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. 2012 Feb 9;109(8):2872-2877. doi: 10.1073/pnas.1113620109

Interdomain dynamics and coactivation of the mRNA decapping enzyme Dcp2 are mediated by a gatekeeper tryptophan

Stephen N Floor a,b,1, Mark S Borja b,c, John D Gross b,2
PMCID: PMC3286934  PMID: 22323607

Abstract

Conformational dynamics in bilobed enzymes can be used to regulate their activity. One such enzyme, the eukaryotic decapping enzyme Dcp2, controls the half-life of mRNA by cleaving the 5′ cap structure, which exposes a monophosphate that is efficiently degraded by exonucleases. Decapping by Dcp2 is thought to be controlled by an open-to-closed transition involving formation of a composite active site with two domains sandwiching substrate, but many details of this process are not understood. Here, using NMR spectroscopy and enzyme kinetics, we show that Trp43 of Schizosaccharomyces pombe Dcp2 is a conserved gatekeeper of this open-to-closed transition. We find that Dcp2 samples multiple conformations in solution on the millisecond-microsecond timescale. Mutation of the gatekeeper tryptophan abolishes the dynamic behavior of Dcp2 and attenuates coactivation by a yeast enhancer of decapping (Edc1). Our results determine the dynamics of the open-to-closed transition in Dcp2, suggest a structural pathway for coactivation, predict that Dcp1 directly contacts the catalytic domain of Dcp2, and show that coactivation of decapping by Dcp2 is linked to formation of the composite active site.

Keywords: enzyme dynamics, methyl groups, mRNA decay, protein NMR

Conformational dynamics in enzymes often comprise the rate-limiting step in the catalytic cycle and thus are prime targets for regulatory cofactors (1–5). Bilobed proteins frequently use an open-to-closed transition to coordinate catalysis on their substrates following cellular cues such as posttranslational modifications or macromolecular interactions (6–8). A recent model proposes that the eukaryotic mRNA decapping enzyme Dcp2 is regulated by such a transition, where a composite active site is formed using conserved surfaces on each of the two N-terminal domains (9). According to this model, stimulating or inhibiting this conformational transition could regulate decapping. However, the structural details of this composite active site and the timescale of interconversion between closed and open states of Dcp2 are currently unknown. Moreover, whether coactivators use the composite active site to effect decapping is unclear.

Degradation of eukaryotic mRNA is critical to many biological processes including development (10), stress response (11), clearance of the products of pervasive transcription (12), and quality control of gene expression (13). For example, it has been suggested that microRNAs (miRNAs) act primarily by destabilizing messages and it is known that Dcp2 is a vital component of miRNA-induced mRNA decay (14, 15). Further, an entire class of unstable transcripts was recently discovered that is sensitive to the exonuclease Xrn1, whose members are therefore likely products of decapping (12, 16). Each of the variety of pathways that utilize decapping relies on coactivator proteins that are believed to recruit messages to the decapping machinery and activate it.

A model of decapping coactivation is emerging following recent work on the Saccharomyces cerevisiae coactivator Edc1 (17). Edc1 is a yeast-specific protein that is required for carbon source changes and is strongly upregulated during such transitions (18–20). It binds directly to Dcp1, which in turn forms a stable complex with the regulatory domain of Dcp2 (17, 21). Dcp1 has an enabled/VASP homology-1 (EVH1) fold and uses a hydrophobic patch to recognize a proline-rich stretch in the C terminus of Edc1, which is also found in other putative coactivators (17, 22). Binding of Edc1 raises the catalytic efficiency of the Dcp1∶Dcp2 complex by up to 3,000 times by enhancing both the KM for mRNA and rate of the catalytic step kmax (17). Interestingly, it appears that Edc1 is modular with the N-terminal region responsible for the KM enhancement and the C-terminal region primarily affecting kmax (17). The mechanism of this enhancement is unknown though it was proposed that Edc1 may stimulate closure and thereby activity of Dcp2.

Many proteins are dynamic on the millisecond-microsecond (ms-μs) timescale and these motions can be intimately tied to activity (1). Dcp2 is known to undergo an open-to-closed transition that leads to formation of the composite active site and it was suggested that both the apo and ligand-bound forms of the enzyme sample multiple conformations (9). Open-to-closed transitions in bilobed proteins like Dcp2 can occur on the ms-μs timescale, which can be monitored with site-specific resolution by NMR spectroscopy (23, 24). Motions on the ms-μs timescale lead to dephasing of transverse magnetization which manifests in a rate constant Rex that, together with R2 for molecular tumbling, determines the resonance linewidth. Carr–Purcell–Meiboom–Gill (CPMG) NMR spectroscopy allows contributions to the resonance linewidth from molecular tumbling to be separated from ms-μs dynamics (25). When coupled with 13C-methyl Ile, Leu, Val, Met, and Ala (ILVMA) labeling CPMG can be applied to large molecular weight complexes and highly dynamic proteins (26). Refocusing pulses applied at various frequencies result in dispersion curves that are fit to either the general two-site equation (SI Methods) or the fast-exchange limit:

graphic file with name pnas.1113620109eq1.jpg [1]

where R2,eff is the effective transverse relaxation rate, R2,0 is the transverse relaxation rate of the major state, pA and pB are the population of the major and minor states, Δω is the chemical shift difference between states, kex is the rate of exchange between states, and νCPMG is the refocusing frequency (27–29). In the general case kinetics (kex), thermodynamics (pA, pB), and structure (Δω) are accessible by CPMG spectroscopy, but in the fast-exchange limit one can only extract the composite term pApBΔω2 and the exchange rate kex (30). CPMG is sensitive to dynamics involving skewed populations with exchange rates between approximately 100 and 5,000 s-1 (28).

Despite its importance in regulation of mRNA decapping, structural, kinetic, and thermodynamic details regarding the open-to-closed transition of Dcp2 are poorly understood. Additionally, it has been suggested, but not tested, that coactivators may stimulate decapping by promoting closure of Dcp2. Here we show that Dcp2 exists in a conformational equilibrium in solution between open and closed states in the absence of ligand that is fast on the NMR timescale. Exchange between states depends on a conserved, solvent-exposed tryptophan previously shown to bind m7G of cap, and implicated as a critical component of the composite active site in Dcp2 (9). Enzyme kinetics and NMR experiments reveal that this tryptophan acts as a gatekeeper that promotes formation of the composite active site and allows Dcp1 and bound coactivators to enhance the catalytic step.

Results

Dcp2 Samples Multiple Conformations in Solution.

To investigate the dynamics of the proposed composite active site in solution, we turned to NMR spectroscopy of the N-terminal 243 residues of Schizosaccharomyces pombe Dcp2 (spDcp2), which contains the regulatory and catalytic Nudix hydrolase domains and is conserved from yeast to humans (31). The first 15N heteronuclear single quantum correlation (HSQC) spectrum acquired of spDcp2 was of marginal quality with many missing resonances and a large variation in peak intensity (Fig. 1), typical of proteins undergoing ms-μs dynamics in solution. Because there is a proposed composite active site between the two domains of Dcp2 (9), and Dcp2 was crystallized with ATP in two conformations (21), we reasoned there might be multiple states coexisting in solution giving rise to the observed resonance broadening. To test this possibility we mutated select sites on both domains of spDcp2 and found that mutation of Trp43 of the regulatory domain induced profound changes in the nitrogen HSQC (Fig. 1). This mutation decouples the two domains because the Trp43Ala spectrum is more similar to the spectra of the isolated domains than the wild-type spectrum (Fig. S1). Tryptophan 43 is required for decapping in vitro and in vivo, closure by small-angle X-ray scattering (SAXS), and cap binding by the regulatory domain (9, 32). These data suggest that it also mediates an interdomain interaction in Dcp2 in the absence of substrate.

Fig. 1.

Fig. 1.

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Mutation of Trp43 in the regulatory domain of spDcp2 alleviates resonance broadening in the 15N HSQC. Shown are the 15N HSQC spectra of wild-type S. pombe Dcp2 residues 1–243 (red) and spDcp2 Trp43Ala (black). Selected residues with significant changes upon mutation are indicated; note that residues following 95 are located on the catalytic domain.

Quantitative Measurement of Methyl Side-Chain Dynamics in spDcp2.

We turned to CPMG spectroscopy to quantify the kinetic and thermodynamic properties of the dynamic behavior observed in the 15N HSQC. Because the cross-peaks for dynamic residues are broadened in the 15N HSQC (Fig. 1) we used 13C-methyl ILVMA side-chain labeling of spDcp2 for quantification. We found that approximately one third of methyl groups in spDcp2 undergo significant ms-μs dynamics that are attenuated by CPMG pulses (Fig. 2, Figs. S2–S4). Exchange rates ranged from approximately 500 s-1 to 3,000 s-1 and all but three are fit well by the fast-exchange limit formula (SI Methods, Fig. S4, Table S1, and Table S2).

Fig. 2.

Fig. 2.

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Quantitative analysis of methyl ILVMA side-chain dynamics in spDcp2. (A) Four representative CPMG dispersion curves are shown with experimental data in circles and fits in lines. Data acquired at 800 and 900 MHz are in black and red, respectively. Error bars are the pooled standard deviation (SI Methods). (B) Methyl groups in spDcp2 1–243 are displayed as spheres on PDB ID code 2QKM. Methyls that were fit are orange, with intensities that were too weak to quantify are red, insignificantly broadened by ms-μs dynamics (Rex less than 5 s-1) are gray and unassigned are black. The regulatory domain is shown in purple, catalytic domain in green, and the catalytic Nudix helix in red with Trp43 in cyan.

To identify the dynamic residues, we assigned the methyl groups of ILVMA residues of spDcp2. Dynamic residues are distributed throughout both domains of the protein (Fig. 2B, orange spheres). Crucially, the exchange broadening (Rex) of nearly all of the methyl groups in the isolated domains of spDcp2 is insignificant (Fig. S5), strongly suggesting that the dynamics measured in spDcp2 are not due to fluctuations within the individual domains, but rather a transient interdomain interaction. Moreover, as the dynamic behavior is largely unchanged by twofold sample dilution, the observed resonance broadening is not due to intermolecular interactions (Fig. S3). Previous SAXS analysis showed that spDcp2 undergoes an open-to-closed transition promoted by nucleotide, but closure was not observed without bound Dcp1 (21). The results presented here show the enzyme samples multiple states in the absence of cofactors.

A Tryptophan Dynamically Links the Regulatory and Catalytic Domains.

If the exchange broadening is due to a concerted process such as an open-to-closed transition, then mutations that disrupt closure should alter the exchange broadening in a uniform manner. To test this possibility we mutated Trp43 of spDcp2 to alanine and measured ms-μs dynamics by 13C-methyl CPMG. This mutant was chosen because it blocks closure by SAXS and has a profound effect on the nitrogen HSQC of spDcp2 (Fig. 1) (9). Remarkably, mutation of Trp43 to alanine strongly damps relaxation in 18 out of 23 methyl groups that are ms-μs dynamic in the wild-type protein (Fig. 3). Of those not affected by mutation of Trp43, two are dynamic in the isolated catalytic domain (Leu113 and Val114) (Fig. S5) and one is in the flexible interdomain linker (Ile102). Mutation of Trp43 quenches relaxation due to dynamics not only on the regulatory domain but also the catalytic domain, consistent with its global effect on the nitrogen HSQC (Fig. 1, Fig. 3). These results implicate Trp43 in an interdomain interface of an excited state of spDcp2 that is transiently sampled in solution. Notably this interdomain interface is different than the one found in the closed crystal structure because mutation of the interaction partner Arg167 does not affect kinetics, closure by SAXS or the nitrogen HSQC (Fig. S6) (9).

Fig. 3.

Fig. 3.

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Mutation of Trp43 in Dcp2 strongly attenuates the majority of ms-μs dynamic methyl groups. CPMG data for four representative residues at 800 MHz for wild-type or Trp43Ala Dcp2 are in black and pink, respectively. All but the following residues had Rex values less than experimental error after Trp43Ala mutation: Val112, Leu113, Val114, Ile179, and Ile102. Error bars are the pooled SD.

The Regulatory and Catalytic Domains Experience Collective Dynamics.

An additional prediction of a concerted dynamic process is that residues on both domains should be able to be fit collectively as a group to one global exchange rate. Alternatively, residues may undergo motions on the ms-μs timescale due to local, independent conformational changes such as loop dynamics. For example, residues Leu113, Val114, Ile193, Leu201, Leu204, and Leu226 are dynamic in the isolated catalytic domain (Fig. S5) as previously observed for the S. cerevisiae ortholog (33). However, many more residues are dynamic in the two-domain spDcp2 than in the isolated domains (23 versus 8, Fig. S4, Fig. S5). This observation, along with the decoupling effect of the Trp43 mutation, suggests the dynamics involve an interdomain interaction.

To rigorously determine if there are collective motions within spDcp2, we isolated a group of fourteen residues to collectively fit CPMG exchange curves to (Fig. 4 and Fig. S7; for group membership rationale see SI Methods). A collective exchange rate fits the group well, with fits extremely close to the individual fits (Fig. 4A dashed versus solid lines, note that Ile130 has the fourth largest difference between individual and group fits). Most interestingly, six of the group members are on the regulatory domain and three are on the catalytic domain, with no such constraint imposed (Fig. 4B, the remaining five are unassigned). We therefore conclude that apo spDcp2 experiences collective interdomain dynamics that depend on the presence of Trp43 with a rate of kex = 2,299 ± 74 s-1, likely the sum of opening and closing rates. Unfortunately, because of protein solubility constraints, we cannot perform similar experiments on substrate-bound protein at present.

Fig. 4.

Fig. 4.

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The regulatory and catalytic domains of spDcp2 are involved in a collective exchange process. (A) Representative CPMG group fits are shown for residues Leu90 and Ile130. Four different fits are shown in these plots: 800 and 900 MHz individual fits (black and red, solid) and 800 and 900 MHz group fits (gray and orange, dashed). The ratio of group to individual F-statistics for Leu90 is 1.02 and for Ile130 is 1.14. (B) The nine assigned members of the fourteen-member group are displayed on PDB ID code 2QKM as orange spheres with other colors as in Fig. 2.

Tryptophan 43 Links Dcp1 and the Catalytic Domain of Dcp2.

Both Dcp1 and Trp43 are thought to promote closure of Dcp2, so we reasoned there might be a coupling between the two (9, 21). To test this possibility we formed complexes between 13C-methyl ILVMA labeled spDcp2 and unlabeled S. pombe Dcp1 and monitored chemical shift changes. Because this heterodimer is dynamic and approaching 50 kDa, which obliterated the nitrogen HSQC, we turned to the 13C-heteronuclear multiple quantum correlation (HMQC), also known as methyl-transverse relaxation optimized spectroscopy (methyl-TROSY) (34). Stable complexes of Dcp1 and spDcp2 for both wild-type and Trp43Ala constructs yield high quality ILVMA spectra with widespread chemical shift changes (Fig. S8). Because we were unable to obtain full resonance assignments of the Dcp1∶Dcp2 complex due to exceedingly high Rex, we used the minimum chemical shift metric to measure chemical shift changes, which places a conservative lower bound on the perturbation of assigned Dcp2 resonances (35). As expected, residues proximal to the Dcp1∶Dcp2 regulatory domain interface observed by crystallography are shifted considerably upon addition of Dcp1, which depended little on mutation of Trp43 (Fig. 5, Fig. S8).

Fig. 5.

Fig. 5.

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Dcp1 causes chemical shift changes on Dcp2 outside its binding site that depend on Trp43. (A) The minimum chemical shift change for select residues is plotted for Dcp1∶Dcp2 (orange), Dcp2 Trp43Ala (black), and Dcp1∶Dcp2 Trp43Ala (gray). Each chemical shift change is calculated with respect to the wild-type Dcp2 HMQC. The horizontal line indicates a chemical shift change of 0.05 ppm (approximately one linewidth). (B) Shifted residues from A are shown in orange sticks on a model of the S. pombe Dcp1•Dcp2 open structure using PDB ID codes 2QKM and 2A6T (SI Methods). Colors are as in Fig. 2 with Dcp1 in gold. (C) An illustration of the Dcp1•Dcp2 complex with Edc1 and cap (m7G) binding sites in gray, regions shifted by Dcp1 addition in orange, and other colors as in B.

A second subset of residues outside the Dcp1 binding site is shifted by the addition of Dcp1 only when Trp43 is not mutated (Fig. 5, Fig. S8). The shift changes are small but larger than a resonance linewidth, consistent with an unfavorable equilibrium in fast exchange, similar to other systems in a preexisting equilibrium that is shifted by posttranslational modifications (36), suggesting either Dcp1 is directly contacting regions of Dcp2 in the closed, Trp43-dependent state, or Dcp1 is enhancing the population of the closed state but can only do so when Trp43 is present. For example, residues Val112, Ile162, Leu201, and Ile233 on the catalytic domain of Dcp2 may be directly influenced by Dcp1 because mutation of Trp43, which should also block formation of the closed state, has little effect on their chemical shift (Fig. 5). Some of the residues outside the Dcp1 binding site on Dcp2 experience Trp43-dependent exchange broadening in apo Dcp2 (Leu68, Leu74, Ile162, Ile196, Leu201, and Ile233; Fig. S2 and Fig. S4). We conclude that there is a coupling between Trp43 and Dcp1 and that Dcp1 might directly interact with regions of the Dcp2 catalytic domain (Fig. 5C).

Tryptophan 43 Couples Coactivators to Catalysis by Dcp2.

Addition of Dcp1 to Dcp2 stimulates catalysis 10-fold in vitro and is believed to enhance formation of the closed, active conformation of Dcp2 (9, 21). Therefore, if Dcp1 interacts with the catalytic domain of Dcp2 in a Trp43-dependent fashion (Fig. 5), then mutation at Trp43 should block kinetic stimulation by Dcp1. We tested this prediction directly by comparing the stimulation of Dcp2 activity afforded by Dcp1 to the stimulation in the Trp43Ala enzyme. Whereas Dcp1 simulates catalysis by wild-type spDcp2 by a factor of ten, the Trp43Ala mutant is refractory to Dcp1 stimulation (Fig. 6A). Therefore Trp43 is critical for both contact between Dcp1 and the catalytic domain in the absence of substrate (Fig. 5), and kinetic stimulation by Dcp1 in the presence of substrate (Fig. 6A).

Fig. 6.

Fig. 6.

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Mutation of Trp43Ala blocks Dcp1 activation in S. pombe and coactivation by Edc1 in S. cerevisiae. (A) Observed rates for 5 μM S. pombe decapping proteins (kmax conditions) at 0.1 °C are 0.02 min-1 for spDcp2, 0.2 min-1 for spDcp1∶Dcp2, 0.002 min-1 for spDcp2 Trp43Ala, and 0.006 min-1 for spDcp1∶Dcp2 Trp43Ala. (B) Observed rates for the S. cerevisiae Dcp1∶Dcp2(1–245) complex with and without Edc1CTR at 20 nM at 4 °C are 0.014 for scDcp1∶Dcp2, 0.18 for scDcp1∶ Dcp2 + Edc1CTR, 0.0019 for scDcp1∶Dcp2 Trp50Ala, and 0.0059 for scDcp1∶Dcp2 Trp50Ala + Edc1CTR. Rates were measured under kmax/KM conditions for budding yeast Dcp2 because reactions were too fast to follow by manual pipetting and previous work showed the Edc1CTR affects KM only threefold (17). Error is SEM from three independent experiments.

Dcp1 is believed to be a protein-protein interaction platform based on its EVH1 fold and essentiality for decapping in yeast (22, 37, 38). The S. cerevisiae protein Edc1 binds to the proline recognition site of Dcp1 and is a model coactivator of decapping (17). The C-terminal 30 residues of Edc1 (Edc1CTR) bind to Dcp1 and stimulate decapping in vitro, largely via enhancement of the rate of the catalytic step, kmax (17). Given the aforementioned results suggesting a coupling of Dcp1 to Dcp2 via Trp43, we hypothesized that mutation of this residue would abolish coactivation by Edc1. To test this hypothesis we made the corresponding mutation in the S. cerevisiae Dcp1∶Dcp2 complex (Trp50Ala) and tested its activity (Fig. 6B). Whereas the wild-type Dcp1∶Dcp2 complex was stimulated by Edc1CTR by a factor of 13, it only enhanced catalysis in the Trp50Ala mutant by a factor of three. These results are consistent with a model where Edc1CTR is contacting either the catalytic domain of Dcp2 or substrate itself, and that an interaction between Trp50 and cap mediates this coactivation.

Discussion

The activity of many enzymes is regulated by macroscopic open-to-closed transitions, for example adenylate kinase (23, 39), imidazole glycerol phosphate synthase (24), DEAD-(Asp-Glu-Ala-Asp) box helicases (40, 41), and, it seems, some ubiquitin E2 enzymes by closure of an attached ubiquitin (42, 43). Nudix enzymes can also be regulated by domains outside the Nudix motif (44, 45). In the case of Dcp2, a composite active site is formed upon cap recognition involving absolutely conserved regions on both domains (9). Tryptophan 43 is intimately tied to formation of this composite active site, as its mutation blocks cap binding to the regulatory domain, enzyme closure, and decapping in vitro and in vivo (9, 32). Ligand binding promotes closure of Dcp2 in a process that is mediated by contacts between the m7G cap structure and both the regulatory and catalytic domains of Dcp2 (9).

Previously we proposed, but did not test, the idea that Dcp1 and associated coactivators like Edc1 use the Trp43-mediated conformational change to enhance catalysis by Dcp2 (9, 17). Furthermore, it was puzzling to us how in the absence of Dcp1 and coactivators the regulatory domain could still contribute 100-fold to kcat, because closure is not observed by SAXS under these conditions (9, 17, 21). Here, we found using NMR that Dcp2 undergoes collective fast-exchange between open and closed states in the absence of any cofactors (Fig. 2, Fig. 4). An important observation is that mutation of Trp43 blocks both exchange observed by NMR and the ability of Dcp1 and Edc1CTR to enhance the catalytic step of Dcp2 (Fig. 3, Fig. 6). Therefore, it follows that cofactors such as Dcp1 and Edc1 stabilize a dynamically labile composite active site. This work provides insights into how decapping can be controlled by multiple layers of protein-protein interactions: Dcp1 binds the regulatory domain and promotes closure via Trp43, possibly by interacting with the catalytic domain of Dcp2; Edc1 likely consolidates through interactions with substrate or the catalytic domain of Dcp2. Altogether, the regulatory domain of Dcp2, its essential activator Dcp1, and the coactivator Edc1 contribute four log-units to the catalytic step (9, 17, 33), which is mediated by Trp43 (Fig. 6). Tryptophan 43 is therefore a gatekeeper of closure with (9) and without bound substrate and is central to conformational changes in Dcp2 (Fig. 1, Fig. 3). It is thus not surprising that, in yeast, mutation of Trp43 phenocopies deletion of the entire regulatory domain (32).

How are the dynamics observed in Dcp2 related to catalysis? For some enzymes like dihydrofolate reductase (5, 46), adenylate kinase (23), triosephosphate isomerase (47, 48), RNase A (49), and cyclophilin A (50) it seems that the rate-limiting step in the catalytic cycle is a conformational change. Some enzymes are even preorganized for catalysis by experiencing dynamics along the catalytic reaction coordinate without substrate (23, 50). For unliganded Dcp2, the exchange rate between conformations is much faster than kcat (∼2,300 s-1, Table S1 versus ∼0.2 min-1, Fig. 6). Because previous studies on Dcp2 showed that product release is fast (33), the catalytic step is the slow step of the catalytic cycle (33), and that closure occurs after substrate binding as a substep of kcat (9, 33), we suggest the population of closed state, not the rates of interconversion per se, imposes a limit on the catalytic rate. Preexisting equilibria play a key role in activation of signaling proteins where phosphorylation shifts a dynamic equilibrium from inactive to active states (36, 51–53). Likewise, protein cofactors of Dcp2 may shift a highly skewed dynamic equilibrium between open, inactive, and closed active states, culminating in formation of the composite active site and catalytic rate enhancement. Confirmation of this prediction requires dynamics studies in the presence of substrate, which remains a challenge for the future because we have not been able to generate stable, concentrated samples of Dcp2 in a buffer compatible with ligand binding.

There are at least two possibilities for the structural nature of the excited state of Dcp2 in the absence of substrate. One possibility is that the excited state is in fact the closed, active conformation. This model, that the apo dynamics are on-pathway, is consistent with the abrogation of cap binding by the regulatory domain, dynamics, stimulation by Dcp1, and coactivation by Edc1 upon mutation of Trp43 to alanine (Fig. 6) (9). However, we cannot exclude the possibility that the dynamics are due to an off-pathway state. For example, the Dcp1∶Dcp2 complex was crystallized in the presence of ATP in a closed conformation, so other states are not unprecedented (21). This ATP-bound closed state is neither the active state nor the excited state though, because mutation of the interdomain interface opposite Trp43 in the closed crystal state does not affect kinetics, closure by SAXS, or the nitrogen HSQC without substrate (Fig. S6) (9). Due to the central nature of Trp43 in apo dynamics, closure, catalysis, and coactivation, we favor the model that the apo dynamics are on-pathway. Independent of these distinctions is the key finding that the conserved residue Trp43 functions as a gatekeeper of the composite active site, allowing decapping activity to be stimulated by multiple layers of protein-protein interactions, which is an important regulatory event for control of 5′-3′ mRNA decay in eukaryotic cells.

Methods

Proteins were expressed in Escherichia coli and purified as described (9, 17, 33). Methyl labeling was achieved by addition of 13C 2H labeled precursors for Ile (50 mg L-1), Leu/Val (100 mg L-1), Met (250 mg L-1), and Ala (100 mg L-1) 40 min prior to induction. All Dcp2 1–243 samples were perdeuterated using 2H glucose as the sole carbon source and D2O, as described in ref. 54. NMR experiments are explained in detail in SI Methods. Briefly, de novo assignment of spDcp2 ILVMA methyls was achieved using the (H)C(CO)NH-total correlation spectroscopy (TOCSY) (55) on independent domains of spDcp2 and a methyl NOE compared to the crystal structure [Protein Data Bank (PDB) ID code 2QKM]. CPMG experiments are described (56); intensities for peaks were fit using Function and Data Analysis (FuDA, http://www.biochem.ucl.ac.uk/hansen/fuda) and CPMG dispersion curves were fit using cpmg_fitd8 (a gift from D. Korzhnev and L. Kay, University of Toronto, Toronto, ON). Enzyme kinetics were measured under single-turnover conditions as described (9, 17, 57). Structure figures were made using PyMOL (http://pymol.org).

Supplementary Material

Supporting Information
supp_109_8_2872__index.html (861B, html)

Acknowledgments.

We thank Mark Kelly and Jeff Pelton for NMR support, along with Dmitry Korzhnev for the cpmg_fitd8 software and advice and D. Flemming Hansen for Function and Data Analysis and fitting advice. This work was supported in part or in full by fellowships from the Sandler Family Foundation for Basic Sciences and the Achievement Awards for College Scientists (ARCS) Foundation (S.N.F.), a US National Institutes of General Medical Sciences predoctoral fellowship 1R25GM56847 (to M.S.B.), a US National Institutes of Health Grant R01GM078360 (to J.D.G.), and a US National Institutes of Health Grant GM68933 for the Central California 900-MHz facility.

Footnotes

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1113620109/-/DCSupplemental.

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