Poly(A) Tail Length Is Controlled by the Nuclear Poly(A)-binding Protein Regulating the Interaction between Poly(A) Polymerase and the Cleavage and Polyadenylation Specificity Factor

Poly(A) tails of mRNAs are synthesized in the cell nucleus with a defined length, ∼250 nucleotides in mammalian cells. The same type of length control is seen in an in vitro polyadenylation system reconstituted from three proteins: poly(A) polymerase, cleavage and polyadenylation specificity factor (CPSF), and the nuclear poly(A)-binding protein (PABPN1). CPSF, binding the polyadenylation signal AAUAAA, and PABPN1, binding the growing poly(A) tail, cooperatively stimulate poly(A) polymerase such that a complete poly(A) tail is synthesized in one processive event, which terminates at a length of ∼250 nucleotides. We report that PABPN1 is required to restrict CPSF binding to the AAUAAA sequence and to permit the stimulation of poly(A) polymerase by AAUAAA-bound CPSF to be maintained throughout the elongation reaction. The stimulation by CPSF is disrupted when the poly(A) tail has reached a length of ∼250 nucleotides, and this terminates processive elongation. PABPN1 measures the length of the tail and is responsible for disrupting the CPSF-poly(A) polymerase interaction.The poly(A) tails present at the 3′ end of almost all eukaryotic mRNAs have two major functions. The first function is in the control of mRNA decay; degradation of the poly(A) tail by a 3′ exonuclease (deadenylation) is the first step in both of the two main pathways of mRNA decay, and the completion of deadenylation triggers the second step, either cap hydrolysis or further 3′–5′ degradation. Because the rate of deadenylation is governed by sequence elements in the mRNA, it is specific for each mRNA species and serves as a major determinant of mRNA half-life (1–3). Obviously, a control of mRNA stability by the rate of deadenylation requires a defined poly(A) length as a starting point. The second function of the poly(A) tail is in the initiation of translation; the cytoplasmic poly(A)-binding protein associated with the poly(A) tail promotes the initiation of translation by an interaction with the initiation factor eIF4G and probably through additional mechanisms (4–7). In this process, poly(A) tail length can also be important. For example, gene regulation during oocyte maturation and early embryonic development of animals depends on translational regulation of maternal mRNAs, and changes in poly(A) tail lengths of specific mRNAs, determined both by deadenylation and by regulated cytoplasmic poly(A) extension, play a major role in this translational regulation. Long poly(A) tails favor translation, whereas a shortening of the tail promotes translational inactivation of the message (8, 9). Similar mechanisms seem to operate in neurons (10, 11) and possibly in other somatic cells (12).Because the length of the poly(A) tail is important for its function, it is not surprising that poly(A) tails are generally synthesized with a defined length, which is species-specific, ∼70–90 nucleotides in Saccharomyces cerevisiae (13, 14) and ∼250 nucleotides in mammalian cells (15). Subtle differences between newly made poly(A) tails of different mRNAs have been described (13), and there is even a class of mRNAs that never receives more than an oligo(A) tail (16, 17). However, the heterogeneous length distribution seen in the steady-state mRNA population is the result of cytoplasmic shortening starting from a relatively well defined initial tail length; heterogeneity of tail length reflects age differences of the mRNA molecules. The oligo(A) tails present on inactive mRNAs in oocytes or embryos are also generated by shortening of full-length tails made in the cell nucleus (18).The poly(A) tail is added during 3′ end processing of mRNA precursors in the cell nucleus (19–21). This reaction consists of two steps: an endonucleolytic cleavage followed by the addition of the poly(A) tail to the upstream cleavage product. Whereas a large protein machinery of some 20 or more polypeptides (22) is required for the cleavage reaction, subsequent polyadenylation has much simpler protein requirements. In the mammalian system, it can be reconstituted from three proteins: poly(A) polymerase, the enzyme catalyzing primer-dependent polymerization of AMP using ATP as a precursor (23–25); the cleavage and polyadenylation specificity factor (CPSF),⁶ which binds the cleavage and polyadenylation signal AAUAAA (26, 27); and the nuclear poly(A)-binding protein (PABPN1), which binds the growing poly(A) tail (28, 29). Note that PABPN1 is distinct from the family of cytoplasmic poly(A)-binding proteins (30). Roles of poly(A) polymerase and CPSF in polyadenylation in vivo have been most clearly demonstrated by genetic analysis of the orthologues in S. cerevisiae (21, 31). PABPN1 has no functional orthologue in budding yeast (32); its function in polyadenylation has been confirmed in mammalian cells (33) and in Drosophila (34).Whereas PABPN1 and poly(A) polymerase are monomeric proteins, CPSF is a hetero-oligomer, which has not yet been reconstituted from recombinant proteins (22, 26, 35–40). Poly(A) polymerase on its own is barely active because of a low affinity for its RNA substrate and thus acts distributively, i.e. it dissociates from the RNA after each polymerization step, and presumably often before it has incorporated any nucleotide; the enzyme also has no significant sequence specificity and will elongate any RNA with a free 3′ OH (24). Both CPSF and PABPN1 enhance the activity of the polymerase by recruiting the enzyme to its substrate through direct interactions (38, 41). Sequence specificity of poly(A) addition reflects the RNA binding specificities of the two stimulatory factors: CPSF recruits the polymerase to RNAs containing the AAUAAA sequence in the vicinity of their 3′ ends (24, 42, 43), and PABPN1 recruits the enzyme to substrate RNAs carrying a terminal oligo(A) tract (29). Each factor alone endows the polymerase with modest processivity, such that it can incorporate maybe two to five nucleotides before dissociating (44). RNAs containing both the AAUAAA sequence and an oligo(A) tail and thus resembling intermediates of the polyadenylation reaction support a cooperative or synergistic stimulation of poly(A) polymerase by both CPSF and PABPN1. Under these conditions, addition of the poly(A) tail occurs in a processive manner, i.e. without intermittent dissociation of the protein complex from its substrate RNA (29, 44).Interestingly, the reconstituted polyadenylation reaction also shows proper length control, generating poly(A) tails of the same length as seen in vivo; tails grow to a relatively well defined length of 250–300 nucleotides in a rapid, processive reaction (29, 44). Length control is due to termination of this processive elongation; extension beyond 250 A residues is largely distributive and therefore slow (45). These kinetics of in vitro poly(A) tail synthesis are fully consistent with the in vivo kinetics derived from pulse-labeling studies (46). In vitro, poly(A) tail elongation rates beyond 250 A residues are similar when either CPSF or PABPN1 or both are present. In other words, substrates with long poly(A) tails no longer support the cooperative stimulation of poly(A) polymerase by both CPSF and PABPN1 that is the basis of processive elongation (45). The termination of processive elongation must be mediated by a change in the RNA-protein complex that remains to be defined. When RNAs carrying poly(A) tails of different lengths are used as substrates for polyadenylation, the tails are always elongated processively to 250 nucleotides, independently of the initial length, whereas extension of a tail of 250 or more nucleotides in length is slow and distributive from the start of the reaction. Thus, poly(A) tail length control is based on some kind of AMP residue counting or length measurement, not on a kinetic mechanism (45).In this paper, we address the two problems outlined above: first, how does the polyadenylation complex change to terminate processive poly(A) tail elongation, and second, how is the length of the tail measured? We provide evidence that PABPN1 is the active component in the mechanism of length control. The protein promotes the interaction between CPSF and poly(A) polymerase when bound to a short poly(A) tail. PABPN1 no longer promotes or even actively disrupts this interaction when bound to a poly(A) tail of 250 nucleotides or longer and thereby terminates the cooperative, processive elongation reaction in a poly(A) tail length-dependent manner. Only poly(A) sequences are counted as part of the tail. Because this reflects the binding specificity of PABPN1 and because disruption of the CPSF-poly(A) polymerase interaction requires complete coverage of the poly(A) tail by this protein, PABPN1 is also the protein that measures the length of the tail.     

Mapping of the Functional Boundaries and Secondary Structure of the Mouse Mammary Tumor Virus Rem-responsive Element

Mouse mammary tumor virus (MMTV) is a complex retrovirus that encodes at least three regulatory and accessory proteins, including Rem. Rem is required for nuclear export of unspliced viral RNA and efficient expression of viral proteins. Our previous data indicated that sequences at the envelope-3′ long terminal repeat junction are required for proper export of viral RNA. To further map the Rem-responsive element (RmRE), reporter vectors containing various portions of the viral envelope gene and the 3′ long terminal repeat were tested in the presence and absence of Rem in transient transfection assays. A 476-bp fragment that spans the envelope-long terminal repeat junction had activity equivalent to the entire 3′-end of the mouse mammary tumor virus genome, but further deletions at the 5′- or 3′-ends reduced Rem responsiveness. RNase structure mapping of the full-length RmRE and a 3′-truncation suggested multiple domains with local base pairing and intervening single-stranded segments. A secondary structure model constrained by these data is reminiscent of the RNA response elements of other complex retroviruses, with numerous local stem-loops and long-range base pairs near the 5′- and 3′-boundaries, and differs substantially from an earlier model generated without experimental constraints. Covariation analysis provides limited support for basic features of our model. Reporter assays in human and mouse cell lines revealed similar boundaries, suggesting that the RmRE does not require cell type-specific proteins to form a functional structure.Mouse mammary tumor virus (MMTV)³ has multiple regulatory and accessory genes (1, 2). The known accessory genes specify a dUTPase (3), which is believed to be involved in retroviral replication in non-dividing cells (4), as well as superantigen (Sag). Sag is a transmembrane glycoprotein that is involved in the lymphocyte-mediated transmission of MMTV from maternal milk in the gut to susceptible epithelial cells in the mammary gland (5, 6). The Sag protein expressed by endogenous (germline) MMTV proviruses has been reported to provide susceptibility to infection by exogenous MMTVs or the bacterial pathogen, Vibrio cholerae (7). These results suggest a role for MMTV Sag in the host innate immune response.MMTV recently was shown to be a complex retrovirus (1). Complex retroviruses encode RNA-binding proteins that facilitate nuclear export of unspliced viral RNA by using a leucine-rich nuclear export sequence (8), which binds to chromosome region maintenance 1 (Crm1)(9), whereas simple retroviruses have a cis-acting constitutive transport element that directly interacts with components of the Tap/NXF1 pathway (10). Similar to other complex retroviruses, MMTV encodes a Rev-like protein, regulator of export/expression of MMTV mRNA (Rem) (1). Rem is translated from a doubly spliced mRNA into a 33-kDa protein that contains nuclear and nucleolar localization signals as well as a predicted RNA-binding motif and leucine-rich nuclear export sequence (1, 2). Our previous experiments indicated that Rem affects export of unspliced viral RNA, and a reporter vector that relies on luciferase expression from unspliced RNAs has increased activity in the presence of Rem (1). Sequences at the MMTV envelope-long terminal repeat (LTR) junction were required within the vector for Rem-induced expression, suggesting that the LTR contains all or part of the Rem-responsive element (RmRE). Very recently, Müllner et al. (11) identified a 490-nt region spanning the MMTV envelope-3′ LTR region, which was predicted to form a highly structured RNA element. This element confers Rem responsiveness on heterologous human immunodeficiency virus type 1 (HIV-1)-based plasmid constructs in transfection experiments.Experiments using other retroviral export proteins have demonstrated considerable variation in the size of the response elements. A minimal Rev-responsive element (RRE) in the human immunodeficiency virus type 1 (HIV-1) genomic RNA is 234 nt, the human T-cell leukemia virus Rex-responsive element is 205 nt (12–14), whereas the Rec-responsive element (RcRE; also known as the K-RRE) of human endogenous retrovirus type K is 416 to 429 nt (15, 16). Most response elements are confined to the 3′-end of their respective retroviral genomes (either to the envelope or LTR regions) (14, 15), but 5′ Rev-response elements also have been identified (17). Studies indicate that the secondary structure is a critical factor for proper function of retroviral response elements (18), and that multiple stem-loops are required. Export proteins multimerize on these elements to allow activity (19).In the current study, we have used deletion mutations within a reporter vector based on the 3′-end of the MMTV genome to define a 476-nt element necessary for maximum Rem responsiveness. This element spans the envelope-LTR junction of the MMTV genome as previously reported (1). However, a secondary structure model generated using digestions of the RmRE by RNases V1, T1, and A as experimental constraints differs significantly from the published structure (11) and more closely resembles complex retroviral response elements. Transfection experiments indicated that the MMTV RmRE could function in both mouse and human cells, suggesting that conserved cellular proteins interact with Rem.     

The 3��-Flap Pocket of Human Flap Endonuclease 1 Is Critical for Substrate Binding and Catalysis

Flap endonuclease 1 (FEN1) proteins, which are present in all kingdoms of life, catalyze the sequence-independent hydrolysis of the bifurcated nucleic acid intermediates formed during DNA replication and repair. How FEN1s have evolved to preferentially cleave flap structures is of great interest especially in light of studies wherein mice carrying a catalytically deficient FEN1 were predisposed to cancer. Structural studies of FEN1s from phage to human have shown that, although they share similar folds, the FEN1s of higher organisms contain a 3′-extrahelical nucleotide (3′-flap) binding pocket. When presented with 5′-flap substrates having a 3′-flap, archaeal and eukaryotic FEN1s display enhanced reaction rates and cleavage site specificity. To investigate the role of this interaction, a kinetic study of human FEN1 (hFEN1) employing well defined DNA substrates was conducted. The presence of a 3′-flap on substrates reduced K_m and increased multiple- and single turnover rates of endonucleolytic hydrolysis at near physiological salt concentrations. Exonucleolytic and fork-gap-endonucleolytic reactions were also stimulated by the presence of a 3′-flap, and the absence of a 3′-flap from a 5′-flap substrate was more detrimental to hFEN1 activity than removal of the 5′-flap or introduction of a hairpin into the 5′-flap structure. hFEN1 reactions were predominantly rate-limited by product release regardless of the presence or absence of a 3′-flap. Furthermore, the identity of the stable enzyme product species was deduced from inhibition studies to be the 5′-phosphorylated product. Together the results indicate that the presence of a 3′-flap is the critical feature for efficient hFEN1 substrate recognition and catalysis.In eukaryotic DNA replication and repair, various bifurcated nucleic acid structure intermediates are formed and must be processed by the appropriate nuclease. Two examples of biological processes that create bifurcated DNA intermediates are Okazaki fragment maturation (1, 2) and long patch excision repair (3). In both models, a polymerase executes strand-displacement synthesis to create a double-stranded DNA (dsDNA)⁶ two-way junction from which a 5′-flap structure protrudes. The penultimate step of both pathways is the cleavage of this flap structure to create a nicked DNA that is then ligated. Because the bifurcated DNA structures that are formed in the aforementioned processes can theoretically occur anywhere in the genome, the nuclease associated with the cleavage of 5′-flap structures in eukaryotic cells, which is called flap endonuclease 1 (FEN1), must be capable of cleavage regardless of sequence. Therefore, FEN1 nucleases, which are found in all kingdoms of life (4), have evolved to recognize substrates based upon nucleic acid structure and strand polarity (5, 6).The Okazaki fragment maturation pathway of yeast has become a paradigm of eukaryotic lagging strand DNA synthesis. In the yeast model, bifurcated intermediates with large single-stranded DNA (ssDNA) 5′-flap structures are imprecisely cleaved by DNA2 in a replication protein A -dependent manner (7). Subsequent to the DNA2 cleavage, Rad27 (yeast homologue of FEN1) cleaves precisely to generate an intermediate suitable for ligation (2). The recent discovery that human DNA2 is predominantly located in mitochondria in various human cell lines (8, 9) suggests that hFEN1 is the paramount 5′-flap endonuclease in the nuclei of human cells. This observation potentially provides a plausible rationale for why deletion of RAD27 (yeast FEN1 homologue) is tolerated in Saccharomyces cerevisiae (10), whereas deletion of FEN1 in mammals is embryonically lethal (11). Recent models wherein mice carrying a mutation (E160D) in the FEN1 gene, which was shown in vitro to alter enzymatic properties (12), have demonstrated that FEN1 functional deficiency in mice (S129 and Black 6) increases the incidence of cancer, albeit different types presumably due to genetic background (13, 14). Thus, the function of mammalian FEN1 in vivo is vital to the prevention of genomic instability. In addition to its importance in the nucleus, hFEN1 has recently been detected in mitochondrial extracts (15, 16) and implicated in mitochondrial long patch base excision repair (15). Considering the pivotal roles of hFEN1 in DNA replication and repair, it is of interest to understand how hFEN1 and homologues achieve substrate and scissile phosphate selectivity in the absence of sequence information.Since its initial discovery as a nuclease that completes reconstituted Okazaki fragment maturation (17) and subsequent rediscovery as a 5′-flap-specific nuclease (DNaseIV) from bacteria (18), mouse (19), and HeLa cells (20), FEN1 proteins ranging from phage to human have been studied biochemically, computationally, and structurally (5, 6, 21). Biochemical characterizations of FEN1 proteins from various organisms have shown that this family of nucleases can perform phosphodiesterase activity on a wide variety of substrates; however, the efficiency of catalysis on various substrates differs among the species. For instance, phage FEN1s prefer pseudo-Y substrates (22, 23), whereas the archaeal and eukaryotic FEN1s prefer 5′-flap substrates (21, 24, 25), which have two dsDNA domains, one upstream and downstream of the site of cleavage, and a 5′-ssDNA protrusion (Fig. 1A). Primary sequence analysis indicates that FEN1 proteins share characteristic N-terminal (N) and Intermediate (I) “domains,” which harbor the highly conserved carboxylate residues that bind the requisite divalent metal ions (26–28). Structural studies of FEN1 nucleases from phage to humans (22, 29–36), have shown that the N and I domains comprise a single nuclease core domain consisting of a mixed, six- or seven-stranded β-sheet packed against an α-helical structure on both sides. The α-helices on either side of the β-sheet are “bridged” by a helical arch that spans the active site groove (supplemental Fig. S1). On one side of the β-sheet, the α-helical bundle (αb1) creates the floor of the active site and a DNA binding motif (helix-3-turn-helix) (32). Similarly, the opposite α-helical bundle (αb2) has also been observed to interact with DNA (35). Based on site-directed mutagenesis studies with T5 phage FEN1 (T5FEN1) (37) and hFEN1 (38, 39), and crystallographic studies of T4 phage FEN1 (T4FEN1) (22) and Archaeoglobus fulgidus FEN1 (aFEN1) (35) in complex with DNA, a general model for how FEN1 proteins recognize flap DNA has emerged. The helix-3-turn-helix motif is involved in downstream dsDNA binding, whereas the upstream dsDNA domain is bound by αb2. The helical arch is likely involved in 5′-flap binding (22).Open in a separate windowFIGURE 1.Secondary structure schematics of hFEN1 substrates. A, illustration of a general flap substrate created using a bimolecular approach whereby a template strand (T-strand), which partially folds into a hairpin, anneals with the duplex strand (d-strand). The T-strand hairpin creates the upstream dsDNA domain, whereas the d-strand base pairs with the T-strand to create the downstream dsDNA domain. The flap or any other structure is created by addition of nucleotides to the 5′-end of the d-strand. The interface between the upstream and downstream dsDNA domains may be viewed as a derivative of a two-way junction (74). Annealing of either the F(5), E, or G(15) d-strands with the T3F T-strand results in the formation of a (B) double flap substrate (Flap of 5-nt d-strand paired with a Template with a 3′-Flap, F(5)·T3F), C, exonuclease substrate with a 3′-extrahelical nucleotide (EXO d-strand paired with a Template with a 3′-Flap, E·T3F), and a D, fork-GEN substrate with a 3′-extrahelical nucleotide and a 15-nt ssDNA gap capped by a 23-nt hairpin structure (fork-Gap of 15-nt d-strand paired with a Template with a 3′-Flap, G(15)·T3F). E, annealing the F(5) d-strand with the T oligonucleotide creates a single flap (Flap of 5-nt d-strand paired with a Template, F(5)·T).Unlike phage FEN1s, studies of FEN1s from eubacterial (40), archaeal (21), and eukaryotic origins (41) have shown that the addition of a 3′-extrahelical nucleotide (3′-flap) to the upstream duplex of a 5′-flap substrate results in a rate enhancement and an increase in cleavage site specificity. Moreover, substrates possessing a 3′-flap, which mimic physiological “equilibrating flaps,” were cleaved exactly one nucleotide into the downstream duplex, thereby resulting in 5′-phosphorylated dsDNA product that was a suitable substrate for DNA ligase I (21, 41). As postulated by Kaiser et al. (21), the structure of an archaeal FEN1 in complex with dsDNA with a 3′-overhang showed that the protein contains a cleft adjacent to the upstream dsDNA binding site that binds the 3′-flap by means of van der Waals and hydrogen bonding interactions with the sugar moiety (35). Once the residues associated with 3′-flap binding were identified, sequence alignment analyses showed that the amino acid residues in the 3′-flap binding pocket are highly conserved from archaea to human. Furthermore, mutation of the conserved amino acid residues in the 3′-flap binding pocket of hFEN1 resulted in reduced affinity for and cleavage specificity on double flap substrates (42). Although the effects of the addition of a 3′-flap to substrates on hFEN1 catalysis are known qualitatively, a detailed understanding of the relationship between changes in catalytic parameters and rate enhancement by the presence of a 3′-flap is unknown. Here, we describe a detailed kinetic analysis of hFEN1 using four well characterized DNA substrates and show that the presence of a 3′-flap on a substrate not only contributes to substrate binding (42), but also increases multiple and single turnover rates of reaction in the presence of near physiological monovalent salt concentrations. We also demonstrate that, like T5FEN1, hFEN1 is rate-limited by product release, and thus multiple turnover rates at saturating concentrations of substrate are predominantly a reflection of product release and not catalysis as was previously concluded (39). Furthermore, this study provides insight into the mechanism of hFEN1 substrate recognition.