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. Author manuscript; available in PMC: 2010 Jan 26.
Published in final edited form as: Mol Cell. 2000 Mar;5(3):479–488. doi: 10.1016/s1097-2765(00)80442-6

Interaction between a Poly(A)-Specific Ribonuclease and the 5′ Cap Influences mRNA Deadenylation Rates In Vitro

Min Gao 1, David T Fritz 1, Lance P Ford 1,†, Jeffrey Wilusz 1,*
PMCID: PMC2811581  NIHMSID: NIHMS168313  PMID: 10882133

Summary

We have used an in vitro system that reproduces in vivo aspects of mRNA turnover to elucidate mechanisms of deadenylation. DAN, the major enzyme responsible for poly(A) tail shortening in vitro, specifically interacts with the 5′ cap structure of RNA substrates, and this interaction is greatly stimulated by a poly(A) tail. Several observations suggest that cap-DAN interactions are functionally important for the networking between regulated mRNA stability and translation. First, uncapped RNA substrates are inefficiently deadenylated. Second, a stem-loop structure in the 5′ UTR dramatically reduces deadenylation by interfering with cap-DAN interactions. Third, the addition of cap binding protein eIF4E inhibits deadenylation in vitro. These data provide insights into the early steps of substrate recognition that target an mRNA for degradation.

Introduction

A 7-methyl-guanosine cap structure at the 5′ end and a 3′ poly(A) tail is a defining characteristic of mammalian mRNA. These terminal mRNA modifications occur cotranscriptionally (Ho and Shuman, 1999), presumably to protect the transcript from exonucleases (Furuichi et al., 1977). Terminal modifications have a broad influence on many aspects of gene expression. The mRNA cap, in association with two nuclear cap binding proteins (Izaurralde et al., 1994; Wilson et al., 1999), influences the efficiency of pre-mRNA splicing (Izaurralde et al., 1994), 3′ end formation/polyadenylation (Cooke and Alwine, 1996; Flaherty et al., 1997), and nucleo-cytoplasmic transport (Izaurralde et al., 1995). In the cytoplasm, the cap structure serves as the binding site for the eIF4F complex of translation initiation factors (Jackson and Wickens, 1997). The major cytoplasmic cap binding protein, eIF4E (Sonenberg and Gingras, 1998), associates with eIF4G and eIF4A to form a complex that serves as the nucleation site for eIF3 and the 43S complex involved in the early stages of ribosome assembly on mRNAs. The activity of eIF4E may be regulated during cell growth and differentiation by phosphorylation (Pyronnet et al., 1999) as well as by eIF4E binding proteins (Gingras et al., 1999). The 5′ cap structure, therefore, is an important recognition element for many posttranscriptional aspects of gene expression.

The 3′ poly(A) tail, in association with specific poly(A) binding protein(s) (PABP), also plays a broad role in gene expression. Polyadenylation influences the splicing efficiency of the terminal exon (Niwa and Berget, 1991) as well as the process of transcription termination (Yonaha and Proudfoot, 1999). Curiously, the poly(A) tail stimulates both cap-dependent and cap-independent translation initiation (Gallie, 1991; Sachs et al., 1997; Preiss et al., 1998). The translational efficiency of many mRNAs during development is directly regulated by changes in length of the poly(A) tail (Richter, 1999). The 3′ poly(A) tail, therefore, plays an active role in the cytoplasmic fate of an mRNA.

The observation that both the cap and poly(A) tail affect translation initiation suggested that they may interact physically as well as functionally. Consistent with this hypothesis, poly(A) appears to stimulate cap-dependent translation through a mechanism involving protein–protein interactions between PABP, eIF4G, and the cap binding protein eIF4E (Tarun and Sachs, 1997; Imataka et al., 1998). These interactions generate a close physical association between the 5′- and 3′- ends of an mRNA, resulting in the observation of circular mRNPs (Wells et al., 1998). Aside from perhaps being a very effective way to recycle ribosomes on an individual mRNA for protein synthesis (Christensen et al., 1987), interactions between the ends of an mRNA may play an important role in additional aspects of mRNA metabolism. Understanding the communication of the 5′ cap structure with other regions of both precursor and mature forms of an mRNA will provide important insights into the regulation of posttranscriptional events.

Relative rates of mRNA turnover play an important role in determining the steady-state level and inducibility of transcripts (Ross, 1995; Caponigro and Parker, 1996; Jacobson and Peltz, 1996). Communication between the cap and distal regions of the mRNA also appears to be required during mRNA turnover. In Saccharomyces cerevisae, turnover of most mRNAs is initiated by deadenylation (Decker and Parker, 1993). Generation of a short oligo-A tail then serves as a signal for decapping of the 5′ end of the transcript in the XRN1-dependent pathway of mRNA turnover (Muhlrad et al., 1994). The presence of inappropriately placed nonsense codons in an mRNA can also activate deadenylation-independent decapping in yeast (Weng et al., 1997; Hilleren and Parker, 1999). Whether or not the cap structure plays a central role in the early events of mRNA turnover in mammalian cells is unclear. Mammalian mRNAs are usually deadenylated before turnover of the body of the transcript (Shyu et al., 1991; Wilson and Treisman, 1998). While indirect evidence suggests that decapping of mammalian mRNAs may occur (Couttet et al., 1997) and a mouse XRN1 homolog has been identified (Bashkirov et al., 1997), mechanistic aspects of mammalian mRNA turnover that occur after deadenylation remain to be elucidated.

We have recently developed an in vitro system using S100 cytoplasmic extracts from HeLa cells that faithfully reproduces in vivo aspects of mRNA stability (Ford et al., 1999). Capped and polyadenylated RNA substrates are deadenylated prior to turnover of the body of the transcript by independent deadenylation and exonucleolytic activities. AU-rich elements (AREs) found in the 3′ untranslated regions of many short-lived mRNAs, including those encoding growth factors and cytokines (Caput et al., 1986), dramatically shorten RNA half-lives in vivo (Shaw and Kamen, 1986; Chen and Shyu, 1995; Chen et al., 1995). Subsequently, the rate of both deadenylation and degradation of RNA substrates in vitro can be significantly stimulated by the inclusion of an ARE. Finally, ARE-binding proteins influence the turnover rates of RNAs in a similar fashion both in vivo and in vitro. HuR protein, for example, binds to and stabilizes ARE-containing transcripts in both transfection assays and in S100 extracts (Jain et al., 1997; Fan and Steitz, 1998; Levy et al., 1998; Peng et al., 1998; Ford et al., 1999). Clearly, our in vitro system has significant potential to yield important clues concerning mechanistic aspects of mammalian mRNA stability.

The identification of the enzymes responsible for deadenylation of eukaryotic mRNAs has been elusive, although several candidate exonucleases with selectivity for poly(A) tracts have been reported (Boeck et al., 1996; Virtanen and Astrom, 1997; Korner et al., 1998). PAN2, a Pab1p-dependent deadenylase, has been characterized from S. cerevisiae (Brown et al., 1996). However, genetic studies indicated that pan2 mutants had only minimal defects on deadenylation (Brown and Sachs, 1998). Recent evidence suggests that PAN2 may be involved in poly(A) tail length regulation in the nucleus (Brown and Sachs, 1998). In human cells, a poly(A)-specific 3′ to 5′ exonuclease (DAN) has recently been shown to have striking similarity with members of the RNase D family of exonucleases (Korner et al., 1998). Curiously, DAN is related to the enzymatic activity that catalyzes default deadenylation during Xenopus oocyte meiotic maturation. Finally, the relationship of poly(A) binding proteins to deadenylation enzymes appears to be different for the mammalian deadenylase than for the yeast PAN2 enzyme. While Pab1p stimulates PAN2, the association of poly(A) binding proteins with human or Xenopus mRNAs appears to protect the transcript from deadenylation (Bernstein et al., 1989; Brown et al., 1996; Wormington et al., 1996; Ford et al., 1997).

Communication between the 5′ and 3′ ends may influence the deadenylation of mRNAs. Furthermore, the role of 5′–3′ interactions in the process of mRNA turnover may be intertwined with the functional interplay between mRNA termini observed in translation (Gallie, 1998). Numerous experiments have shown that translation can influence the efficiency of mRNA turnover in both yeast and mammalian systems (Ross, 1995). The addition of protein synthesis inhibitors, mutation of the initiation codon, or insertion of stem-loop structures in the 5′ untranslated region have all been shown to stabilize mRNAs (Aharon and Schneider, 1993; Schiavi et al., 1994; Ross, 1997). Translation, however, is not absolutely required to observe regulated mRNA stability (Chen et al., 1995; Fan et al., 1997). Perhaps a key aspect in the functional association between translation and mRNA stability involves a competition between translation initiation factors and mRNA degradative enzymes for communication with both the cap and poly(A) tail.

In this study, we have used our in vitro stability system to define key aspects of mRNA deadenylation, the first regulated step in mammalian mRNA turnover. We have demonstrated that DAN, a powerful poly(A)-specific exonuclease (Korner et al., 1998), is responsible for deadenylation in the system. Curiously, the deadenylase directly and specifically interacts with the 5′ cap structure on RNA substrates. The interaction of DAN with the 5′ cap is greatly stimulated by the presence of a poly(A) tail on RNA substrates. Cap–deadenylase interactions influence the efficiency of deadenylation in vitro, and can be inhibited by both 5′ stem-loop structures in the RNA substrate as well as the cap binding protein eIF4E. These data provide important insights into the regulation of mRNA deadenylation as well as the interplay between mRNA translation and turnover.

Results

DAN Is the Major Deadenylase in S100 Extracts

Deadenylation precedes turnover of the body of RNA substrates in our in vitro stability system (Ford et al., 1999). Poly(A) shortening occurs in the system due to a poly(A)-specific 3′ to 5′ exonuclease and is not the result of the action of random exonucleases. The first step in determining the mechanism of regulated deadenylation in vitro is the identification of the deadenylase enzyme. Candidate 3′ to 5′ exonucleases that demonstrate preferential activity on poly(A) tracts have been previously identified using a biochemical approach (Virtanen and Astrom, 1997). The Wahle and Wormington laboratories recently purified, cloned, and prepared antibodies to a candidate deadenylase (DAN) of ∼74 kDa that contains homology to the RNase D family of 3′ to 5′ exonucleases (Korner et al., 1998). Preimmune and DAN-specific rabbit antisera were used to immunodeplete HeLa cytoplasmic S100 extracts prior to their use in in vitro assays. Western blot analysis demonstrated that most, but not all, of DAN protein was specifically removed by immunodepletion (Figure 1A). As seen in Figure 1B, immunodepletion with α-DAN antisera specifically reduced deadenylation of a model polyadenylated substrate RNA (SVARE-A60). The deadenylation rate of all polyadenylated RNA substrates was dramatically reduced in the in vitro system using several independent extracts that were immunodepleted with α-DAN antisera (data not shown). The reason that deadenylation was not totally inhibited is due to difficulties in removing 100% of DAN activity from S100 extracts by immunodepletion (Figure 1A). Finally, the addition of purified, recombinant DAN produced in E. coli restored efficient deadenylase activity to immunodepleted extracts (Figure 1C). We conclude that DAN is the major deadenylase that is active in the in vitro system.

Figure 1. Immunodepletion Using α-DAN Antibodies Specifically Inhibits Deadenylation in the In Vitro System.

Figure 1

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(A) Specific immunodepletion of DAN from S100 extracts. The level of DAN protein was determined by Western blotting using α-DAN polyclonal antisera in S100 extracts before (lane Ext) or after immunodepletion using α-DAN antisera (lane α-DAN) or rabbit preimmune sera (lane Preimm). Arrows to the right indicate the position of DAN and the heavy chain of IgG (which is present due to the antisera used for immunodepletion).

(B) SVARE-A60 RNA was incubated in the in vitro system using extracts that had been immunodepleted using either preimmune rabbit serum (Pre-immune lanes) or α-DAN antisera (α-DAN lanes). Reaction mixtures were incubated for the indicated times and products were analyzed on a 5% acrylamide gel containing 7 M urea.

(C) Recombinant DAN restores efficient deadenylation to immunodepleted extracts. GemARE-A60 RNA (Input lane) was inefficiently deadenylated after 30 min in the in vitro system using extracts that were immunodepleted using α-DAN antisera (lane 0). The addition of the indicated amounts (in ng) of purified recombinant DAN protein restored efficient deadenylation. The positions of the input GemARE-A60 RNA and the fully deadenylated intermediate are indicated on the left.

Deadenylase Specifically Interacts with the 5′ Cap Structure on Polyadenylated RNA Substrates

A variety of physical and functional interactions have been recently noted between the 5′ and 3′ ends of mRNAs. The poly(A) tail influences cap-dependent translation initiation (Sachs et al., 1997) and cap binding protein eIF4E interacts with PABP through eIF4G (Imataka et al., 1998; Wells et al., 1998). Furthermore, mRNA deadenylation influences decapping of mRNAs in S. cerevisiae (Decker and Parker, 1993). Because communication between the 5′ and 3′ ends clearly influences other aspects of mRNA metabolism (Gallie, 1998), we tested whether the cap structure could influence deadenylation through a titratable interaction. Unlabeled 7meGpppG was added in increasing amounts to the in vitro system and deadenylation was monitored by gel electrophoresis. As seen in Figure 2A, increasing amounts of 7meGpppG inhibited deadenylation of a model capped and polyadenylated RNA substrate (GemARE-A60), while GpppG had no effect even at the highest concentration tested. Furthermore, 7meGpppG inhibited deadenylation of all RNA substrates tested, regardless of whether or not they contained an AU-rich element (data not shown). A similar specific inhibition of deadenylation was observed by the addition of 7meGTP to reaction mixtures or by passing extracts over a 7meGTP-Sepharose column (data not shown). In conjunction with this chromatographic removal of deadenylation activity in extracts, Western blot analysis demonstrated that the DAN enzyme can be bound and purified on 7meGTP-Sepharose (Figure 2B). The addition of cap analogs or chromatography on 7meGTP-Sepharose, therefore, sequesters factor(s), most notably DAN, that is required for deadenylation in S100 extracts.

Figure 2. Cap Analog Inhibits Deadenylation in the In Vitro System and Directly Sequesters the Deadenylase.

Figure 2

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(A) GemARE-A60 RNA was incubated in the in vitro mRNA stability system for 30 min in the presence of the indicated amounts of 7meGpppG or GpppG. Reaction products were analyzed on a 5% acrylamide gel containing 7 M urea. The positions of the input GemARE-A60 RNA and the fully deadenylated intermediate are indicated on the left.

(B) DAN can be purified on 7meGTP-Sepharose. HeLa S100 extracts were mixed with 7meGTP-Sepharose beads and bound proteins were washed five times in buffer D before elution by denaturation. Samples of the input, wash, and eluted fractions were run on a 7.5% acrylamide/SDS gel and analyzed by Western blotting using α-DAN polyclonal sera.

We hypothesized that 7meGpppG may be blocking deadenylation by directly sequestering DAN itself. In order to test whether DAN can specifically interact with the cap structure present on our polyadenylated RNA substrates, RNAs were radiolabeled exclusively at the α-phosphate of the cap structure using guanylyltransferase and 32P-GTP. Cap-labeled transcripts were incubated in the in vitro stability system for 5 min to allow the assembly of protein-RNA complexes and then were irradiated with UV light. Following RNase treatment, proteins that were radiolabeled due to cross-linking to 32P-capped RNA oligomers were identified on SDS-acrylamide gels either before (Figure 3A, total lane) or after immunoprecipitation with preimmune or α-DAN antisera. As seen in Figure 3A, DAN cross-linked specifically to the cap-structure of GemARE-A60 RNA. In order to determine if the AU-rich instability element present in GemARE-A60 RNA was required for the deadenlyase to cross-link to the cap structure, a variant that lacks the AU-rich element (Gem-A60) was tested. As seen in Figure 3B, the cap of Gem-A60 RNA cross-linked to the deadenylase with similar efficiency as GemARE-A60 RNA. DAN cross-linked to the cap structure of all polyadenylated RNA substrates we tested, regardless of the size or internal sequence content of the substrate (Figures 3 and 5, other data not shown). In addition, we wished to determine if the DAN-cap interaction we have detected by UV cross-linking was indeed specific, especially since cap-labeled RNA substrates cross-linked to several proteins in our assays (Figure 3A, total lane). Competition experiments using unlabeled cap analog demonstrated that most of the protein-cap labeled substrate interactions detected in the total cross-linked lane of Figure 3A were not cap-specific (data not shown). These cross-linked proteins are likely detected due to their interaction with the body of the RNA substrate near, but not directly with, the 5′ cap structure. However, as seen in Figure 3C, DAN-cap interactions were readily competed by unlabeled 7meGpppG. The concentration of cold cap analog required to compete for cap-DAN interactions was similar to the amount required to inhibit deadenylation in vitro. Therefore, the deadenylase specifically interacts with the cap structure of RNA substrates in the in vitro system. Finally, purified recombinant DAN could also be cross-linked to cap-labeled RNA substrates (data not shown), suggesting that the enzyme directly interacts with the cap structure in the absence of accessory proteins.

Figure 3. Specific Interactions between the Deadenylase and the 5′ Cap of RNA Substrates Occur in a Poly.

Figure 3

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(A) Tail–Dependent Fashion All of the RNAs used in these experiments were exclusively radiolabeled at the α-P of the 5′ cap structure. (A) Cap-labeled GemARE-A60 RNA was incubated in the in vitro mRNA stability system for 5 min. RNA-protein complexes were covalently cross-linked by UV irradiation and reaction mixtures were treated with RNase. Proteins cross-linked to short radioactive RNA oligomers were analyzed on a 10% acrylamide/SDS gel before (Total lane) or after immunoprecipitation using either rabbit prebleed (Pre lane) or a rabbit polyclonal α-DAN sera (α-DAN lane).

(B) UV cross-linking analysis was performed using either GemARE-A60 or Gem-A60 RNAs as described above. Cross-linked proteins were immunoprecipitated using α-DAN antisera and analyzed on a 10% acrylamide/SDS gel.

(C) UV cross-linking analysis was performed using GemARE-A60 RNA as described in (A) in the absence (lane 0) or the presence of the indicated final concentration of 0.4 mM 7meGpppG (lane 5). Cross-linked proteins were immunoprecipitated using α-DAN antisera and analyzed on a 10% acrylamide/SDS gel.

(D) UV cross-linking analysis was performed using either GemARE-A60 (lane A60) or a deadenylated variant GemARE-A0 (lane A0) as described in (A). Cross-linked proteins were immunoprecipitated using α-DAN antisera and analyzed on a 10% acrylamide/SDS gel. The numbers on the left of each panel denote molecular weight markers and the arrow at the right denotes the position of cross-linked DAN.

Figure 5. The Insertion of a Stable Hairpin Structure near the 5′ End of an RNA Substrate Inhibits both Deadenylation and Cap-Deadenylase Interactions.

Figure 5

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(A) Diagrammatic representation of SVARE-A60 RNA and a derivative (SV-SL-ARE-A60 RNA) that contains a 20 base–hairpin structure located at position 4 from the 5′ end.

(B) SVARE-A60 RNA or SV-SL-ARE-A60 RNAs were incubated in the in vitro stability system for the times indicated. Reaction products were analyzed on a 5% acrylamide gel containing 7 M urea. The positions of the polyadenylated input RNAs (A60) and deadenylated intermediates (A0) are indicated at the left of each panel.

(C) UV cross-linking assays were performed using SVARE-A60 or SV-SL-ARE-A60 RNAs as described in Figure 5. Cross-linked proteins were immunoprecipitated using α-DAN antisera and analyzed on a 10% acrylamide/SDS gel.

Since DAN was first identified by its poly(A) substrate specificity (Korner et al., 1998), we next investigated whether a poly(A) tail was required for efficient DAN-cap interactions. Cap-labeled GemARE RNA derivatives were prepared that lacked a poly(A) tail (GemARE-A0) or contained 60 adenylates at the 3′ end (GemARE-A60), incubated in the in vitro stability system, and subjected to cross-linking/immunoprecipitation analysis as described above. As seen in Figure 3D, deadenylase-cap interactions were readily detected on polyadenylated transcripts, but DAN failed to efficiently cross-link to the cap of the GemARE-A0 RNA that lacked a poly(A) tail. We conclude that a poly(A) tail is required for the deadenylase to stably interact with the cap structure of RNA substrates in vitro.

The 5′ Cap Affects Deadenylation Efficiency

The next key question to address was the functional significance of cap–deadenylase interactions. If cap–DAN interactions affect deadenylase activity or kinetics, then the deadenylation rate of polyadenylated RNA substrates should be affected by removal of the 5′ cap. To address this, GemARE-A60 and SVARE-A60 RNAs were prepared with or without a cap structure at their 5′ ends and incubated in the in vitro stability system. As seen in Figures 4A and 4B, capped transcripts were rapidly deadenylated. Uncapped GemARE-A60 or SVARE-A60 RNAs, on the other hand, were deadenylated very inefficiently. Although some deadenylation was still observed on uncapped RNA substrates, the rate of poly(A) shortening was dramatically reduced. Therefore, the presence of a 5′ cap structure on RNA substrates stimulates deadenylation of the poly(A) tail.

Figure 4. Uncapped ARE-Containing RNAs Are Inefficiently Deadenylated In Vitro.

Figure 4

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(A) GemARE-A60 RNA with a 7meGppp 5′ cap (Capped lanes) or containing only a 5′ triphosphate (Uncapped lanes) were incubated in the in vitro stability system for the times indicated. Reaction products were analyzed on a 5% acrylamide gel containing 7 M urea. The positions of the polyadenylated input RNAs (A60) and deadenylated intermediates (A0) are indicated at the left of each panel.

In addition to outright removal of terminal modifications, other ways of interfering with 5′ cap–3′ poly(A) communication may also affect deadenylation rates. Previous data have demonstrated that the insertion of stem-loop structures in the 5′ untranslated region of mRNAs can dramatically stabilize some transcripts in vivo (Aharon and Schneider, 1993; Ross, 1995). While these results were originally interpreted as evidence that mRNA degradation is dependent on the process of translation, our data presented above suggest an alternative interpretation. Perhaps 5′ stem-loop structures can interfere with mRNA degradation by disrupting interactions between the deadenylase and the cap structure. To test this hypothesis, a variant of SVARE-A60 RNA was created that contained a 20 base hairpin structure beginning at position 4 from the 5′ end (Figure 5A). Capped, polyadenylated SVARE-A60 RNA and the variant containing the 5′ hairpin were incubated in the in vitro stability system. While SVARE-A60 RNA was efficiently deadenylated (Figure 5B, right side), the variant containing the 5′ hairpin structure was deadenylated with markedly reduced kinetics (Figure 5B, left side). A similar dramatic decrease in deadenylation efficiency was observed with a derivative of SVARE-A60 RNA that contained a hairpin structure 23 bases from the 5′ end (data not shown). Furthermore, UV cross-linking/immunoprecipitation analysis using cap-labeled transcripts showed that the variant containing the 5′ stem-loop structure blocked interaction of DAN with the cap structure (Figure 5C). S100 extracts, therefore, reproduce an in vivo observation with regard to the effect of 5′ stem-loop structures on RNA turnover. In addition, these data further suggest that cap-deadenylase interactions play a significant role in determining rates of poly(A) tail removal.

Protein-RNA interactions involving the 5′ cap structure may also influence 5′-3′ end communication. For example, cap binding proteins, such as the translation initiation factor eIF4E, may compete with the deadenylase for binding of the 5′ cap. In order to test this hypothesis, the effect of adding purified mouse eIF4E protein (Figure 6A) to the in vitro RNA stability system was monitored. As seen in Figure 6B, the addition of eIF4E protein inhibited deadenylation of capped RNA substrates in vitro. The addition of an equivalent amount of unrelated recombinant proteins (GST, hnRNP H) had no effect on deadenylation rates (data not shown). We conclude that eIF4E may influence RNA stability by prohibiting the association of the deadenylase with the mRNA cap structure. In summary, these observations suggest that at least a portion of the basis for the interplay between translation and mRNA turnover may lie in a competition between factors for association with the mRNA cap structure. Disruption of these interactions can possibly lead to changes in mRNA half-lives in vivo.

Figure 6. The Cap Binding Protein eIF4E Inhibits Deadenylation.

Figure 6

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(A) Coomassie-stained SDS gel of the eIF4E preparation demonstrates its purity.

(B) GemARE-A60 RNA was incubated in the in vitro RNA stability system in the presence (+eIF4E lanes) or absence (extract lanes) of 100 ng of recombinant cap binding protein for the indicated times. Reaction products were analyzed on a 5% acrylamide gel containing 7 M urea. The positions of the polyadenylated input RNAs (GemARE-A60) and deadenylated intermediates (GemARE-A0) are indicated at the left of each panel.

Discussion

This study suggests that communication between the 5′ and 3′ ends of transcripts plays a key role in determining the efficiency of mRNA turnover. DAN, the enzyme responsible for poly(A) shortening, interacts with the 5′ cap structure of mRNAs in a poly(A)-dependent fashion. This interaction is functionally significant as removal of the cap, blocking access to the cap by a nearby secondary structure, or addition of cap binding protein eIF4E as competitor, dramatically reduce deadenylation of RNA substrates. Since deadenylation is likely to be the rate-limiting step in the turnover of most mRNAs, these data suggest that cap-deadenylase interactions may also be an important factor in regulated mRNA decay.

Our data suggest a model for two separate pathways for the initial steps in deadenylation of mRNA substrates, as well as for the interplay between mRNA turnover and translation (Figure 7). Limited poly(A) tail shortening may occur throughout the lifetime of the transcript due to exonucleolytic trimming. The deadenylation activity addressed by this model is the relatively rapid poly(A) shortening that occurs in association with mRNA degradation. A variety of factors, including mRNA binding proteins and subcellular localization (Gray and Wickens, 1998), determine the translatability of the mRNA and, therefore, the pathway of assembly of deadenylation factors. Communication between the 5′ and 3′ ends of the transcript is proposed to be a key determinant of mRNA fate. The association of translation factors, such as eIF4E, eIF4G, and PABP with the mRNA influences communication between the termini, determining the pathway of deadenylation.

Figure 7. Protein-RNA Interactions Involving both the 5′ Cap and Poly(A) Tail Are Proposed to Play a Key Role in the Interplay between Translation and mRNA Deadenylation.

Figure 7

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A detailed discussion can be found in the text.

The association of cap binding protein, eIF4E, with the mRNA may be the rate-limiting step in determining the pathway for deadenylation of the transcript. eIF4E is present in substoichiometric amounts relative to the amount of mRNA in HeLa cells (Rau et al., 1996), and its association with mRNA cap structures is further regulated by eIF4E binding proteins (Gingras et al., 1999). In the translation factor independent pathway, the deadenylation apparatus assembles on the mRNA termini before eIF4E, eIF4G, and PABP can form a stable complex. AU-rich element mediated mRNA decay, which has been shown by some studies to be independent of translation (Chen et al., 1995; Fan et al., 1997), may occur through this pathway. AU-rich elements may promote the direct association of DAN with mRNAs, resulting in their rapid deadenylation and relatively short half-lives. Stability factors, such as HuR protein, can interact with AU-rich elements and prohibit the association of DAN with the mRNA, perhaps by promoting interaction of PABP with the poly(A) tail (Ma et al., 1997) or by competing for the binding of the factors responsible for instability to the AU-rich element (Fan and Steitz, 1998; Peng et al., 1998). The in vitro system that we have developed using cytoplasmic S100 extracts (Ford et al., 1999) likely reproduces this pathway of regulated mRNA deadenylation.

The alternative pathway for the assembly of the deadenylation apparatus on mRNAs begins with the association of translation factors with the 5′ and 3′ ends of the mRNA. Translatable mRNAs are likely to be found in circular forms due to interaction between PABP, eIF4G, and the cap binding protein eIF4E (Wells et al., 1998). These circular mRNPs would be refractory to deadenylation due to the inefficient association of DAN with the transcript since both the cap and poly(A) tail are masked by protein factors. The efficiency of cap-dependent translation is influenced by eIF4E, eIF4G, and PABP (Merrick and Hershey, 1996). These initiation factors attract eIF3 and the small ribosomal subunit to begin scanning the mRNA for the open reading frame and commencement of protein synthesis. The movement of the ribosome along the transcript may result in the transient disruption of the eIF4E-eIF4G-PABP complex with the mRNA termini, briefly exposing the 5′ and 3′ ends to the deadenylation apparatus. Although steric and topological forces clearly may transiently destabilize this terminal complex during translation, it is important to point out that the fate of the ribosome, eIF4E, eIF4G, and PABP following translation termination remains to be determined. Instability elements that require translation to reduce mRNA half-lives (Ross, 1995) may function by promoting the disruption of the eIF4E-eIF4G-PABP complex during translation. In addition, AU-rich elements and associated binding factors can also influence mRNA stability in this pathway by regulating the association of the deadenylation apparatus with the transiently exposed ends of the mRNA. In summary, protein-RNA interactions involving the terminal modifications of mRNAs may play a key role in networking the process of mRNA translation and turnover in the cell.

Several observations suggest that interfering with protein complexes involving mRNA termini may play a role in destabilizing transcripts. Disruption of poly(A) binding protein complexes by ribosome read-through into the Constant Spring variant of α-globin, for example, has been proposed as a destabilizing mechanism for this short-lived transcript (Weiss and Liebhaber, 1994). Consistent with a role for the cap in poly(A) tail shortening, yeast ceg-1 mutants that are defective in mRNA capping accumulate poly(A)+ mRNAs in an xrn1 strain (Schwer et al., 1998). Finally, PABP prevents the access of exonucleases to the poly(A) tail in vertebrates (Bernstein et al., 1989; Wormington et al., 1996; Ford et al., 1997).

Regulation of cap binding protein eIF4E appears to have a significant impact in cell biology. Both phosphorylation and eIF4E binding proteins have been proposed as regulators of the cap binding protein eIF4E through a number of signal transduction pathways (Gingras et al., 1999; Pyronnet et al., 1999). The aberrant growth and cellular transformation observed when eIF4E is over-expressed also demonstrate the importance of factor in cellular regulation (Lazaris-Karatzas and Sonenberg, 1992). Perhaps some of this regulation is mediated not only by the effects of eIF4E on translation, but also by the influence of eIF4E in regulating the deadenylation of mammalian mRNAs.

DAN is an apparently highly processive cytoplasmic enzyme that rapidly degrades poly(A) in a 3′ to 5′ manner (Korner et al., 1998). Immunodepletion experiments clearly demonstrate that the major deadenylase in S100 extracts at least shares an epitope with DAN, and most likely, is DAN itself. Direct addition of the α-DAN antisera to in vitro assays had no effect on deadenylation rates (data not shown). The identification of DAN as the major deadenylase makes it the first enzymatic activity that can be linked to the turnover of RNAs in our in vitro system.

The interaction of DAN with the cap structure is greatly stimulated by a poly(A) tract at the 3′ end of the RNA (Figure 3). The presence of poly(A) binding proteins such as PABP, therefore, may influence cap-deadenylase interactions. As noted earlier, omission of poly(A) competitor RNA from extracts results in the interaction of poly(A) binding proteins to the 3′ end of polyadenylated RNA substrates (Ford and Wilusz, 1999). In order to test whether the presence of poly(A) binding proteins inhibits cap-deadenylase interactions, RNA substrates that contain a 60 base poly(A) tail were incubated with S100 extracts in the presence or absence of 500 ng poly(A) competitor RNA. DAN cross-linked to the cap structure of GemARE-A60 RNA in both the presence and absence of poly(A) competitor RNA (data not shown). These data suggest that DAN can assemble on polyadenylated substrates and interact with the 5′ cap even in the presence of poly(A) binding proteins on the 3′ end. Perhaps terminal poly(A) regions that are not efficiently occluded by PABP serve as assembly sites for the deadenylase under these conditions.

In conclusion, we have used the regulated in vitro mRNA deadenylation/degradation system that we have recently developed to identify an interaction between the 3′ and 5′ ends of RNA substrates that has both mechanistic and regulatory implications for the process of mRNA deadenylation. This powerful biochemical approach should continue to yield mechanistic insights in the future. We are currently addressing questions concerning the fate of the cap structure in the turnover of mammalian mRNAs, the role of the trimeric PABP, eIF4G, and eIF4E complex in the regulation of mRNA turnover, and the identification of novel enzymatic and regulatory factors that impact on the process.

Experimental Procedures

RNAs

SVARE-A0 RNA, which contains the 34 base AU-rich element from TNF-α inserted into the BamHI-BclI fragment representing the 3′ portion of SV40 late mRNAs, was transcribed from HindIII linearized templates as previously described (Ford et al., 1999). SVARE-A60 RNA, a variant that contains a 60 poly(A) tract at its 3′ end, was prepared as previously described (Ford and Wilusz, 1999). Templates to synthesize SV-SL-ARE-A60, a variant that contains a 20 base–stem loop at its 5′ end, were prepared by PCR of pSVARE (Ford et al., 1999) using the primers 5′-CATACGATTTAGGTGACAC TATGAAAATTCCGTGTATACACGGAATTCGAGCTCGCCCGGGGA TCCAGAC and 5′-TACCTCGAGCACTC. Gem-A60 RNA, which contains sequences from the pGem4 polylinker region followed by 60 A residues, was prepared as previously described (Ford et al., 1999). GemARE-A60, a variant that contains the 34 base AU-rich element from TNF-α, was prepared by inserting the oligonucleotide 5′-ATT ATTTATTATTTATTTATTATTTATTTATTTA and the appropriate complement into the PstI and HindIII sites of pGem4. Transcription of HindIII linearized templates yields GemARE-A0 RNA. Sequences encoding a 60 base poly(A) tail were added to DNA using a ligation-PCR approach previously described (Ford et al., 1997).

RNAs were transcribed in vitro using SP6 polymerase as described previously (Wilusz and Shenk, 1988). Transcripts were radiolabeled at U residues by incorporating α-32P-UTP into the reaction mixtures. Transcripts were capped cotranscriptionally by the addition of 7meGpppG to reaction mixtures. To generate uncapped transcripts that contain a triphosphate at their 5′ ends, cap analog was omitted from the reaction mixtures. To prepare RNAs exclusively labeled at their cap structures, in vitro transcription reactions were performed in the absence of cap analog and radiolabeled nucleotides. Transcription products were then capped using recombinant vaccinia guanyltransferase and α-32P-GTP. All RNAs were purified on 5% acrylamide gels prior to use.

In Vitro mRNA Deadenylation/Degradation Assays

RNA substrates were incubated in the in vitro RNA stability system as described previously (Ford and Wilusz, 1999; Ford et al., 1999). Briefly, RNAs were incubated in the presence of S100 HeLa cell cytoplasmic extract, polyvinyl alcohol, ATP, phosphocreatine, and 500 ng of poly(A). Reactions were incubated at 30°C for the times indicated and products were analyzed on a 5% acrylamide gel containing 7 M urea.

To elucidate the role of DAN–cap interactions in deadenylation, extracts were immunodepleted using rabbit preimmune or α-DAN polyclonal antisera (Korner et al., 1998) by adding 3 μl of antisera and 1 μl of RNAsin to S100 extracts. Immune complexes were precipitated using protein A Sepharose and depleted extracts were used in the in vitro stability system as described above. Recombinant his-tagged DAN was purified from E. coli and added back directly to immunodepleted extracts at the levels indicated. For cap competition studies, 7meGpppG or GpppG were added directly to reaction mixtures. For 7meGTP-Sepharose purification of DAN, extracts were incubated with swelled beads in a batch mode, washed five times in Buffer D (10 mM HEPES [pH 7.9], 100 mM KCl, 1 mM EDTA, 20% glycerol, and 1 mM DTT), and bound proteins eluted by boiling in protein gel loading buffer. For eIF4E cap binding protein studies, 100 ng of recombinant mouse eIF4E purified in E. coli, obtained from C. Williams and S. Peltz, was added directly to reaction mixtures.

UV Cross-Linking/Immunoprecipitation/Western Blotting

UV cross-linking analysis was performed as described (Wilusz and Shenk, 1988). Briefly, 20–50 fmol of cap- or U-radiolabeled RNA was incubated in the in vitro RNA stability system for 5 min in the presence of EDTA (to prevent turnover and allow for accurate comparisons). Reaction mixtures were irradiated on ice for 10 min using a 15 W germicidal light. RNases A and T1 were added and proteins covalently attached to short radioactive RNA oligomers were analyzed on a 10% acrylamide gel containing SDS.

For analysis of UV cross-linked products by immunoprecipitation, 400 μl of NET2 buffer (50 mM Tris [pH 7.6], 150 mM NaCl, and 0.01% NP40) was added to reaction mixtures following RNase treatment and reaction mixtures were centrifuged for 4 min. Precleared samples were incubated on ice with 2–5 μl of specified rabbit polyclonal antisera for 1 hr. Antigen antibody complexes were collected on protein A-positive Staphylococcus aureus cells, washed five times in RIPA buffer (50 mM Tris [pH 7.6], 150 mM NaCl, 0.1% SDS, 1% NP40, and 0.5% deoxycholate), and immunoprecipitated cross-linked proteins were analyzed on a 10% acrylamide gel containing SDS. Polyclonal and preimmune antisera specific for DAN were obtained from M. Wormington (Korner et al., 1998).

Western blot analysis of proteins separated on 7.5% acrylamide SDS gels was performed using rabbit polyclonal DAN antisera at a 1:1000 dilution and detected using a goat anti-rabbit IgG secondary antibody and ECL reagents.

Acknowledgments

We wish to thank Michael Wormington and Elmar Wahle for the generous gift of α-DAN antisera and the DAN expression plasmid, Stuart Peltz and Carol Williams for providing eIF4E and guanyltransferase, Robert Donnelly for oligonucleotide synthesis, and M. Mathews, V. Bellofatto and C. Williams for critical comments. This work was supported by NIH grants GM56434 and CA80062 (to J. W.). L. P. F. was supported by cancer training grant CA09665.

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