chung et al. 1999). Clf1 has a human homolog, Crn,

which has been shown to associate with spliceosomes (Chung et al. 2002), but has not been found to associate with human PRP19C.

What bridges U2AF and PRP19C? Despite some effort, we have been unable to identify a core PRP19C subunit that interacts with U2AF65 (unpublished data). One possibility is that a more loosely associated factor bridges the two complexes. Far western blotting using highly

purified active fractions revealed a protein of approximately 75 kDa that specifically interacts

16 151 with U2AF65 (unpublished data). The only protein we identified by mass spectrometry in that

region was HSP73, making HSP73 a candidate to bridge U2AF65 and PRP19C. Interaction

studies have shown that HSP73 is the most loosely associated component of the PRP19C (Grote

et al. 2010), which is consistent with the salt-lability of the of the U2AF65-PRP19C complex.

However, using purified E. coli-expressed proteins, we have been unable to detect an interaction between U2AF65 and HSP73 (unpublished data). Additional studies will be necessary to identify conclusively the factor that bridges U2AF65 and PRP19C.

Our results may also have implications for regulation of alternative splicing (AS). Since it

appears likely that increased PRP19C recruitment can counteract PTB inhibition, Prp19C

recruitment may serve as a regulatory point in the control of AS. Consistent with this idea, it was

recently shown that the AS regulatory protein hnRNP M can directly interact with PRP19C, and

this interaction appears to be important for regulation of AS by this protein (Lleres et al. 2010).

Additionally, our results point to the possibility that splicing of different introns might be differently affected by modulation of PRP19C levels, which occurs during processes such as neuronal differentiation (Urano et al. 2006).

In addition to the association between U2AF and PRP19C, we have also identified a

novel interaction between U2AF65 and the CTD. This raises the possibility that U2AF65 plays

an important role in the physical coupling of transcription and splicing. The fact that U2AF65

binds the CTD is consistent with previous studies. For example, U2AF65 has been reported to

associate very tightly with RNAPII transcription elongation complexes (TECs) in vitro (Ujvari

and Luse 2004). Importantly, these experiments showed that U2AF65 in the TEC was positioned

to interact with RNA immediately upon its extrusion from the polymerase. Such positioning

appears consistent with CTD binding, as the base of the CTD is located near the RNA channel

17 152 (Cramer et al. 2001). Additionally, U2AF65 has been shown by chromatin immunoprecipitation to associate with transcriptionally active genes, including transcribed regions well upstream of the first 3’ splice site, consistent with RNAPII-mediated recruitment of U2AF65 to intron- containing genes (Listerman et al. 2006). Similar experiments with yeast have shown that Mud2 is co-transcriptionally recruited to active genes, raising the possibility that the interaction we identified here is evolutionarily conserved (Gornemann et al. 2005; Lacadie et al. 2006).

Our results, coupled with previous studies, provide an attractive model to explain how the efficiency of splicing in vivo can be enhanced by coupling it to transcription. Specifically, since

U1 snRNP has been previously shown to associate with RNAPII (Das et al. 2007), the interaction between U2AF65 and elongating RNAPII, via direct CTD binding, means that both major factors that recognize the 5’ and 3’ splice sites are likely recruited to nascent transcripts.

We envision a scenario (Figure 7) in which RNAPII-associated U1 snRNP, and SR proteins, first recognize a 5’ splice site in the nascent RNA, resulting in tethering of the exon to RNAPII (Dye et al. 2006; Das et al. 2007). In the meantime, CTD-bound U2AF65 is positioned to interact with the 3’ splice site as soon as it is synthesized. Such an arrangement would result in rapid splice site recognition, the immediate juxtaposition of the two ends of the intron, as well as PRP19C recruitment, through interaction with U2AF65, thereby facilitating efficient formation of mature spliceosomal complexes on nascent transcripts. In any event, our results have defined key molecular interactions that underlie the coupling of RNAPII transcription and splicing, and the role of the CTD in this process.

MATERIALS AND METHODS

In vitro splicing

18 153 In vitro splicing assays were performed using S-100, appropriate nuclear fractions, and

SRSF1 or SRSF1-CTD as described (Millhouse and Manley 2005). Imagequant software was

used to quantify the amount of spliced product (mRNA + intron) in each lane, with the

equivalent regions in buffer D-containing reactions set as background.

Chromatography

NE for chromatography was prepared as described (Dignam et al. 1983). NF20-40 was

prepared as described (Millhouse and Manley 2005). Chromatography was performed in buffer

D (20 mM HEPES, pH 7.9, 5% glycerol, 0.2 mM EDTA, 100 mM KCl, 0.5 mM DTT, 1 mM

PMSF), except for the Mono Q, which was run with buffer containing 20 mM tris-HCl, pH 7.9 instead of Hepes. For chromatography, 120mg NF20-40 was loaded on a 40 ml Butyl-FF Hitrap column (GE healthcare), and eluted with an 800-0 mM AS gradient in buffer D. Active fractions were pooled, concentrated by AS precipitation, desalted, then loaded on an 8 ml Mono Q column

(GE healthcare). The Mono Q column was developed with a 0-250 mM AS gradient. Mono Q fractions were pooled, concentrated by AS precipitation, dialyzed in a microdialyzer (Gibco), then assayed. Active fractions were loaded on a 1 ml Mono S column, which was developed with a 0-250 mM AS gradient, followed by a step to 800 mM AS. Mono S fractions were dialyzed as above, prior to CTD-dependent splicing assay. Excised proteins were identified using MALDI-

TOF mass spectrometry essentially as described (Shevchenko et al. 2006).

Plasmids and recombinant proteins

SRSF1-CTD, and SRSF1, SRSF1ΔRS-CTD, SRSF1ΔRS, and IgMA3 splicing substrate were prepared as described (Millhouse and Manley 2005). Baculovirus producing his-U2AF65 and his-U2AF65/35, a gift of Brenton Graveley, was used to infect Hi5 cells and purified as described (Graveley et al. 2001). To produce U2AF65 and Prp19C in mammalian cells, U2AF65 and SPF27 were cloned into p3xFlag-CMV-14, and transfected into 293T cells. The following day, cells were split 1:5, then 48h after transfection, cells were collected and used to make NE.

Purification of complexes was performed overnight using M2-agarose beads (Sigma), washed

19 154 with Buffer D containing the specified concentration of KCl three times, then eluted with

0.2mg/ml 3xFlag peptide (Sigma). GST-U2AF65 and deletion mutants were cloned into pGEX-

6P1, and induced in Rosetta cells at 16˚C overnight and purified using glutathione sepharose (GE

healthcare)

Antibodies and immunoblotting

The following antibodies were used: U2AF65 (MC3; Sigma), PLRG1 (Novus- NBP1-

06556), SPF27 (Novus- NB110-40681), anti-CTD 8WG16 (Lab stock), TFII-I (a gift of Ananda

Roy, Tufts Univ.), U2AF35 (a gift of Tom Maniatis, Columbia University Medical Center), GST

(Molecular probes A5800), anti-Flag M2 (Sigma F1084). A Prp19 antibody from Bethyl (A300-

102A) was used in Figures 1E, 2B and 5B, and a Prp19 antibody provided by R. Luhrmann was

used in Figures 1D and 2D. A rabbit anti-CDC5L antibody from Abcam (ab31779-100) was used

for the co-IP experiments in Figure 2A, while a mouse anti-CDC5L antibody from Santa Cruz

(sc-81220) was used for immunodepletion.

Immunoblotting was performed Li-cor secondary antibodies, and quantified using Li-cor software.

GST pulldown

For GST-pulldowns using GST-tagged U2AF65 or truncated derivatives, 5 µg immobilized protein was used for pulldown in 250µg S-100/NF20-40. Proteins were incubated

for 3h at 22˚C, then washed with buffer D. Proteins were eluted with 15 mM glutathione prior to

SDS-PAGE and immunoblot analysis. GST-CTD was prepared and phosphorylated as described

(Hirose and Manley 1998). For pulldowns, 5µg of each CTD fusion protein was used for each

250µg of extract or column fraction, and incubated at 22˚C for 3 hours, then washed three times

with buffer D and eluted with 15 mM glutathione. For GST-CTD pulldown with recombinant

U2AF65, 10µg his-U2AF65 was incubated with 5 µg GST-CTD and incubated, washed and

eluted as above. For pulldowns performed with RNase pre-treatment, extracts were pre-

20 155 incubated in the presence of 10µg/ml RNase A at 37˚C for 30 minutes prior to incubation with

GST-p-CTD.

Immunoprecipitations and depletions

For co-IPs in Figure 2, 2µg of each antibody (anti-Prp19 or CDC5L) was used to IP

100µg of NF20-40. Beads were washed with buffer D before boiling and SDS-PAGE. For immunodepletions, 12µg of each antibody (CDC5L or U2AF65) was used for each 50µg of

NF20-40 or Mono Q fraction. For each, 12µg of anti-flag M2 antibody served as a mock.

Depletions were carried out for 2h at 4˚C, in buffer D containing 500 mM NaCl. Following depletion, proteins were precipitated by addition of AS to 85%, redissolved in buffer D and dialyzed against buffer D. Poly-U depletion of U2AF was performed as described (Page-McCaw et al. 1999). Immunoprecipitation of spliceosomes was performed as described (Blencowe et al.

1994).

UV crosslinking

UV crosslinking- U2AF65 immunoprecipitation was performed by setting up splicing reactions using 32P-UTP labeled IgMA3 substrate as described (Millhouse and Manley 2005), but omitting PVA, and incubating for 10 minutes. The splicing reactions were placed on ice then irradiated with UV light in a Stratalinker (Stratagene), followed by 30 minutes of RNase A treatment (10µg/ ml). Immunoprecipitation of crosslinked proteins was performed as described

(Kashima and Manley 2003). For UV crosslinking with purified proteins, 10 µl reactions contining 32P-UTP labeled IgMA3 substrate, and 50 nM his-U2AF65 were incubated on ice for

10 minutes in the presence of buffer D, or 200 nM SRSF1ΔRS/SRSF1ΔRS-CTD before exposure to UV light and RNase A treatment.

ACKNOWLEDGEMENTS

We thank B. Graveley, T. Maniatis, A. Roy, and R. Luhrmann for reagents. We thank N.

Rao and M. Chen for nuclear extracts. J.-P. Hsin, P. Richard, M. Chen, Y. Shi and other Manley

21 156 lab members are acknowledged for helpful discussions. This work was supported by NIH grant

R01-GM048259.

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FIGURE LEGENDS

Figure 1. U2AF65 and PRP19C co-fractionate with the CTD-dependent splicing activity.

(A) Schematic diagrams of the SRSF1-CTD (top) and IgMA3 constructs used in this study.

IgMA3 contains 3 copies of a high affinity binding site for SRSF1 (A3). (B) NF20-40 was

fractionated by Butyl-FF, Mono Q, followed by Mono S chromatography, then Mono S fractions

were assayed for CTD-dependent splicing. Splicing reactions were incubated in the presence of

SRSF1-CTD, S-100 and the fraction indicated for 2h, then RNA was isolated, separated by 6%

denaturing PAGE, then visualized by autoradiography. (C) Mono S fractions from (B) were separated by 10% SDS-PAGE gel and stained by colloidal commassie. Protein bands in fraction

18 were excised and identified by MALDI-TOF mass spectrometry. Protein identities are indicated at right. (D) NF20-40, S-100, and the indicated Mono S fractions from (B) were immunoblotted with the indicated antibodies. Comparisons of protein levels in NF20-40 and S-

100 are provided at left. (E) NF20-40 was fractionated on a Mono Q column, then gradient fractions were immunoblotted with the indicated antibodies. (F) Mono Q fractions from (E) were used in IgMA3 splicing assays in the presence of SRSF1-CTD and S-100, then processed as in

(B).

Figure 2. U2AF65 interacts with PRP19C in an RNA-independent manner.

(A) Schematic of U2AF65 and deletion constructs used for GST-pulldowns (left). These constructs were expressed in E. coli, then purified by GSH-sepharose beads. The beads were run on SDS-PAGE and coomassie stained (right). (B) GST pulldown was performed using GST-

29 164 tagged U2AF65, U2AF65ΔRS, or U2AF65ΔUHM. After a 3 hour incubation at 22˚ C with the

indicated extract, beads were washed, then eluted with 15 mM glutathione. Eluted proteins were

immunoblotted with an anti-Prp19 antibody. (C) A vector expressing flag-tagged SPF27 was

transfected into 293T cells, cells were harvested, used to make NE, and PRP19C was purified

using anti-flag agarose, and washed with buffer D containing the indicated salt concentration. In parallel, beads were incubated with NE from non-transfected cells and washed with buffer D

containing 100 mM KCl. Beads were eluted with flag peptide, then separated by SDS-PAGE and

coomassie stained (left), or immunoblotted with the indicated antibody (right). (D) Co-IPs were

carried out in NF20-40, using anti-Prp19 antibodies, or anti-GST antibodies (mock). Beads were

washed with buffer D, then boiled, and immunoblotted for U2AF65. (E) Co-IP was performed

using anti-CDC5L antibodies or anti-GST antibodies as above, except NF20-40 was mock- or

RNase A- treated beforehand for 30 minutes at 37˚ C.

Figure 3. U2AF and PRP19C are required for the CTD-dependent splicing activity.

(A) NF20-40 was depleted of U2AF65 at 1M KCl using Poly-U sepharose beads, followed by

immunoblotting against U2AF65 and CDC5L. Band intensities quantified using Li-cor software

are indicated below. (B) Mock and depleted extracts from (A) were used in the CTD-dependent

splicing assay (lanes 2 and 3). 100 nM U2AF65 and U2AF produced in baculovirus-infected

insect cells were tested for their ability to complement the depleted NF20-40 (lanes 4 and 5), or

to activate splicing when added alone (lanes 6 and 7). Relative intensities of spliced product were

quantified using Imagequant software and indicated below. (C) PRP19C was depleted from an

active Mono Q fraction at 500 mM NaCl using an anti-CDC5L antibody, or mock depleted using

an anti-flag antibody. Mock and depleted samples were immunoblotted for U2AF65 and CDC5L.

Relative CDC5L levels were normalized to U2AF65, and indicated below. (D) CTD-dependent

splicing assays were performed using the mock- and PRP19C-depleted samples from (C).

Relative amounts of spliced product are indicated below.

30 165

Figure 4. Association of U2AF65 and PRP19C is necessary for CTD-dependent IgMA3

splicing.

(A) S-100 was separated on a Mono Q column under conditions identical to those used in Figure

1E. Immunoblotting for U2AF65 and CDC5L was performed using S-100 (load) and Mono Q fractions. (B) Co-IP was performed using an anti-CDC5L antibody in NE and S-100, followed by anti-U2AF65 immunoblotting. (C) NF20-40 was fractionated on a Superdex 200 column.

Fractions were directly assayed for CTD-dependent splicing. Elution locations of gel filtration standards are indicated at top. (D) Immunoblot of Superdex 200 fractions for U2AF65, CDC5L, and Prp19.

Figure 5. U2AF65 affinity purified from mammalian cells can reconstitute CTD-dependent splicing.

(A) NF20-40 (lane 1), anti-flag eluate prepared from non-transfected cells (lane 2), and PRP19C preparations from (2C) were used in the CTD-dependent splicing assay. For each, 40, 80 or 120 nM was used in splicing reactions. (B) Anti-flag eluate prepared from 293T cells transfected

with U2AF65-flag was separated by SDS-PAGE and coomassie stained (top panel) or

immunoblotted for the indicated proteins (bottom panel). The band marked with * in the

U2AF65 preparation is an N-terminally proteolyzed U2AF65 species. (C) NF20-40 (lane 1), anti-flag eluate prepared from non-transfected cells (lane 2), and 50 nM of the U2AF65-flag

from (B) was used in the CTD-dependent splicing assay.

Figure 6. The CTD binds directly to U2AF65 to recruit U2AF65-PRP19C to pre-mRNA.

(A) GST-CTD was expressed in E. coli, purified and in vitro. 5 µl beads were separated by SDS-

PAGE to ensure equal loading. (B) GST pulldowns were performed using unphosphorylated or

NE-phosphorylated GST-CTD. Pulldowns were performed using crude NF20-40 (top panel),

31 166 U2AF65 partially purified by Mono Q chromatography (middle panel), and recombinant

U2AF65 purified from baculovirus infected insect cells (lower panel). Input and bound samples

were immunoblotted for U2AF65. (C) GST pulldown was performed using GST-U2AF65

constructs used in Figure 2A, and phosphorylated CTD cleaved from GST by thrombin. After

elution, the bound p-CTD was detected by immunoblotting using the anti-CTD antibody 8WG16.

(D) Splicing reactions were set up with 32P-UTP labeled IgMA3 in the presence of NE, or S-100

and SRSF1, SRSF1-CTD or buffer D. After 10 minutes, reactions were exposed to UV light,

followed by RNase treatment. Reactions were then IPed using an anti-U2AF65 or anti-HA

antibody (mock), crosslinked proteins were then separated by 10% SDS-PAGE and visualized by

autoradiography. (E) UV crosslinking reactions using IgMA3 were set up in the presence of

purified U2AF65, SRSF1ΔRS-CTD, or both. After crosslinking and RNase treatment, proteins

were separated by 8% SDS-PAGE, and visualized by autoradiography. (F) Splicing reactions were performed in the presence of S-100, NF20-40, and SRSF1 or SRSF1-CTD. After a 30 minute incubation at 30˚ C, reactions were IPed with anti-GST antibodies (mock) or anti-PLRG1 antibodies. After IP, RNA was extracted and separated by denaturing 6% PAGE and visualized by autoradiography.

Figure 7. Model for activation of CTD-dependent splicing by a U2AF-PRP19C complex.

(A) At promoters, RNAPII is present in preinitiation complexes, but the CTD is unphosphorylated and unable to recruit splicing factors. (B) Transcription initiation results in

CTD phosphorylation by multiple kinases, resulting in the association of splicing factors, including SR proteins, U1 snRNP, through unknown interactions, as well as the U2AF-PRP19C complex via a direct interaction with U2AF65. RNAPII-associated U1 and SR proteins recognize a transcribed exon, resulting in its tethering to the RNAPII elongation complex through multiple interactions. (C) Transcription of the 3’ splice site results in a transition from protein-protein interactions between U2AF65 and the p-CTD to protein-RNA interactions, resulting in efficient

32 167 recognition of the 3’ splice site. This facilitates rapid transition to a mature spliceosomal complex, promoted by U2AF65-associated PRP19C.

SUPPLEMENTAL FIGURES

Supplementary Figure 1. Scheme used to purify the NF20-40 CTD-dependent splicing activity.

12 ml of NF20-40 was first loaded on a butyl-FF column and eluted with a decreasing AS gradient. Active fractions from 400-0 mM AS were collected, and loaded on a Mono Q column.

The Mono Q was developed with a 0-250 mM AS gradient, and active fractions from 40-60 mM

AS were loaded on a Mono S column. The Mono S column was developed with a 0-250 mM AS gradient, followed by a step to 800 mM AS.

Supplementary Figure 2. U2AF65 immunodepletion results in decreased CTD-dependent splicing activity.

(A) U2AF65 in NF20-40 was immunodepleted at 500 mM NaCl using the MC3 monoclonal antibody. Extracts were then immunoblotted for CDC5L and U2AF65. After quantification,

U2AF65 levels were normalized to CDC5L. (B) CTD-dependent splicing assay using mock- and

U2AF65-depleted NF20-40. Relative levels of spliced product were quantified using Imagequant software, indicated below.

Supplementary Figure 3. Prp19 interacts with the p-CTD in NF20-40 but not S-100.

(A) GST-CTD pulldowns were performed in NF20-40 and S-100, and immunoblotted for the presence of Prp19. (B) GST-CTD pulldowns were performed as above, and immunoblotting was performed for U2AF65. The left panel is reproduced from Figure 6A.

Supplementary Figure 4. p-CTD-U2AF65 interaction is salt and RNase resistant.

33 168 (A) Salt resistance of U2AF65-GST-p-CTD interaction. GST-pCTD pulldown was performed in a Mono Q fraction (as in 6B, middle panel), then washed with buffer D containing increasing concentrations of salt. (B) GST-p-CTD was used to pull down U2AF65 present in NF20-40, after treatment of the extract with RNase A for 30 minutes at 37˚C.

34 169 170 171 172 173 174 175 176 177 178