Repositioning of the Reaction Intermediate within the Catalytic Center ...

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Feb 17, 2006 - transitions (reviewed in Staley and Guthrie [1998] and. Burge et al. ...... M.M.K., by grant BMC2002-00157 and program ''Ramón y Cajal''.
Molecular Cell 21, 543–553, February 17, 2006 ª2006 Elsevier Inc.

DOI 10.1016/j.molcel.2006.01.017

Repositioning of the Reaction Intermediate within the Catalytic Center of the Spliceosome Maria M. Konarska,1,* Josep Vilardell,2 and Charles C. Query3,* 1 The Rockefeller University New York, New York 10021 2 Centre de Regulacio´ Geno`mica 08003 Barcelona Spain 3 Department of Cell Biology Albert Einstein College of Medicine Bronx, New York 10461

Summary Conformational change within the spliceosome is required between the first catalytic step of pre-mRNA splicing, when the branch site attacks the 50 splice site (SS), and the second step, when the 50 exon attacks the 30 SS. Little is known, however, about repositioning of the reaction substrates during this transition. Whereas the 50 SS is positioned for the first step by pairing with the invariant U6 snRNA-ACAGAG site, we demonstrate that this pairing interaction must be disrupted to allow transition to the second step. We propose that removal of the branch structure from the catalytic center is in competition with binding of the 30 SS substrate for the second step. Changes in the relative occupancy of first and second step substrates at the catalytic center alter efficiency of the two steps of splicing, allowing use of suboptimal intron sequences and thereby altering substrate selectivity. Introduction Removal of introns from pre-mRNA is catalyzed by the spliceosome, a large, multicomponent complex whose assembly and function require multiple conformational transitions (reviewed in Staley and Guthrie [1998] and Burge et al. [1999]). Splicing catalysis consists of two consecutive transesterification reactions that directly involve three sites in the pre-mRNA: 50 SS, 30 SS, and branch site (BS). In the first step, the BS adenosine nucleophilically attacks the 50 SS, producing a lariat intermediate; in the second step, the 50 exon attacks the 30 SS, yielding spliced mRNA and lariat intron products (Figure 1A). Based on accumulated genetic and biochemical data, several models of the spliceosome catalytic center have been proposed in which a network of conserved RNA:RNA interactions involving snRNAs and pre-mRNA implicate U2 and U6 snRNAs in critical contacts with the splice sites and BS (Figure 1B; reviewed in Nilsen [1998], Burge et al. [1999], and Collins and Guthrie [2000]). These models are supported by the ability of a U2-U6 core structure to catalyze a reaction that resembles the first step of splicing (Valadkhan

*Correspondence: [email protected] (M.M.K.); query@ aecom.yu.edu (C.C.Q.)

and Manley, 2001). The 50 SS is positioned for first step catalysis in part by base pairing with the conserved ACAGAG region of U6 snRNA, and the region surrounding the BS is paired with the conserved GUAGUA sequence of U2 snRNA. In addition, U5 snRNA interacts with both exons, contributing to their positioning for catalysis (reviewed in Newman [1997]). Conformational change within the spliceosome is required between the first and second catalytic steps; this transition is assisted by Prp16, a member of the DExD/H family of ATPases ([Burgess et al., 1990]; reviewed in Rocak and Linder [2004]), which is required for the second step in vitro (Schwer and Guthrie, 1991). Little is known, however, about repositioning of the reaction substrates and products during this conformational change or, apart from exon interactions with U5, about substrate positioning for second step catalysis. Genetic contributions to current models of the catalytic center have relied predominantly on analysis of defects caused by one mutant allele and suppressed or exacerbated by another allele. Such compensatory effects of two independent mutations (in RNA, protein, or both) have often been interpreted as evidence for direct interactions between the affected factors. We have recently formulated a ‘‘two-state model’’ of the catalytic spliceosome, according to which structural conformations of the complex that define the two catalytic steps exist in kinetic competition (Query and Konarska, 2004; Konarska and Query, 2005). Our model suggests mechanistic similarities between the ribosome and spliceosome. In both cases, equilibrium between conformational states (e.g., the open/closed form of the ribosomal 30S subunit or the first/second step conformation of the spliceosome) can be modulated by several factors; these transitions are facilitated by the action of EF-Tu GTPase or Prp16 ATPase, respectively. Specific contacts between the tRNA substrate and the ribosome also contribute to the ribosomal transition through an induced fit mechanism, and similar induced fit interactions may contribute to transitions in the spliceosome. Our results argued that mutant alleles can affect each other’s phenotype by altering the equilibrium of the reaction, without physical contact between mutated sites (Query and Konarska, 2004). For example, a number of prp8 and nonprp8 alleles share the common property of improving second step efficiency for multiple intron mutants; concomitant first step inhibition by these alleles suggests that the first and second catalytic steps require different conformations of the spliceosome. Destabilization of interactions that favor the first step conformation inhibits the first step and promotes rearrangement into the second step conformation, thus suppressing second step defects. Thus far, our analysis focused on protein factors that most likely affect the stability of enzyme conformations; however, alteration of substrate:enzyme interactions would also be expected to affect the equilibrium between these conformations. Here, we examine interactions between the 50 SS and U6 snRNA and conclude that changes in the stability

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Figure 1. The 50 SS A3C Mutation Is Rate Limiting In Vivo for the Second Step of Splicing (A) Two chemical steps of splicing, depicting the ACT1-CUP1 reporter pre-mRNA with intron position +3 and +4 mutations used in (C), (D), and (E). As shown in (C), A3C lariat intermediates stall prior to the second step of splicing, but additional change of U4A relieves this second step block. (B) Schematic of RNA:RNA interactions that contribute to the first step of splicing. (C) Splicing of A3C is limiting in vivo but improved by an additional U4A mutation. Top, primer extension analysis of RNA from cells containing ACT1-CUP1 reporters, as indicated. Primer complementary to the 30 exon was used to reveal levels of pre-mRNA, mRNA, and lariat intermediate (indicated by icons to the left). Middle, quantitation of primer extension results presented above. For each reaction, the first step efficiency (light bars) was calculated as (M + LI) / (P + M + LI) and normalized to the first step efficiency of the wt reporter, set at 100. The second step efficiency (dark bars) was calculated as M / (M + LI) and normalized to the wt reporter second step efficiency, set at 100. Abbreviations: P, pre-mRNA; M, mRNA; and LI, lariat intermediate. Although these quantitations do not reflect absolute levels of RNA because of inherent losses due to discard pathways (see Figure S1), they nevertheless provide a useful illustration of relative differences between the tested reporters. Bottom, copper growth phenotype of the reporters used above. Comparable numbers of cells were spotted onto plates containing CuSO4, ranging from 0 to 2.0 mM. Relevant concentrations are shown, and the highest concentration on which each reporter supported growth is indicated below. (D and E) Splicing of the A3C mutant is improved by the U6-U57A allele, worsened by the U6-U57C allele, and improved by second step suppressor prp8-161 and 8-162 alleles. Primer extension analysis, quantitation, and growth on copper as in (C).

of these interactions alter the equilibrium between the first and second step conformations. Whereas pairing between 50 SS and the conserved U6-ACAGAG sequence facilitates the first step of splicing, hyperstabilization of this interaction inhibits the second step. Conversely, destabilization of this interaction improves the second step for introns that are otherwise suboptimal for the second step, suggesting that the 50 SS:U6-ACA GAG duplex must be disrupted in order for the branch structure to be removed from the first step catalytic center and to allow the reaction to proceed through the second step. These findings underscore the dynamics of RNA:RNA interactions during the catalytic phase of pre-mRNA splicing.

Results The 50 SS A3C Mutation Results in an Inefficient Transition between the First and Second Steps of Splicing The intron sequence surrounding the 50 SS is highly conserved among the w250 introns of S. cerevisiae, with /GUAUGU as the consensus. Mutation of A to C at position +3 (A3C) inhibits the second step of splicing, yet at least one gene (RPL30) naturally contains A3C, whose pre-mRNA is nevertheless spliced efficiently (Dabeva and Warner, 1987; Spingola et al., 1999). The RPL30 50 SS deviates from the consensus at both positions +3 and +4 (/GUCAGU); this covariation is conserved among

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various yeast species, including S. cerevisiae and K. lactis RPL30 genes (Eng and Warner, 1991). To investigate whether this represents a functional covariation, we tested the effects of position +3 and +4 mutations in the well-characterized S. cerevisiae actin intron. To monitor splicing of intron mutants in vivo, we used ACT1-CUP1 reporters (Lesser and Guthrie, 1993a; Figure 1A) and analyzed splicing both by resistance to copper in the growth medium and by primer extension of RNA isolated from cells. The 50 SS A3C mutation resulted in a defective second step of splicing in vivo, as evidenced by accumulation of lariat intermediates and a low level of spliced mRNA, consistent with poor growth on copper (Figure 1C, cf. lanes 2 and 1; also see Luukkonen and Se´raphin [1998] and Collins and Guthrie [2001]). Analysis of the A3C mutation in a strain lacking debranching enzyme (dbr12) showed strong accumulation of lariat intermediates, indicating that the first step was not defective compared to the wild-type (wt) pre-mRNA but rather that A3C lariat intermediates are discarded from the spliceosome (and degraded in the wt DBR1, but not in the dbr12, strain; Figure S1 available in the Supplemental Data with this article online). Changing 50 SS position +4 from U to A (U4A) had no detectable effect by itself (lane 3); however, in combination with the A3C mutation (A3C, U4A), it improved the second step relative to A3C, as indicated by reduced stalled lariat intermediates and increased mRNA levels, consistent with improved growth on copper (cf. lanes 4 and 2). These results suggested a relationship to the spliceosome ‘‘two-state model’’ (Query and Konarska, 2004) in which suppression of second step defects results from alteration of the equilibrium between the first and second step conformations. Spliceosomal mutations known to influence the relative efficiencies of the first versus the second step include alleles of U6 snRNA, PRP8, and other factors (Query and Konarska, 2004). We have tested U6-U57A and U57C alleles in the presence of a wide spectrum of intron mutations at the 50 SS, BS, and 30 SS and demonstrated that U6-U57A generally inhibits the first step but improves the second and U6-U57C acts in the opposite manner, improving the first step but inhibiting the second (data not shown; McPheeters, 1996; Query and Konarska, 2004). Consistent with the A3C defect affecting the first-to-second step transition, U6-U57A improved the second step for the A3C reporter (Figure 1D, cf. lanes 2 and 1), whereas U6-U57C acted in an opposite manner, inhibiting the second step (lane 3). The third allele of U6 at this position, U6-U57G, inhibited both steps of splicing, as observed for other reporters (lane 4; data not shown). The second step of the A3C mutant was also improved by the strongest prp8 and nonprp8 second step suppressor alleles described in Query and Konarska (2004) (Figure 1E and data not shown). Thus, the effects of known spliceosomal second step suppressors on A3C are similar to those of the U4A mutation. Taken together, these results are consistent with the transition between the first and second steps of splicing being kinetically limiting for the A3C substrate. Although the stability of 50 SS pairing with U1 snRNA contributes to 50 SS recognition (e.g., Roca et al. [2005] and references therein), the above-described effects cannot be attributed to U1:50 SS pairing. The U4A muta-

tion improved the second step efficiency for A3C (and other reporters defective for the second step; see below) and not the first step, as would be expected if it stabilized base pairing to U1 and thereby stimulated spliceosomal assembly, consistent with the finding that the U4A mutation does not alter binding to U1 snRNP (Libri et al., 2002). Mutations at 50 SS Position +4 Facilitate the Second Step for Position +3 Mutants The interplay between 50 SS A3C and U4A raised two questions: whether other position +3 mutants display similar second step defects, and whether other position +4 mutants may compensate for these defects. We tested all four bases at position +3 in combination with all four bases at position +4 (Figures 2A and 2B). When position +3 was wt, changes at position +4 did not have any detectable effect (lanes 1–4). When position +3 was C, the second step was limiting only for the wt U at position +4 (indicated by accumulation of lariat intermediates) and was improved by any non-U position +4 (lanes 5–8; indicated by reduced stalled lariat intermediates relative to mRNA). Although inhibition of the first step contributes to reduced levels of lariat intermediates, the ratio between these two signals indicates that in addition to inhibition of the first step there must also be improvement of the second step. A3G mutants were generally defective for the first step (or earlier; also see Figure S1), and thus, second step efficiencies were not measurable for some combinations (lanes 9– 12); however, with wt position +4 (U), A3G was not limiting for the second step (lane 12). A3U was also limiting for the first step, but not for the second (lanes 13–16 and Figure S1). In all cases, growth in the presence of copper correlated with mRNA levels detected by primer extension. Importantly, the only combination resulting in accumulation of lariat intermediates was A3C with the wt U at position +4 (Figure 2B, lane 8), and any non-U at position +4 suppressed the second step defect of A3C. Hyperstabilization of the 50 SS:U6 Duplex Inhibits the Second Step of Splicing A mechanistic basis for the defect of the 50 SS A3C mutation can be proposed from the model of first step catalytic core interactions shown in Figure 1B. Genetic evidence suggests a role for base pairing between 50 SS positions +4–6 (UGU) and U6 positions 47–49 (ACA) (Kandels-Lewis and Se´raphin, 1993; Lesser and Guthrie, 1993b), and crosslinking data suggest close juxtaposition of 50 SS positions U+2 and U6-A51 (Sontheimer and Steitz, 1993; Kim and Abelson, 1996; Ryan et al., 2004). Thus, U6-G50 is likely positioned in proximity to intron position A+3, consistent with described genetic interactions between these two bases (Luukkonen and Se´raphin, 1998; Collins and Guthrie, 2001). Mutation of A+3 to C would potentially create a Watson-Crick C:G base pair with U6-G50 and thereby hyperstabilize the 50 SS:U6 duplex (Figure 2C). The above-described U+4 mutations, which facilitate the second step for the A3C mutant, would disrupt a base pair with U6-A49 (Figure 2C), suggesting the possibility that 50 SS:U6 hyperstabilization inhibits the second step of splicing.

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and 8). This is consistent with the A3C second step defect resulting from a Watson-Crick interaction with U6G50 that stabilizes the 50 SS:U6 duplex but does not form in the context of U6-G50 mutants (Figure 3E). If the second step defect of A3C is due to position +3:G50 interaction, then A3G in combination with U6G50C might exhibit a second step defect similar to that of A3C in the presence of wt-U6. Indeed, in the presence of the U6-G50C allele, A3G became defective for the second step (Figure 3D, lane 7). Therefore, A3G:G50C displays a similar defect to that of A3C:wt-G50 (cf. lanes 2 and 7). Consistent with this observation, in a different reporter system, A3G mutations have been observed to inhibit the second step in combination with U6-G50C or G50U alleles (Luukkonen and Se´raphin, 1998). Thus, there is reciprocity in base pairing combinations at position +3 and U6-position 50 that inhibit the second step. Using the actin reporter, we have analyzed all combinations of U6-G50 alleles and mutations at positions +3 and +4; in this context, second step inhibition attributable to U6:50 SS hyperstabilization is observed only with the more stable G:C pairs and not with A:U or G:U pairs (data not shown). To further test the notion that the observed effects result from alterations in 50 SS:U6 duplex stability, we analyzed the effect of temperature on splicing of mutant pre-mRNAs. Lowering growth temperature (30ºC, 23ºC, and 17ºC) resulted in a progressive inhibition of the second step for A3C mutants (Figure 4, lanes 1–3), and this effect was maintained in strains harboring the prp8-161 second step suppressor allele (lanes 4–6). By contrast, lower temperatures did not inhibit the second step for wt or for A3C+U4A reporters (data not shown). These results are consistent with the A3C mutation increasing the stability of the 50 SS:U6 duplex and disruption of this hyperstabilized duplex becoming rate limiting for the transition between the first and second steps.

Figure 2. 50 SS Position +4 Mutations Improve the Second Step of the A3C Mutant (A) Schematic of ACT1-CUP1 reporter pre-mRNA, indicating intron position +3 and +4 mutations used in (B). (B) The only combination of all 16 position +3 and +4 mutants that results in a defective second step of splicing is A3C with wt position +4 U. Top, middle, and bottom panels are as in Figure 1C. An asterisk (*) represents reporters for which the second step efficiency is not measurable, due to low first step efficiency. (C) The A3C mutation creates a potential Watson-Crick C:G base pair between the 50 SS and U6 snRNA, extending and stabilizing the 50 SS:U6 duplex (outlined box). Mutation of position +4 would disrupt a U:A pair, thus reducing the stability of this duplex.

To investigate this, we tested the role of pairing between 50 SS A3C and all four alleles of U6-G50 (Figures 3A and 3B). Strikingly, any change of U6-G50 (to A, C, or U) resulted in strong improvement in the second step of A3C splicing, indicated by increased levels of mRNA, reduced levels of lariat intermediates, and improved growth on copper (Figure 3B, cf. lanes 2, 4, 6,

Destabilization of the 50 SS:U6 Duplex Facilitates the Second Step of Splicing Purines at 5 0 SS Position +4 Improve the Second Step for Multiple 3 0 SS and BS Second Step Mutants The A3C mutation worsened the second step defect of 30 SS mutations, such as gAG/ and UuG/ (data not shown, and see Collins and Guthrie [2001]). If hyperstabilization of the 50 SS:U6 duplex is responsible for A3C second step defects, then its destabilization might generally improve the second step. To investigate this possibility, we tested the effect of disrupting 50 SS:U6 pairing on a variety of mutants defective for the second step. First, we tested all nucleotides (nt) at 50 SS position +4 in combination with 30 SS gAG/ (Figures 5A and 5B). Whereas the gAG/ mutation with a wt 50 SS (U4) exhibited a significant second step defect (Figure 5B, cf. lanes 5 and 1), gAG/ in combination with U4A or U4G showed an improved second step (cf. lanes 2 and 4 with 5). This is consistent with disruption of the position +4:U6-A49 base pair improving the second step. U4C, however, did not improve the second step for the gAG/ mutant (lane 3), which may be explained by the stability of an A+:C wobble pair, similar to that of either a G:U wobble or A:U Watson-Crick pair (Strobel et al., 1994), stabilizing the 50 SS:U6 duplex. Importantly, the

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Figure 3. The Second Step Defect of the 50 SS A3C Mutation Is Relieved by Any U6-G50 Mutation, and A3G in Combination with G50C Recapitulates This Second Step Defect (A and C) Schematic of ACT1-CUP1 reporter pre-mRNA and U6 snRNA, indicating 50 SS position +3 and U6-G50 mutations used in (B) and (D), respectively. (B and D) Primer extension analysis of RNA recovered from cells, quantitation, and growth on copper as in Figure 1C. (E) The A3C mutation creates a potential Watson-Crick C:G base pair between 50 SS and U6 snRNA, extending and stabilizing the 50 SS:U6 duplex (outlined box). Mutation of U6-G50 would disrupt this C:G pair, thus reducing the stability of this duplex. A similar second step defect is recapitulated by A3G:U6-G50C.

presence of either purine at position +4 facilitated the second step for the gAG/ mutant. Second, we tested whether the second step effect of position +4 purines was general to a variety of mutants defective for the second step of splicing. Purines at position +4 improved the second step of all 30 SS mutants at position 22 (UcG/, UgG/, and UuG/; Figures 5A and 5B, lanes 9–17), as evidenced by increased mRNA levels and improved growth on copper. Third, U4A improved the second step for BS-G and BS-C mutants, which are also defective for the second step—in these cases, overall splicing and growth on copper were diminished due to a strong first step inhibition that accompanied the second step improvement (cf. lanes 19 and 18, and data not shown). The effect of the U4G mutation in combination with BS-G could not be determined because of even stronger inhibition of the first step (lane 20). We conclude that position +4 purines improve the second step of splicing for a wide spectrum of BS and 30 SS second step mutants.

Mutation at 5 0 SS Position +5 Improves the Second Step for Multiple 3 0 SS Mutants 50 SS position +5 (G5) provides the central pair of the 50 SS-UGU:U6-ACA interaction (Kandels-Lewis and Se´raphin, 1993; Lesser and Guthrie, 1993b). If position +3 and +4 mutations affect the second step by altering stability of this duplex, then a similar effect should also be observed with position +5 mutants. The G5A mutation destabilizes the 50 SS:U6 interaction, activating nearby cryptic 50 SS (25 and 210 in Figure 5C), which are used in addition to the original +1 50 SS (M.M.K., unpublished data; Lesser and Guthrie, 1993b). To analyze the effect of the G5A mutation on the second step, we combined G5A with either 30 SS gAG/ or UuG/ mutants (Figures 5C and 5D and data not shown) and quantified the level of second step occurring at the original +1 50 SS. Because the cryptic sites contain mutations that block the second step, only those lariat intermediates derived from the original +1 50 SS can proceed through the second step to yield functional mRNA. Primer extension analysis

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strate for first catalysis, it must be disrupted to allow the 50 end of the intron (now covalently attached to the BS) to be removed from the catalytic center at the stage of transition to the second catalytic step. Discussion The transition between the first and second steps of splicing requires a conformational change. We propose that the substrates for the reaction are repositioned during this transition, such that removal of the branch structure from the catalytic center is in competition with binding of the 30 SS substrate for the second step. We demonstrate that the previously described interaction between the 50 SS and U6 snRNA (positions 46–52 ACA GAG, the first step binding site for the 50 SS) is disrupted during transition to the second step. Destabilization of first step U6:50 SS pairing improves the efficiency of the second step, whereas hyperstabilization of this pairing inhibits the second step. These conclusions are drawn from studies of the in vivo spliceosome, using a genetic system that is highly sensitive to small changes in the stability of enzyme:substrate interactions. These findings strengthen the argument that the two catalytic conformations of the spliceosome exist in kinetic competition and indicate that substrate repositioning likely makes a major structural contribution to the transition between the first and second steps. Figure 4. Splicing of the A3C Mutant In Vivo Is Temperature Sensitive Lowering of growth temperature exacerbates the A3C second step defect. Primer extension analysis of RNA from cells grown for 3 hr at the indicated temperature and quantitations, as in Figure 1C.

using a 30 exon-specific primer detects all lariat intermediates formed by using the 50 SS at either the original position +1 or cryptic position 25 or 210 (Figure 5D, top gel). An intron-specific primer distinguishes these three lariats (Figure 5D, bottom gel). As expected, almost no signal was detected for wt reporter with the intron primer, consistent with no accumulation of lariat intermediates (lane 1). Lariat intermediates that accumulated for the G5A mutant represented only molecules branched at the 25 or 210 cryptic 50 SS (lane 2), whereas those accumulated in the 30 SS UuG/ reporter used the +1 50 SS (lane 3). The combination of G5A+UuG/ improved the second step for lariat intermediates branched at the functional +1 50 SS, as shown by a decrease in the +1 50 SS signal (Figure 5D, bottom, cf. lanes 3 and 4); this resulted in an overall increase in mRNA and improved growth on copper. Similar results were obtained for the 30 SS gAG/ mutant (data not shown). We also tested combinations of G5A with several 50 SS and BS mutants; however, these combinations strongly inhibited the first step of splicing, preventing analysis of the effect of G5A on the second step for these reporters (data not shown). Thus, mutations at position +4 (U4A, U4G) and position +5 (G5A) that destabilize the 50 SS:U6 interaction improve the second step for both BS and 30 SS position 22 and 23 mutants, indicating that destabilization of the 50 SS:U6 duplex generally improves the second step of splicing. By contrast, mutations that stabilize this duplex inhibit the second step. Therefore, whereas the 50 SS:U6 duplex is required for positioning of the sub-

The First Step Binding Site for 50 SS Current understanding of RNA:RNA interactions at the catalytic center of the spliceosome is substantially better at describing the first than the second step. The presence of base pairing between the 50 SS and the U6-ACAGAG site is strongly supported by numerous studies in yeast and mammalian cells and further strengthened by an analogous 50 SS:U6atac interaction in minor spliceosomes. Similarly, the importance of base pairing between U2 snRNA (U12 in minor spliceosomes) and the branch site region has been demonstrated for multiple systems (reviewed in Patel and Steitz [2003]). Extensive interactions between U2 and U6 snRNAs have been firmly established, although the detailed structure assumed by these snRNAs throughout splicing is not known. Our results demonstrate that whereas destabilization of 50 SS:U6-ACAGAG pairing inhibits the first step, it facilitates the second step for a variety of intron mutants defective for the second step. These findings confirm previous results that this 50 SS:U6 interaction is required for the first step (Kandels-Lewis and Se´raphin, 1993; Lesser and Guthrie, 1993b); however, they argue that this interaction must be disrupted for transition into the second step. This conclusion is based on the effects of mutations that stabilize 50 SS interactions with U6 snRNA (A3C with wt-U6 G50; A3G with U6-G50C) and inhibit the second step of splicing. By contrast, mutations that destabilize 50 SS interactions with U6 snRNA (U4A or U4G, G5A) improve the second step (while inhibiting the first) for a variety of reporters. Because U4-to-purine mutations suppress second step defects caused by numerous BS and 30 SS mutations, this effect is clearly not nucleotide or position specific—rather, it results from altering

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Figure 5. Disruption of Potential Base Pairing between U6 and 50 SS Position +4 or +5 Improves the Second Step for Multiple 30 SS and BS Mutants (A and C) Schematic of ACT1-CUP1 reporter pre-mRNAs and U6 snRNA, indicating intron mutations used in (B) and (D), respectively. (B and D) Primer extension analysis of RNA from cells, quantitation, and growth on copper as in Figure 1C. Lower gel of (D) shows primer extension analysis with an intron-specific primer, thus indicating the site of 50 SS cleavage. Quantitation in (D) represents the second step efficiency only for products having used the +1 (wt) 50 SS, normalized to wt reporter (lane 1). (E) Disruption of 50 SS:U6-ACAGAG interactions improves the second step for BS and 30 SS mutants.

a general feature of the first step conformation. Interestingly, the mammalian 50 SS consensus contains A or G at position +4 and 15% non-G at position +5 (Burge et al., 1999), suggesting that removal of the branch structure from U6-ACAGAG is less limiting for these introns, perhaps compensating for suboptimal second step interactions. Formally, alterations in the 50 SS:U6 interaction could affect the outcome of splicing not by altering the stability of the duplex per se but rather by modifying interactions

of this helix with other components of the catalytic center. In this view, mutation of the duplex would destabilize the first step catalytic center, facilitating transition into the second step conformation. Because genetic analysis is limited to U6 positions that can be mutated while maintaining cell viability, we could not alter U6 positions 48–49 and, thus, cannot exclude such a contribution to second step improvement for these positions of the duplex. However, the reciprocity of the effects of A3G:U6G50C and of A3C:wt-U6-G50 argues that stability of this

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Figure 6. Model of Structural Rearrangements within the Spliceosome at the Time of Transition between the First and Second Steps of Splicing (A) The first and second catalytic steps require different conformations of the spliceosome, the equilibrium between these conformations being modulated by the stability of the 50 SS:U6-ACAGAG duplex, in addition to interactions of Isy1 (Villa and Guthrie, 2005), U6 snRNA, Prp8, Prp16, and other factors (Query and Konarska, 2004). By analogy to the ribosomal decoding process, the initial signal for this conformational change may be presented by the branch structure formed in the first catalytic step. This induced-fit conformational change is facilitated by a weakened 50 SS:U6-ACAGAG duplex. Transition between first and second step conformations requires disruption of 50 SS:U6-ACAGAG pairing; thus, strong 50 SS:U6 pairing disfavors exit from the first step conformation, improving first step catalysis and inhibiting the second step; conversely, weak 50 SS:U6 pairing promotes progression into the second step conformation, improving splicing of mutant intermediates defective in this step. (B–D) Summary of 50 SS-U6 snRNA crosslinks prior to the first catalytic step (B) (Johnson and Abelson, 2001; Chan et al., 2003; Chan and Cheng, 2005), in the first step conformation (C), (Sontheimer and Steitz, 1993; Kim and Abelson, 1996), and lariat product-U6 crosslinks after the first step (D) (Sawa and Abelson, 1992). Crosslinks in S. cerevisiae are shown by gray bolts, those in HeLa extracts by black bolts.

duplex itself significantly affects the first-to-second step transition. The concomitant inhibition of the first step and improvement of the second (and vice versa) by mutations that alter the stability the 50 SS:U6-ACAGAG duplex further supports the two-state model, according to which structural conformations of the spliceosome active in the two catalytic steps exist in competition (Figure 6A; Query and Konarska, 2004). Destabilization of interactions that favor the first step conformation inhibits the first step and promotes rearrangement into the second step conformation, thus suppressing second step defects. Because the stability of substrate binding at the first step catalytic center alters efficiencies of the first and second steps, and because the substrates must be repositioned between the two steps of splicing, an attractive possibility is that partitioning between first and second step conformations primarily results from competition between first step products (i.e., the branch structure of the lariat intermediate) and second step substrates (30 SS region of the lariat intermediate) binding at the catalytic center. (The free 50 exon, both a product of the first step and a substrate for the second, need not reposition.) Assessment of a detailed model clearly requires consideration of the first and second step binding sites for both the branch structure and the 30 SS. Further analysis is needed to unequivocally identify these substrate:enzyme interactions, in particular those required for the second step. Repositioning of the Lariat Intermediate Enzymes that catalyze a series of reactions in which a product of one step is used as a substrate for the

next typically require repositioning of at least one of the reaction products prior to proceeding to the next reaction. In the case of the spliceosome, a single active site has been proposed; as a consequence, placement of the 30 SS into the second step catalytic center would require prior displacement of the branch nucleophile (Steitz and Steitz, 1993). Stereochemical analysis of splicing is consistent with some rearrangement being required between the two steps (Moore and Sharp, 1993). In support of this view, our data argue that placement of the 30 SS at the second step catalytic center requires removal of the branch structure formed during the first step. Genetic interactions between 50 SS position +1 and 30 SS position 21 (Parker and Siliciano, 1993) and 50 SS position +3 and 30 SS position 23 (Collins and Guthrie, 2001) suggested a different model of RNA:RNA interactions at the catalytic center, according to which the 50 and 30 ends of the intron are simultaneously bound and aligned in antiparallel fashion, with intron position +3 juxtaposed with position 23 (Collins and Guthrie, 2000, 2001). This model was based on several independent lines of evidence suggestive of interactions between the 50 SS, U6-ACAGAG, and 30 SS. Among these were exacerbation of defects caused by intron mutations at positions +3 and 23 (i.e., mutations of A3 exacerbate defects of rAG/ 30 SS mutants; r = purine) and suppression of A3C+aAG/ double mutation by U6-G50C and U6-G50U alleles (Collins and Guthrie, 2001). Although our data confirm these genetic interactions, they do not overall support such a model. First, genetic interaction between the 50 and 30 SS is not specific to positions +3 and 23. Rather, mutations

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of position +3 generally exacerbate second step defects, not only those due to mutation at position 23 but also at other intron positions (such as 30 SS position 22 and BS; data not shown). More importantly, mutations of 50 SS position +4 or +5 also exhibit similar nonposition-specific genetic interactions, in that they generally rescue second step defects caused by mutations at 30 SS positions 23, 22, or at the branch site (Figure 5). Second, suppression of A3C by U6-G50 alleles is not allele specific. The second step defect of the A3C mutation results from hyperstabilization of the 50 SS:U6-ACA GAG pairing and is suppressed by all mutations of U6G50, which destabilize this duplex. We propose that the hyperstabilized 50 SS:U6 duplex inhibits the second step by slowing exit of the branch structure (first step product) from the first step catalytic center. A mutant 30 SS (second step substrate) competes inefficiently with such a hyperstabilized first step product, exacerbating the A3C defect. Thus, U6-G50 alleles suppress A3C+30 SS double mutants not because of a simultaneous interaction between the three nucleotides but rather because they improve exit of first step products from the catalytic center, facilitating placement of 30 SS mutants for the second step. The same mechanism explains suppression of 30 SS and BS defects by U4A, U4G, and G5A mutants. These considerations argue against direct simultaneous interactions between the ends of the intron (in agreement with Luukkonen and Se´raphin [1997] and Dietrich et al. [2005]). Instead, both the previously described genetic interactions and our new results can be explained by alterations of equilibrium between two catalytic conformations that differ in substrate positioning for the two reactions. Whereas the 50 end of the intron is present in the catalytic center during the first step, it must be removed during the transition into the second step, when the 30 SS is placed at the catalytic center. Thus, their presence at the catalytic center is mutually exclusive. This view is supported by the demonstrated accessibility of the 30 SS to nuclease digestion prior to the second step (Schwer and Guthrie, 1992), by the fact that first step catalysis does not require the presence of the 30 SS (Rymond and Rosbash [1985], Vijayraghavan et al. [1986], Anderson and Moore [1997], and references therein), and by the ability of U5 loop nt to crosslink to the 30 exon only after the first step (Sontheimer and Steitz, 1993). It should be noted, however, that the first and second step substrates need not be distant in the three-dimensional space of the spliceosome. Strikingly similar conclusions have been reached for group II self-splicing introns, whose splicing proceeds by the same pathway of two-transesterification steps as does pre-mRNA splicing. Modification interference studies of group II introns suggested that a conformational change between the first and second steps results in displacement of the branch structure from the active site, which was proposed to be required for positioning of the 30 SS for the second step (Chanfreau and Jacquier, 1996). Although our results directly address only the fate of the 50 SS:U6-ACAGAG interaction, at the time of transition, the 50 SS is covalently attached to the branch site and one must therefore question the fate of the branch

site:U2 interaction. One possibility is that the 50 SS becomes unpaired from U6 and the branch site remains at the catalytic center. Alternatively, the entire branch structure is removed from the first step catalytic core. In this case, U2 snRNA may either become unpaired from the branch structure and retain its position in the catalytic core or be repositioned together with it (resembling the group II intron situation in which the entire domain VI that contains the branch site is thought to be rearranged in the transition to the second step [Chanfreau and Jacquier, 1996]). In either case, the repositioned branch structure is likely to interact with a nearby second step site (akin to the ribosomal ‘‘exit’’ site). Positioning of the Lariat Intermediate during the Second Step A common, direct method to monitor substrate positioning within the catalytic center relies on RNA:RNA crosslinking. A number of studies, using both HeLa and yeast extracts, have used site-specific placement of photoreactive moieties or mapping of UV-induced crosslinks to identify close contacts between substrate and snRNAs. In support of the notion that the branch structure is repositioned after the first step, in HeLa extracts, U5 snRNA loop nt have been crosslinked to 50 SS position +2 in pre-mRNA and chased into lariat intermediate (McConnell and Steitz, 2001) and U5:50 SS exon position 21 crosslinks could be chased through both the first and second steps (Sontheimer and Steitz, 1993); however, crosslinks from U5 to lariat products were not observed, and the U5:50 SS+2 crosslink was not chased into lariat product (McConnell and Steitz, 2001). Similar analysis has identified two distinct sets of UV crosslinks formed between the 50 SS and U6 at different stages of the splicing reaction. The first of these corresponds to U6 position U47-A51 juxtaposed against the 50 SS during the first step (Sontheimer and Steitz, 1993; Kim and Abelson, 1996), consistent with the 50 SS:U6 pairing demonstrated by genetic interactions (Kandels-Lewis and Se´raphin, 1993; Lesser and Guthrie, 1993b; Figure 6C). The second set corresponds to U6 position G39-A44 crosslinked to the 50 SS both in early complexes (Johnson and Abelson, 2001; Chan et al., 2003; Figure 6B) and within the branch structure of the lariat intermediate (Sawa and Abelson, 1992; Figure 6D). This latter set of crosslinks likely reflects the new location of the branch structure after repositioning from the first step site. These physical constraints relative to U6 snRNA have been provided both in S. cerevisiae and HeLa systems. In yeast, U6-A51 crosslinked to pre-mRNA at the 50 SS from positions 22 to +3, and U6-U46 crosslinked to 50 SS from positions +4 to +7 (Kim and Abelson, 1996). Importantly, the U6-A51 crosslinks formed to pre-mRNA could be chased through the first step, confirming that the 50 SS is positioned near U6-A51 during the first catalytic step. By contrast, crosslinking of U6 position G39-A44 (U6-GAAACA) was observed to the 50 SS within lariat intermediate (Sawa and Abelson, 1992). In addition, in HeLa extracts, position U+2 in lariat intermediates crosslinked exclusively to U6-A51 (hU6-A45), whereas the same position in lariat products crosslinked most prominently to two U6 nt, U6-A51 and U6-U46 (hU6-A45 and hU6-U40) (Sontheimer and Steitz, 1993). Intriguingly, the same two sites on U6 snRNA were shown to interact

Molecular Cell 552

with the 50 end of the intron in pre-mRNA, i.e., prior to the first catalytic step (Johnson and Abelson, 2001; Chan et al., 2003; Chan and Cheng, 2005). Crosslinking to the U6-GAAACA site occurred in the absence of the NTC (Prp19-associated complex), whereas in the presence of the NTC, the U6-ACAGAG site was crosslinked (Chan et al., 2003). Formation of crosslinks does not necessarily require direct base pairing between two RNAs but only their physical proximity. Thus, U6-GAAACA may form part of a binding pocket; alternatively it may be positioned in proximity of the binding site without contributing directly to binding of the 50 SS (both prior to and after the first step). Although our results do not directly address the 50 SS interaction with the U6-GAAACA site, the first step inhibition observed for some position +3 and +4 mutants may in part reflect defective 50 SS interactions prior to the first catalytic step (e.g., Figure 2B, lanes 6 and 7, 9–13, and 15). These results suggest two distinct locations of the 50 SS at different stages in the reaction: one in proximity of U6-ACAGAG (demonstrated both genetically and by crosslinking experiments) and the other in proximity of U6-GAAACA (detected by crosslinking). Crosslinking data suggest that initially the 50 SS is positioned close to U6-GAAACA (Johnson and Abelson, 2001; Chan et al., 2003); the NTC is then required for the repositioning of the 50 SS into the first step catalytic center, where it interacts with U6-ACAGAG (Sontheimer and Steitz, 1993; Kim and Abelson, 1996; Chan et al., 2003). Subsequently, after first step catalysis, the 50 end of the intron (in the branch structure) is moved again; our genetic data argue that it is moved out of the catalytic center, and crosslinking data suggest that it moves into proximity of U6-GAAACA (Sawa and Abelson, 1992). Although we do not yet understand the molecular basis of this repositioning of the branch structure, one distance restraint is the shortest functional BS to 30 SS distance, which can be 10 nt for some S. cerevisiae introns (e.g., MATa1) or as short as 7 nt in other species, e.g., S. pombe, G. lamblia, and T. vaginalis (Wood et al. [2002], ova´ et al. [2005], and references therein). HowVanˇa´c ever, the movement of the branch structure could be much smaller, as long as it provides enough space for positioning of the 30 SS in the catalytic center for the second step. This raises a related question: where is the 30 SS located during the first step? The same distance considerations that restrain movement of the branch structure also apply to movement of the 30 SS from its first step site to the second step catalytic center. The above restraints represent maximum distances possible, but either one or both sites (first step location of 30 SS and second step location of branch structure) may be directly adjacent to the catalytic center. Whereas these considerations highlight dynamics of the catalytic phase of splicing, they also raise a number of additional questions. Clearly, future experiments will need to identify components (RNA and/or proteins) of the binding sites for both the branch structure and 30 SS. Some of the 30 SS binding components may already have been identified (e.g., Prp8, Prp16, Prp22, and U2 snRNA [Newman et al., 1995; Umen and Guthrie, 1995; McPheeters and Muhlenkamp, 2003]), but better understanding of the temporal contributions of these interactions is needed. In addition, clarification of the mo-

lecular nature of repositioning will be needed, including identification of spliceosomal interactions, whose disruption/formation may be required at the time of transition between the first and second steps, and further characterization of the consequences of Prp16 action, which facilitates this transition. Experimental Procedures Strains and Reporter Plasmids S. cerevisiae strains were derived from yCQ05 (MATa, ade2 cup1D::ura3 his3 leu2 lys2 prp8D::LYS2 trp1 U6D::KAN; pCC130 [U6 URA3 CEN ARS]; and pMK8-1 [PRP8 HIS3 CEN ARS]) containing plasmid-borne alleles of prp8 and U6 (snr6) as indicated in the figures. ACT1-CUP1 reporter plasmids (Lesser and Guthrie, 1993a) with 50 SS, BS, or 30 SS mutations were as described (Query and Konarska, 2004) or prepared by overlapping PCR and in vivo gap repair cloning. Copper Assays Cultures were grown to midlog phase in 2Leu medium, diluted to 0.2 OD, and equal volumes were dropped onto 2Leu plates containing CuSO4 ranging from 0 to 2.0 mM (Lesser and Guthrie, 1993a). Plates were scored and photographed after 3 days at 30ºC or after 4 days at 28ºC for experiments comparing U6-G50 mutants. RNA Analysis Primer extensions were carried out as described (Query and Konarska, 2004), using primer YAC6 50 -GGCACTCATGACCTTC-30 , complementary to exon 2 of ACT1, or YAC5 50 -CGAGCAATTGGGAC CGTGC-30 , complementary to the intron. Extension products were separated in 7% polyacrylamide/8 M urea gels and visualized by autoradiography. Supplemental Data Supplemental Data include one figure and can be found with this article online at http://www.molecule.org/cgi/content/full/21/4/543/ DC1/. Acknowledgments We are grateful to Tim Nilsen, Duncan Smith, and Jon Warner for helpful discussions and critically reading the manuscript and to David McPheeters for providing U6-G50 alleles. This work was supported by National Institutes of Health (NIH) grant GM49044 to M.M.K., by grant BMC2002-00157 and program ‘‘Ramo´n y Cajal’’ of the Spanish MEC to J.V., by NIH grant GM57829 to C.C.Q., and by a Cancer Center Support (core) grant from the National Cancer Institute to AECOM. Received: August 11, 2005 Revised: December 2, 2005 Accepted: January 5, 2006 Published: February 16, 2006 References Anderson, K., and Moore, M.J. (1997). Bimolecular exon ligation by the human spliceosome. Science 276, 1712–1716. Burge, C.B., Tuschl, T.H., and Sharp, P.A. (1999). Splicing of precursors to mRNAs by the spliceosomes. In The RNA World, Second Edition, R.F. Gesteland, T.R. Cech, and J.F. Atkins, eds. (New York: Cold Spring Harbor Laboratory Press), pp. 525–560. Burgess, S., Couto, J.R., and Guthrie, C. (1990). A putative ATP binding protein influences the fidelity of branchpoint recognition in yeast splicing. Cell 60, 705–717. Chan, S.P., and Cheng, S.C. (2005). The Prp19-associated complex is required for specifying interactions of U5 and U6 with pre-mRNA during spliceosome activation. J. Biol. Chem. 280, 31190–31199. Chan, S.P., Kao, D.I., Tsai, W.Y., and Cheng, S.C. (2003). The Prp19p-associated complex in spliceosome activation. Science 302, 279–282.

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