Introduction
mRNA degradation plays a key role in the regulation of gene expression. Several mRNA decay pathways exist in eukaryotic cells (
Caponigro and Parker, 1996). In the 5′–3′ mRNA decay pathway, poly(A) shortening is followed by cleavage of the mRNA cap, exposing the mRNA body to 5′–3′ exonucleases. The nonsense‐mediated decay (NMD) pathway involves, likewise, decapping followed by 5′–3′ digestion of the mRNA. However, during NMD these steps are preceded by the recognition of a premature stop codon present in the (pre‐)mRNA rather than by deadenylation (
Czaplinski et al., 1999). The two other known eukaryotic mRNA turnover pathways rely on the activity of the exosome, a multisubunit complex endowed with 3′–5′ exonuclease activity (
Butler, 2002). These are the 3′–5′ decay and the non‐stop decay (NSD) pathways. The former process involves deadenylation followed by 3′–5′ decay of the mRNA body. In contrast, during NSD, the exosome‐mediated degradation of the mRNA is triggered by the absence of a stop codon. In yeast and human cells, NMD and NSD allow the removal of aberrant mRNAs in the cytoplasm and unspliced pre‐mRNAs that could be deleterious to the cell (
He et al., 1993;
Frischmeyer et al., 2002;
van Hoof et al., 2002). Various proposals have been made for the location of NMD in metazoan cells: some models suggest that NMD occurs in the cytoplasm while others suggest that it happens exclusively, or in addition, in the nucleus where translation and ribosomes have been detected (
Iborra et al., 2001;
Ishigaki et al., 2001;
Brogna et al., 2002). The 3′–5′ mRNA turnover pathway has also been shown to be involved in the decay of pre‐mRNAs in the yeast nucleus (
Bousquet‐Antonelli et al., 2000).
As in yeast, both the 5′–3′ and 3′–5′ pathways probably contribute to mRNA decay in mammals. Indeed, deadenylated and decapped mRNAs, which are characteristic decay intermediates of the 5′–3′ pathway, have been detected (
Couttet et al., 1997). In contrast,
in vitro experiments indicated that substrate mRNAs are degraded mostly by the 3′–5′ pathway (
Chen et al., 2001;
Mukherjee et al., 2002). This pathway is activated
in vitro by the presence of AU‐rich elements (AREs) in the mRNA substrate, consistent with observations demonstrating that these sequences increase mRNA instability. These latter observations led to the suggestion that the 3′–5′ pathway is the major pathway for mRNA degradation in human cells. However, the proportion of (specific) mRNAs degraded by the 3′–5′ or the 5′–3′ decay pathway
in vivo remains to be established.
Several cellular factors implicated in mRNA decay have been identified. These include deadenylases (
Daugeron et al., 2001;
Tucker et al., 2001) and exonucleases (
Larimer et al., 1992;
Mitchell et al., 1997). Two different decapping activities have been described. In yeast, Dcp1 was reported to be the major decapping enzyme cleaving capped RNA to release m7GDP and a 5′‐phosphorylated downstream product that is a substrate for the Xrn1 5′–3′ exonuclease (
Beelman et al., 1996). dcp1 mutants are unable to perform this process, leading to the stabilization of numerous mRNAs. dcp1 mutants are also defective in NMD. Dcp2 was identified as a multicopy suppressor of dcp1 mutants (
Dunckley and Parker, 1999). dcp2 mutants also abolish a large fraction of mRNA turnover in yeast cells. Dcp2 was proposed to be an activator of Dcp1 since no intrinsic decapping activity was observed for Dcp2 (
Dunckley and Parker, 1999). Interestingly, the Dcp2 protein contains a highly conserved MutT/Nudix motif. This motif has been found in a wide variety of proteins that possess the capability to hydrolyse a
nucleoside
diphos phate linked to some other moiety,
X (nudiX) and has been shown to be critical for catalytic activity (
Bessman et al., 1996). The Dcp2 protein is also required for NMD and was shown to interact with Upf1 (
He and Jacobson, 1995). It is noteworthy that the decapping activity mediated by Dcp1 and Dcp2 is required, and tightly controlled both for the 5′–3′ and NMD mRNA decay pathways. Pat1 and a group of seven Lsm proteins are required for mRNA decapping in the 5′–3′ mRNA decay pathway but not in the NMD pathway (
Bouveret et al., 2000;
Tharun et al., 2000). Conversely, the Upf1, Upf2 and Upf3 factors specifically required to initiate NMD do not affect decapping in the 5′–3′ pathway (
Czaplinski et al., 1999).
Recently, a different decapping activity was identified in human cell extracts. This activity, cleaving caps that have been released from mRNA through the activity of 3′–5′ exonucleases, has been named DcpS denoting the scavenger decapping activity of the corresponding factor (
Wang and Kiledjian, 2001). The products of the reaction mediated by DcpS and using m7GpppG as a substrate are m7GMP and GDP and differ thus from the products of the Dcp1‐mediated reaction. Because DcpS activity requires prior degradation of the mRNA body it is likely to act downstream of the exosome in the 3′–5′ and NSD mRNA decay pathways.
To gain further insight into the process of mRNA decapping and turnover, we set out to identify and characterize human homologues of the yeast Dcp1 and Dcp2 proteins. Surprisingly, analysis of recombinant human Dcp2 (hDcp2) revealed that it has decapping activity, identifying it as new decapping enzyme. Interestingly, hDcp2 co‐localizes with hDcp1 in specific sub‐structures in the cytoplasm of human cells suggesting that mRNA decay may occur in a dedicated compartment.
Discussion
Our results demonstrate that hDcp2 is a new mRNA decapping enzyme. Recombinant hDcp2 catalyses the decapping of capped RNA generating m7GDP and a 5′‐phosphorylated RNA fragment that is a substrate for 5′–3′ exonucleases such as Xrn1. hDcp2 differs from human DcpS activity by the nature of the product and by its ability to cleave cap structures attached to long mRNA molecules. Recombinant yeast Dcp2 has been reported to be inactive for decapping
in vitro (
Dunckley and Parker, 1999). We were unable to test the activity of full‐length yeast Dcp2 as expression of this large (108 kDa) protein in
E.coli was not successful (F.Wyers, personal communication). Our work indicates, however, that a recombinant fragment of yeast Dcp2 containing the conserved N‐terminal moiety was active for decapping (
Figure 5). The region required for hDcp2 activity is also located at the N‐terminus of the protein, which is the most conserved part. This region contains a MutT/Nudix domain and mutation of conserved residues in this domain ablates the decapping activity of hDcp2. Similar mutations completely abolish the activity of yeast Dcp2 in
vivo, thus blocking decapping (
Dunckley and Parker, 1999). The decapping activity of hDcp2 has an alkaline pH optimum, a property that is also consistent with characteristics of known enzymes harbouring an active MutT/Nudix catalytic domain. The various enzymes of this family have been shown to cleave substrates carrying a pyrophosphate group to release a phosphate containing product and the remaining part of the substrate molecule. Similarly, the reaction catalysed by hDcp2 produces m7GDP and a downstream 5′‐phosphorylated RNA. Interestingly, hDcp2 requires the presence of a long RNA molecule attached to the cap and is stimulated by the presence of the methyl7 group. This substrate specificity differs from what is known for other MutT/Nudix proteins, demonstrating that capped mRNAs are natural substrates for Dcp2. Taken together, these data strongly suggest that hDcp2 is a bona fide decapping enzyme.
The yeast Dcp1 protein has been reported to be a decapping enzyme generating m7GDP (
Beelman et al., 1996;
LaGrandeur and Parker, 1998;
Fischer and Weis, 2002). The involvement of the yeast Dcp1 protein in decapping
in vivo is well established as deletion or mutation of the corresponding gene blocks decapping. Recombinant yeast Dcp1 expressed in
E.coli was originally reported to be inactive in decapping (
Beelman et al., 1996;
LaGrandeur and Parker, 1998). Yeast Dcp1 overproduced and purified from yeast was, however, active. The difference in activity of the two protein preparations was attributed to the lack of phosphorylation of the recombinant protein. More recently, the preparation of active recombinant yeast Dcp1 was reported (
Vilela et al., 2000). Thus, while there is no doubt that in yeast, Dcp1 is required for decapping
in vivo, two models can be proposed to explain the function of Dcp1 in the decapping reaction. There is first the possibility that Dcp1, like Dcp2, is an active decapping enzyme. Our inability to detect activity associated using recombinant hDcp1 produced in
E.coli or by
in vitro translation under a variety of conditions (
Figure 1 and data not shown) would thus reflect a lack of modification or aberrant folding of the protein. Alternatively, it remains possible that the decapping activity originally attributed to Dcp1 was not mediated by Dcp1 itself. Purification of yeast Dcp2 reveals that it copurifies with yeast Dcp1 but is quite unstable leading to the generation of a smear of truncated products probably through the action of non‐specific proteases during the purification (F.Caspary and B.Séraphin, unpublished results). It remains therefore possible that Dcp1 fractions obtained from yeast contained a significant amount of truncated yeast Dcp2 responsible for the observed activity (even if the corresponding polypeptide was not detected by staining after gel electrophoresis). However, this possibility is somewhat difficult to reconcile with the fact that denaturing gel electrophoresis followed by renaturation of Dcp1 yields a protein capable of decapping (
Beelman et al., 1996;
LaGrandeur and Parker, 1998). The detection of cap‐cleavage in the presence of recombinant yeast Dcp1 also suggests that this protein might be catalytically active. It is noteworthy, however, that the exact nature of the products generated by the recombinant protein purified from
E.coli was not assessed. We have observed a background cap‐cleaving activity from
E.coli cleaving free cap but only weakly affected by cap methylation (data not shown). In this vein, it is important to note that recombinant Dcp1 proteins produced in
E.coli appeared to have low activities per weight unit of protein. In summary, while we cannot exclude that hDcp1 is a second decapping enzyme, our results clearly demonstrate that both yeast and human Dcp2 are active. The exact function of hDcp1 remains therefore to be established. The phenotype of yeast Dcp1 mutants, the co‐purification of yeast Dcp1 and Dcp2, sequence conservation throughout evolution and the co‐localization of hDcp1 and hDcp2 in the cytoplasm leave however little doubt that hDcp1 is required for some aspect of the decapping reaction or more generally for mRNA decay. While this work was in progress, hDcp1 was also identified as a factor interacting with Smad4 and implicated in transcriptional regulation (
Bai et al., 2002). Further studies will reveal whether Dcp1 affects mRNA degradation and transcriptional regulation independently.
In yeast, the 5′–3′ mRNA decay pathway appears to be the major mode of mRNA degradation
in vivo (
van Hoof and Parker, 2002). The situation is less clear for human cells. mRNA decay intermediates lacking a cap structure have been identified, indicating that the 5′–3′ mRNA decay pathway is also active in these cells (
Couttet et al., 1997). In addition, a decapping activity potentially generating m7GDP has been reported (
Gao et al., 2001). The finding that Dcp2 is active in decapping supports this conclusion and suggests further that it might have been responsible for the production of the intermediates detected. Previous studies have revealed that only a small proportion (3%) of poly(A)
− RNA population lacks a cap (
Couttet et al., 1997). However, given that the steady state level of intermediates will be governed by the relative kinetics of their formation and disappearance, further work will be required to determine the quantitative contribution of the 5′–3′ pathway to overall mRNA decay in human cells. Incubation of synthetic mRNAs in human cell extracts indicated that those are essentially degraded in a 3′–5′ manner
in vitro and suggested that this pathway is specifically regulated by ARE elements (
Chen et al., 2001;
Mukherjee et al., 2002). It remains to be demonstrated, however, whether the natural mRNA degradation pathway with intricate regulation (e.g. deadenylation by bona fide enzymes preceding decapping or 3′–5′ exonucleolytic degradation by the exosome) is faithfully reproduced
in vitro in the absence of processes such as translation. Indeed, while 5′–3′ degradation is the main route for mRNA degradation in yeast, RNA introduced into cells appear to be degraded by the 3′–5′ mRNA decay pathway, suggesting that degradation directionality may be highly sensitive to small perturbations of the system (
Brown and Johnson, 2001). Overall, the existence of mRNA decay intermediates lacking a cap together with the conservation of factors implicated in the 5′–3′ pathway (Lsm proteins, Pat1, Xrn1) in human cells (
Bouveret et al., 2000) clearly demonstrate that an mRNA decapping activity is functional in these cells. However, the contribution of hDcp2 to the overall mRNA decay process as well as its involvement in specific pathways remains to be established.
Our investigation of the intracellular localization of hDcp1 and hDcp2 led to the surprising observation that these proteins co‐localize in specific substructures in the cytoplasm. While a cytoplasmic localization supports a role for both of these proteins in mRNA decay, their presence in specific substructures was unexpected. Two major suggestions can be made for the role of these structures. First, these structures could represent a storage compartment for mRNA decay factors. Alternatively, these structures may correspond to the location of an active site of mRNA degradation. We favour the latter hypothesis because there is no evidence for storage of mRNA decay factors that would have to be activated under specific conditions. It is worth mentioning that related structures containing RNA and RNA‐binding proteins have been described recently in human cells (
Eystathioy et al., 2002). If these two structures are indeed identical, substrate molecules would be present together with mRNA decay enzymes, supporting an active rather than a storage function. Interestingly, the Xrn1 protein has also been shown to be enriched in specific dots in the cytoplasm of mouse cells (
Bashkirov et al., 1997). Beside the identification of a new pattern of cytoplasmic staining, the observation that mRNA decay factors are unevenly distributed in the cytoplasm suggests that mRNA decay may be precisely localized. Such a situation would enable cells to remove specific mRNAs from particular locations in the cytoplasm. Defining the exact function and composition of these structures in the near future should provide new insights into the mechanism and control of mRNA decay.
Material and methods
Plasmid construction
cDNA clones for hDcp1 (IMAGp998B07136Q2, pBS2154) and hDcp2 (IMAGp998B233419Q2, pBS2155) were obtained from the IMAGE EST project (
Lennon et al., 1996).
To construct an expression clone of hDcp1, the original cDNA was PCR amplified with a 5′ oligo containing a NcoI site and the first five nucleotides (nt) of the hDCP1 coding sequence lacking in the cDNA; the 3′ oligo contained a XhoI site. The PCR product digested with NcoI and XhoI was cloned in the corresponding sites of an expression vector carrying a His6 and a GST tag. The coding sequence of the resulting plasmid (pBS2175) was free of mutations. Digestion of pBS2175 with NheI and XhoI, blunting and religation generated a C‐terminal truncation of 154 amino acids (aa). The resulting plasmid (pBS2176) encodes the first 428 aa of hDcp1 followed by eight amino acids derived from the vector.
Approximately 200 nt lacking at the 5′ end of the original Dcp2 cDNA were recovered by RACE from HeLa cell poly(A)+ mRNAs using specific oligonucleotides (pBS2183). This fragment was combined with a cDNA containing the downstream Dcp2 ORF and inserted in an expression vector yielding pBS2204.
Point mutations were introduced in the hDcp2 coding sequence using the Quikchange kit (Stratagene). Two A/C nucleotide substitutions were introduced changed residues E147 and E148 to alanine (pBS2270). The region coding for the first 250 aa of hDcp2 was PCR amplified using appropriate primers containing a stop codon and XhoI site. The product was digested with BsrGI and XhoI and inserted into the same sites of plasmid pBS2204, giving pBS2271.
Modification of pBS2204 allowed the insertion of the hDcp2 coding sequence between a N‐terminal GST tag and a C‐terminal His6 tag. The His6 tag of pBS2204 was deleted by replacing the XbaI–SwaI fragment by an XbaI–SwaI fragment from pGSTevCBP (L.Minvielle‐Sebastia) yielding pBS2311. An NcoI–AvaII fragment of the hDcp2 coding sequence was fused with an adapter containing an AvaII site, a His6 coding sequence, a stop codon and an XhoI site. The fusion product was cloned in the NcoI–XhoI sites of pBS2311 yielding pBS2312.
For in vitro translation, the hDcp1 and hDcp2 ORF were cloned in the NcoI–SmaI sites of plasmid pIVEX2.4a (Roche) yielding plasmid pBS2229 and pBS2230, respectively.
In vitro decapping assays
Unlabelled and uncapped RNA was produced by
in vitro transcription with T7 polymerase (Promega). Plasmid pBS2266 (a pGEM‐3Zf+ derivative containing a cloned synthetic insert) digested with
FokI was used as template, yielding a transcript of 49 nt. Digestion of pBS2266 with
SmaI or
EcoRI generated transcripts of 23 and nine nt. RNAs were purified from a 7 M urea–8% polyacrylamide gel, eluted overnight in elution buffer (0.3 M NaAc, 50% phenol, 1 mM EDTA pH 5.2), chloroform‐extracted and ethanol‐precipitated. Between 20 and 50 pmol of these RNAs were cap‐labelled using the vaccinia virus capping enzyme (
Higman et al., 1994) in capping buffer (50 mM Tris–HCl pH 7.9, 1.2 mM MgCl
2, 6.0 mM KCl, 2.5 mM DTT) in the presence of 1 mM SAM, 2.5 μM unlabelled GTP, 0.132 μM [α‐
32P]GTP, 40 units RNasin and capping enzyme. Specific activity, determined after gel‐filtration and ethanol precipitation, was measured by counting; 5000–9000 c.p.m. used per reaction corresponded to 0.01–0.02 pmol RNA.
Recombinant proteins were expressed and purified on Ni‐NTA (Qiagen). For the GST–hDcp2‐His6 protein, the Ni‐agarose purification was followed by GST purification using glutathione–Sepharose 4B beads (Amersham). Purified proteins were dialysed against 20 mM HEPES–KOH pH 7.6, 0.01 % NP‐40, 20 mM KCl, 1 mM MgCl2, 10% glycerol, 0.1 mM EDTA, 1 mM DTT. In vitro translated full‐length hDcp1 and hDcp2 were obtained using the Rapid Translation System RTS500 E.coli HY kit (Roche) and the RTS500 Instrument (Roche).
The decapping reactions (
Zhang et al., 1999) were performed at 37°C for 30 min in decapping buffer (45 mM Tris–HCl pH 8, 27 mM (NH
4)
2SO
4, 9 mM MgAc) in the presence of 0.25 μg/μl tRNA and 26 ng to 2 μg of recombinant protein. While an absolute specific activity was not determined, we observed that, at non‐saturating substrate concentrations, <26 ng of recombinant Dcp2 were sufficient for the decapping of 0.1 pmol of substrate in 10 min. Aliquots of the reactions were run on PEI‐cellulose TLC plates (Merck) in 0.3–0.5 M LiCl, 1 M formic acid. TLC plates were dried and exposed to PhosphorImager screens (Molecular Dynamics). Nuclease P1 treatment of cap‐labelled RNA was carried out with 1 U enzyme in 10 mM Tris pH 7.5.
Downstream product analysis
Uniformly labelled RNAs were transcribed in vitro with T7 polymerase (Promega) in the presence of [α‐32P]UTP with or without m7GpppG for capped or uncapped RNA, respectively, using pBS2266 linearized with SmaI as template. RNAs were purified as described above. 3000 c.p.s. were used for in vitro decapping. Reactions were stopped with formamide loading dye and run on 7 M urea–8% polyacrylamide denaturing gels. Dried gels were exposed to PhosphorImager screens. CIP (Biolabs) treatment was carried out with 10 U of enzyme for 30 min at 37°C.
Cell culture and transfection for GFP localization
The cDNAs encoding hDcp1 or hDcp2 were cloned in peGFP‐C2, peCFP‐C1 and peYFP‐C1 plasmids (Clontech). This yielded the following constructs: hDcp1‐GFP (pBS2201), hDcp1‐CFP (pBS2278), hDcp1‐YFP (pBS2265), hDcp2‐GFP (pBS2315), hDcp2‐CFP (pBS2269) and hDcp2‐YFP (pBS2277). As a control, the hSnu30‐2 coding sequence from pBS2108 was inserted in the peYFP‐C1 vector giving plasmid pBS2293.
For stable transfection, 3 × 106 HEK293 cells were transfected with 1 μg plasmid DNA using the Effectene Transfection Reagent (Qiagen) and were cultured for 2 days without selection agent. Cells were then treated with 400 μg/ml G418 (Invitrogen) and selected for 2–3 months on selective media. Cells were cloned and selected for expression of the fluorescent protein. Transient transfections were performed using the same protocol but using 10 μg of plasmid DNA. For the co‐localization experiments, stably transfected cell lines were transiently transfected, either with CFP‐hDcp1, CFP‐hDcp2 and/or YFP‐hSnu30‐2, as described above. Next, the preparations were analysed using a confocal microscope (Leica RCS SP2) on an inverted stand using objectives HC Plan APO OS 100X oil NA 1.4.
Immunofluorescence
HEK293 cells were grown in DMEM Glutamax (Invitrogen) and 10% FCS on clean glass slides to 70% confluence. The cells were washed in PBS and fixed for 20 min in 4%
p‐formaldehyde. After several washes, cells were permeabilized in 0.1% Triton X‐100 for 10 min. Then, cells were saturated for 30 min with 0.2% BSA/PBS and after several washes cells were immunostained for 1 h at 37°C with a rabbit anti‐hDcp1 serum (1:1000). After subsequent washes, cells were incubated 1 h at 37°C with secondary antibody [Fluorolink Cy 5‐labelled goat anti‐rabbit IgG (Amersham) or goat anti‐rabbit IgG FITC‐conjugated (Sigma) (1:1000)]. The anti‐hDcp1 serum used for immunofluorescence revealed specifically hDcp1 in western blotting analyses (
Figure 6A). Nuclei were stained with propidium iodide (5 μg/ml) for 30 min in a dark chamber. Finally preparations were analysed by confocal microscopy as described above.
Note added in proof
While this work was in progress, mRNA decapping by hDcp2 was independently reported by Wang et al. (Proc. Natl Acad. Sci. USA, 2002, 99, 12663–12668) and Lykke‐Andersen (Mol. Cell. Biol., 2002, in press).