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The activity of C. elegans scavenger decapping enzyme (DcpS) on its natural substrates and dinucleotide cap analogues modified in the nucleoside’s base or ribose moiety, has been examined. All tested dinucleotides were specifically cleaved between β and γ phosphate groups in the triphosphate chain. The kinetic parameters of enzymatic hydrolysis (Km, Vmax) were determined using fluorescence and HPLC methods, as complementary approaches for the kinetic studies of C. elegans DcpS. From the kinetic data, we have determined which parts of the cap structure are crucial for DcpS binding and hydrolysis. We show that m32,2,7GpppG and m32,2,7GpppA are cleaved with higher rates than their monomethylated counterparts. However, C. elegans DcpS higher specificity for MMG caps is illustrated by lower Km values. Modifications of the first transcribed nucleotide did not affect the activity, regardless of the type of purine base. Our finding suggests C. elegans DcpS flexibility in the first transcribed nucleoside-binding pocket. Moreover, while C. elegans DcpS accomodates bulkier groups in the N7 position (ethyl or benzyl) of the cap, either 2′-O-or 3′-O-methylations of the 7-methylguanosine result in two orders of magnitude reduction in hydrolysis.
mRNA turnover is one critical determinant in the regulation of gene expression [1–3]. Degradation of normal transcripts in eukaryotes occurs along two major pathways, 5′→3′ and 3′→5′decay, both initiated by shortening the poly(A) tail [4,5]. In the 5′→3′ decay pathway, deadenylation is followed by Dcp1/Dcp2 mediated decapping, which exposes the body of the transcript to Xrn1 exonuclease [6,7]. In the 3′→5′decay pathway, deadenylation facilitates access to the mRNA 3′ end by a complex of nucleases, known as the exosome, which degrades the mRNA chain 3′→5′ until it reaches the cap-containing dinucleotide or a short capped oligonucleotide [8,9]. The residual cap structure m7GpppN (7-methylGpppN) is further hydrolyzed by the scavenger decapping enzyme DcpS . Capped dinucleotides or oligonucleotides accumulated in cells could bind to cap-binding proteins such as eIF4E and inhibit translation . Hydrolysis of cap dinucleotides in this context is thought to be important. However, mutations in DcpS are generally not lethal suggesting the possibility that other undiscovered and redundant scavenger enzyme activities may be present [11,12].
Decapping scavengers have been characterized in yeast (Saccharomyces cerevisiae and Saccharomyces pombe), nematode (Caenorhabditis elegans and Ascaris suum) and mammalian (mouse and human) cells [13–15]. DcpS proteins constitute their own branch within the HIT family of pyrophosphatases with decapping activity as the main, well defined biological function [16,17]. All these enzymes exhibit high specificity for cap structure and limited activity towards non-methylated dinucleotides (e.g. ApppA and GpppG). Decapping scavengers utilize an evolutionary conserved Histidine Triad motif (HIT) to cleave the 5′-ppp-5′ pyrophosphate bond within the cap, releasing m7GMP [15–17]. Sequence alignment of DcpS proteins from different organisms demonstrated the presence of a conserved hexapeptide containing a histidine triad (HIT) with three histidines separated by hydrophobic residues (His-ϕ-His-ϕ-His-ϕ). Structural analysis has revealed that HIT proteins exist as homodimers containing nucleotide binding pockets with respect to the three histidine residues of the catalytic HIT motif [18–20]. A high degree of identity observed in the HIT region of different scavengers supports the functional significance of this domain in decapping activity. Substitution mutagenesis of the central histidine in human and nematode decapping scavengers inactivates their hydrolytic properties, demonstrating that the central HIT motif is critical for catalysis [14,20]. This histidine is involved in the formation of a covalent nucleotidyl phosphohistidyl intermediate, the nucleophile agent for the γ phosphate group of dinucleoside triphosphate substrates [19,20].
The process of mRNA turnover is more complicated in nematodes, because they have two populations of mRNAs, each with a distinct cap structure. ~70% of nematode mRNAs possess a trimethylguanosine cap m32,2,7GpppG, while ~30% have a typical cap structure m7GpppG . Both types of mRNA interact with polysomes and undergo translation [12,22]. The presence of two populations of mRNAs has profound implications for proteins that recognize specifically each mRNA . The eIF4E protein in C. elegans exists in 5 different isoforms, with different affinity to m7GpppG and m32,2,7GpppG [20,21]. Human and yeast DcpS can effectively hydrolyze only the m7GpppG cap, and human DcpS has activity on capped oligonucleotides up to 10 nucleotides [22–24]. In contrast, initial studies on the nematode decapping scavenger indicated that both trimethylated and monomethylated caps and oligonucleotides up to 4 nucleotides were hydrolyzed .
Previous data suggest that the substrate specificity of C. elegans DcpS differs from its human and yeast orthologs [3,14,25,26]. However, neither detailed kinetic analysis of enzymatic cleavage nor mechanisms of substrate recognition have been carried out on C. elegans DcpS. Here, we have studied substrate specificity and kinetic analysis of recombinant C. elegans scavenger decapping enzyme. Various dinucleotide cap analogues, natural and chemically modified within 7-methylgunosine moiety and the first transcribed nucleoside have been investigated as potential substrates. Kinetic parameters (Km, Vmax and Vmax/Km) were determined to characterize the hydrolytic activity of C. elegans DcpS.
To identify the DcpS hydrolysis products of all investigated dinucleotides presented in Fig. 1, HPLC chromatograms were analyzed. As an example, chromatographic analysis for the cleavage of MMG, TMG and GpppG are shown on Fig. 2. For m32,2,7GpppG, the peak corresponding to the substrate disappeared after 10 minutes of reaction (panel A). MMG was almost completely hydrolyzed during 20 minutes (panel B). The hydrolysis of GpppG was much slower, after 120 minutes a considerable amount of the substrate was still observed in the reaction mixture (panel C). The analysis of hydrolysis products (Table 1) demonstrates that the cleavage of cap analogues occurs exclusively between β and γ phosphate groups within the triphosphate bridge. These data confirm the earlier observations that nematode DcpS utilizes the same mechanism of catalysis as it was proposed for the other HIT pyrophosphatases cleaving the cap structure, and the highly conserved HIT motif is involved in binding of the substrates and catalysis [19,20].
Initial studies on the substrate specificity of recombinant C. elegans DcpS suggested that the protein was specific for 7-methylguanosine nucleotides. The first quantitative experiments characterizing this enzyme were reported by Kwasnicka et al. . However, specificity of C. elegans DcpS was defined with m7GpppBODIPY, GpppBODIPY and ApppBODIPY, but not with natural caps m7GpppG or m32,2,7GpppG. Methylated mono-and dinucleotides (m7GDP, m7GTP, m7GpppG, m32,2,7GpppG) have been only examined as inhibitors of C. elegans scavenger in the hydrolysis process of m7GpppBODIPY. Inhibition constant calculated for m32,2,7GpppG (Ki = 28.1 ± 2.5 μM), 8-fold higher than for m7GpppG (Ki = 3.47 ± 0.84 μM), indicated less efficient inhibitory properties of trimethylated cap in comparison with its monomethylated counterpart. On the basis of these findings it was concluded that TMG cap may not be a substrate for C. elegans DcpS. In subsequent studies both MMG and TMG caps were shown to be hydrolyzed by C. elegans scavenger (cellular extract and recombinant protein), but the substrate affinity and kinetics of this reactions with the substrates were not quantitatively determined . To make detailed comparison of C. elegans DcpS activity for the natural mono- and trimethylated caps, we carried out kinetic studies of hydrolysis of m7GpppG, m7GpppA, m32,2,7GpppG and m32,2,7GpppA, using a fluorimetric method. The Michaelis-Menten curves (vo versus co) obtained for these compounds are presented in Fig. 3.
The initial velocity data showed that the kinetics for MMG and TMG cap are hyperbolic in the investigated concentration range, 0.5 – 86 μM for m7GpppG and 0.5 – 97 μM for m32,2,7GpppG. The kinetic parameters derived for these reactions, Michaelis constants (Km), maximum velocities (Vmax) and pseudo-first order rate constants (Vmax/Km) are summarized in Table 1. The Km and Vmax values are about 3 times higher for TMG cap than for MMG cap, whereas the Vmax/Km is almost the same. This indicates that C. elegans DcpS has slightly different substrate specificity for these natural compounds, with a preference for m7GpppG, as previously suggested [25,26]. However, the rate of hydrolysis catalyzed by C. elegans DcpS is higher for TMG cap.
To further examine the substrate specificity of C. elegans DcpS, the hydrolysis of several other dinucleotide cap analogues was examined. Substitution of adenine for guanine as the second nucleotide in MMG and TMG caps did not significantly changed the substrate properties of m7GpppA and m32,2,7GpppA for DcpS catalysis when compared to m7GpppG and m32,2,7GpppG, respectively (Table 1). Similarly, monomethylated cap dinucleotides of the type m7GpppN, modified within the first transcribed nucleoside (N = m6A, m7G, 2′dG, m2′-OG) were all good DcpS substrates as illustrated by the kinetic data (Fig. 3, Table 1). The Km and Vmax values for these four compounds are similar to that obtained for MMG cap, indicating that C. elegans DcpS tolerates different modifications within the first transcribed nucleoside. The data presented here show that the second nucleotide of the cap structure is not crucial for catalytic mechanism of C. elegans DcpS.
The next interesting part of our studies concerning substrate requirements for C. elegans DcpS revealed that the enzyme tolerates different sized substituents at the N7 position of the m7Guo (7-methylguanosine). The kinetic data (Km, Vmax and Vmax/Km) calculated for m7GpppG (7-methylGpppG), et7GpppG (7-ethylGpppG) and bn7GpppG (7-benzylGpppG) clearly show that all these three compounds are similarly recognized as substrates by nematode scavenger (Table 1). These findings suggest plasticity within the C. elegans DcpS cap-binding pocket.
We also examined m27,2′-OGpppG and m27,3′-OGpppG (bearing additional methylation at the 2′ or 3′ oxygen of the m7Guo) as C. elegans DcpS substrates (Fig. 3). Km values determined by the fluorimetric and HPLC methods for both compounds are significantly higher than for m7GpppG (Table 1). Furthermore, for m27,3′-OGpppG the rate of hydrolysis is drastically reduced. This compound has been previously studied as an effective inhibitor of m32,2,7GpppA hydrolysis catalyzed by C. elegans DcpS, with Ki = 1 μM , significantly lower than Km value (~14 μM) determined in this study (Table 1). Such low Ki value indicates tight binding of the m27,3′-OGpppG with DcpS, whereas Km involves contribution from dissociation step including product release, which may be very slow in m27,3′-OGpppG hydrolysis. As the inhibition type has not been determined, it is not obvious, that m32,2,7GpppA and m27,3′-OGpppG compete for the same binding site in the inhibitory experiment .
The kinetic parameters obtained for m27,2′-OGpppG and m27,3′-OGpppG indicate that 2′-OH and 3′-OH positions in the ribose ring of 7-methylguanosine moiety play a significant role in catalytic activity of C. elegans DcpS.
A series of modified dinucleotide cap analogues studied in this work defined several structural requirements for substrate specificity towards C. elegans DcpS. We find that cleavage of the cap structure occurs exclusively between β and γ phosphate groups in the triphosphate chain. We examined the ability of the enzyme to act on various cap analogues in a quantitative manner, employing two independent methods (fluorescence and HPLC) to determine kinetic data.
Among different scavengers investigated (human, nematode, yeast), C. elegans DcpS has a unique property, i.e. the possibility to hydrolyse both monomethylated (m7GpppG and m7GpppA) and trimethylated (m32,2,7GpppG and m32,2,7GpppA) cap structures. Our kinetic data demonstrate that trimethylated cap analogues (m32,2,7GpppG and m32,2,7GpppA) are cleaved with higher rates than their monomethylated counterparts (Table 1). However, MMG caps are recognized with higher specificity indicating the two additional methyl groups at N2 position in TMG caps account for the Km differences for these substrates.
In agreement with previous data for nematode and human DcpS [14,20], we observed very low activity of C. elegans DcpS for the unmodified dinucleotide GpppG (Fig. 2). These results clearly show that for tight and specific binding of the base moiety to the enzyme the positive charge is required at N7 position, introduced by a methyl or any alkyl group. Different sized substituents (methyl, ethyl, benzyl) introduce positive charge into the base moiety, which is a key feature for recognition of the cap structure. Amino acids involved in the stacking interactions with the methylated base are not conserved in different organisms (Fig. 4) and thus apparently not crucial for hydrolytic activity, as indicated by the mutation L206A retaining over 90% of wild-type activity of human DcpS . Substrate properties of N7 alkylated dinucleotides (m7GpppG, et7GpppG, bn7GpppG) do not significantly differ as indicated by kinetic parameters presented in Table 1. These data indicate that the cap-binding pocket of C. elegans DcpS is inherently flexible and able to accommodate different cap structures. This flexibility may explain why significantly large groups as ethyl or benzyl can interact with nematode scavenger and be hydrolyzed with comparable rate.
To investigate the catalytic mechanism of C. elegans DcpS with respect to the first transcribed nucleoside of the cap structure, we made a detailed, quantitative comparison of the kinetic parameters for various cap analogues modified in the first transcribed nucleoside. We established that modifications introduced into the first transcribed nucleoside do not significantly influence nematode DcpS kinetic parameters. The substitution of adenine for guanine in m7GpppG or m32,2,7GpppG does not affect the Km values. Other cap analogues bearing modifications of Guo, such as m6A, m7G, m2′-OG and 2′dG, have similar kinetic parameters to m7GpppG, indicating that modifications of the base or ribose moiety within the first transcribed nucleotide are not crucial for substrate recognition or the rate of hydrolysis. What’s more, the Km value for m7GpppG, (1.17 ± 0.14) μM, is remarkably similar to the Km reported for m7GpppBODIPY, (1.21 ± 0.05) μM, containing an artificial fluorescent probe BODIPY instead of guanine . C. elegans DcpS thus can accept different, even non biological substituents instead of the first transcribed nucleotide, which affect neither the substrate specificity nor the hydrolysis rate.
Similar effect was observed for human DcpS. Mutagenesis of the human DcpS amino acids responsible for the contacts with the first transcribed nucleoside had little effect on enzyme activity, suggesting the structure of the binding pocket recognizing the first transcribed nucleoside is more flexible than the cap-binding pocket . As it was shown in Fig. 4, the amino acids recognizing the first transcribed nucleoside are not conserved in DcpS homologs, indicating that interaction with this nucleoside is not very important for decapping activity. We thus propose that DcpS proteins exhibit structural plasticity for the first transcribed nucleoside, which has no affect on the enzyme hydrolysis.
Substrates modified by additional methylation at the 2′ or 3′ oxygen of the m7Guo Kinetic parameters obtained for m27,2′-OGpppG and m27,3′-OGpppG demonstrated the crucial role of 2′-OH and 3′-OH groups of 7-methylguanosine moiety for C. elegans DcpS hydrolysis. The 2′-O-Me and 3′-O-Me analogues are so called ARCA (Anti-Reverse Cap Analogues) which are commercially available and used as substrates for in vitro transcription reactions [27,28]. Such analogues prevent their reverse incorporation into mRNAs, thus produce transcripts which are more efficiently translated than those prepared with m7GpppG. The transcripts obtained by this method are commonly used for numerous studies because they mimic well natural transcripts, e.g. in initiation of translation (methylation of ribose of 7-Guo do not disturb the interaction with eIF4E) . We have established that in some studies ARCA-prepared transcripts may not be a good mimic of natural transcripts (this DcpS study is a good example). As indicated by high Km values and very low Vmax/Km values, both these compounds are poor substrates for C. elegans DcpS. Interestingly, 2′-O- and 3′-O- methylations mean various susceptibility of the cap to enzymatic hydrolysis. Despite the fact, that efficiencies of hydrolysis are two order of magnitude reduced comparing to natural substrate, the kinetic parameters (Km and Vmax) are significantly different. Although the leaving group is the same as in MMG cap (GDP), the rate of hydrolysis observed for m27,3′-OGpppG is significantly lower, suggesting that slow dissociation of enzyme-product complex might be a control step in the hydrolysis process. With respect to substrate specificity, the loss of a hydrogen bond with the CH3 substitution is more important in the 2′-O- position, leading to a significant reduction in substrate specificity. These results are the first evidence for the fact that 2′-O- and 3′-O- methylations of m7Guo may influence cap-binding proteins action in a different way. Our new finding could be a good starting point for elucidation of detailed mechanism of action on molecular level, studying inhibition and designing effective inhibitors (especially that human DcpS was selected as a therapeutic target for Spinal Muscular Atrophy treatment ). Moreover, the differences between the hydrolytic activity of m27,2′-OGpppG and m27,3′-OGpppG may be a crucial feature for their biotechnological application.
The crucial role of the region associated with binding the ribose moiety arises also from a sequence alignment of different DcpS proteins (Fig. 4). Amino acids interacting with m7Guo in human DcpS (Asn110, Trp175, Glu185, Asp205, Lys207) are highly conserved in the illustrated organisms. Mutations of these crucial amino acids resulted in enzyme inactivation or significant decrease of its activity . Two of them, Asp205 and Lys207, are involved in interactions with 2′-O- and 3′-O- positions of ribose moiety of m7Guo in the human protein.
DcpS orthologs reported in different species (human, yeast and nematode cells) share significant sequence similarity (Fig. 4), however they differ in their ability to hydrolyze different cap structures. Yeast and human scavengers recognize only monomethylated cap analogues as substrates, whereas C. elegans DcpS is capable for efficient cleavage of both MMG and TMG caps. Kinetic data for enzymatic hydrolysis of m7GpppG catalyzed by S. cerevisiae Dcs1 (Km = 0.14 μM)  and C. elegans DcpS (Km = 1.3 μM) (Table 1) illustrate their high specificity for monomethylguanosine cap. From such low Km values one can conclude that DcpS enzymes are capable of maintaining high specific hydrolytic activity down to sub-micromolar intracellular concentrations of capped dinucleotides and short mRNA fragments. It therefore seems to be appropriately adopted to clear various capped species from the cells.
Despite their well-known decapping function in cytoplasmic mRNA turnover, yeast and human scavengers have been detected predominantly in the nucleus . This may suggest that yeast and mammalian DcpS are involved primarily in nuclear degradation of the cap structure. Their high specificity for MMG cap is crucial for quick removal of methylated nucleotides from the nucleus, preventing their misincorporation into RNA chain during transcription . In contrast, nematode DcpS is predominantly cytoplasmic protein . Although some regions of more intense DcpS labeling have been observed, DcpS scavengers are not components of specific degradation foci, processing bodies. The fact that C. elegans mRNAs are in the majority (~70%) trimethylated may explain the most of detectable DcpS protein observed in the cytoplasm  and a higher hydrolytic activity towards TMG cap determined in this study (Table 1). Dual activity of C. elegans DcpS is required for efficient degradation of mono- and timethylated species which may interact with eIF4E proteins during translation.
The ability of DcpS proteins to compete with eIF4E for the cap structure supports the idea that scavenger decapping enzymes may play modulatory roles at different levels of mRNA metabolism (cap-dependent translation, miRNA guided translation repression, 5′→3′ degradation). Recently, it has been demonstrated that human DcpS is a nucleocytoplasmic shuttling protein with a broad functionality as a modulator of cap-dependent processes . It was also suggested that decapping activity in C. elegans and S. cerevisiae is required for responses to heat shock and genotoxic stress [25, 31].
Kinetic studies presented in this paper provide insight into the mechanism of interaction of MMG and TMG cap with the binding pocket of C. elegans DcpS. Detailed characteristic of DcpS scavenger presented in this study is essential to understand a key step in mRNA turnover and may enable the design and synthesis new cap analogues that are selective inhibitors for parasitic nematodes scavenger decapping enzymes, without affecting their mammalian counterparts.
Recombinant C. elegans DcpS in pET16b  was grown in E. coli Rosetta (DE3) cells (Novagen, Madison, WT) at 37 °C until it reached OD600 = 0.5. Protein production was induced by adding 0.4 mM isopropyl β-D- thiogalactopyranoside (IPTG) and by shaking the bacterial culture for 16 h at 20 °C. The culture was centrifugated and bacterial pellets resuspended in ice-cold lysis buffer (20 mM HEPES, pH 7.5, 300 mM NaCl, 30 mM urea, 10% glycerol, 1% Triton X-100, 10 mM imidazole), lysozyme was added to a final concentration of 1 mg/ml, the suspension incubated on ice for 30 min, and then sonicated on ice (15 × 30 s every 1 min). The 6x-His-tagged DcpS was bound to Ni2+- nitrilotriacetic acid (NTA)-agarose (Novagen, Madison, WT) for 60 min at 4 °C and then, unbound proteins were removed with a washing buffer (20 mM Tris HCl, pH 7.5, 300 mM NaCl). The bound protein was eluted by 2 ml portions of elution buffer (20 mM Tris HCl, pH 7.5, 300 mM NaCl) containing increasing concentration of imidazole (from 20 to 300 mM). Fractions containing DcpS activity were dialyzed against 20 mM Tris HCl, pH 7.2, 50 mM KCl, 0.2 mM EDTA, 20% glycerol, 1 mM DTT, and stored at −80 °C. The enzyme activity was checked before each set of experiments. The concentration of DcpS was estimated by the method of Bradford  and spectrophotometrically from its molar absorption coefficient ε280 = 38900 M−1cm−1 (calculated from amino acid composition of a monomer using an algorithm on the ExPASy Server).
Cap analogues investigated in this work (m7GpppG, m32,2,7GpppG, m7GpppA, m32,2,7GpppA, m27,2′-OGpppG, m27,3′-OGpppG, bn7GpppG, et7GpppG, m7Gpppm7G, m7Gppp2′dG, m7Gpppm2′-OG, m7Gpppm6A) were prepared according to the methods described earlier [27,28,33–36].
Dinucleotide cap analogues and their hydrolysis products were identified using absorption and emission spectroscopy and HPLC analysis. The concentrations of investigated substrates were determined on the basis of their absorption coefficients: ε255(m7GpppG) = 22600 M−1cm−1, ε259(m7GpppA) = 21300 M−1cm−1, ε262(m7Gpppm6A) = 21100 M−1cm−1, ε259(m7Gpppm7G) = 16000 M−1cm−1, ε255(m7Gpppm2′-OG) = 19600 M−1cm−1, ε255(m7Gppp2′dG) = 19300 M−1cm−1 , ε255(m27,2′-OGpppG) = 20800 M−1cm−1, ε255(m27,3′-OGpppG) = 22000 M−1cm−1 (J. Zuberek, unpublished data), ε255(et7GpppG) = 21900 M−1cm−1, ε256(bn7GpppG) = 17800 M−1cm−1, ε258(m32,2,7GpppG = M−1cm−1 . The coefficient for m32,2,7GpppA ε260 = 28900 M−1cm−1 was calculated in this study. Absorption spectra were recorded in 0.1 M phosphate buffer, pH 7.0, on a Lambda 20UV/VIS spectrophotometer (Perkin Elmer Co.) at 20 °C.
Hydrolytic activity of the recombinant C. elegans DcpS was assayed at 20 °C in 50 mM Tris buffer containing 20 mM MgCl2, 30 mM (NH4)2SO4 (final pH 7.2). Scavenger decapping enzymes have been reported to share a neutral pH range (7 – 8) as the optimum reaction medium for their activity [14,25,26]. We previously demonstrated that the kinetic parameters of enzymatic hydrolysis catalyzed by C. elegans DcpS do not change significantly in this pH range . However, the fluorescence intensity and stacking interactions of dinucleotide cap analogues are strongly pH-dependent. The cationic (N1 protonated) form of the 7-alkylated residue exhibits higher fluorescence quantum yield and more efficient stacking than its zwitterionic counterpart [38–40]. A lower pH is thus more favourable to observe the fluorescence increase during the cleavage of the pyrophosphate bridge. Consequently, pH 7.2 was adopted for the enzymatic hydrolysis assays monitored by the fluorimetric method, as well as for the HPLC measurements.
The initial substrate concentration ranged from 0.5 μM to 120 μM, depending on the analyzed compound. DcpS cleavage assays were carried out with 0.11 – 1.98 μg of recombinant protein. The products of enzymatic hydrolysis were examined by analytical HPLC (Agilent Technologies 1200 Series) using a reverse-phase Supelcosil LC-18-T column (4.6 mm × 250 mm, 5 μm), UV/VIS and fluorescence detector. After a sample injection the column was eluted at room temperature with a linear gradient of methanol from 0% to 25% in aqueous 0.1 M KH2PO4 over 15 min at a flow rate of 1.3 ml/min. The fluorescence at 337 nm (excitation at 280 nm) and absorbance at 260 nm were continuously monitored during the analysis.
For all investigated dinucleotides, the spectrofluorimetric method was used to determine the kinetic parameters. The fluorescence measurements were performed on a LS 55 spectrofluorometer (Perkin Elmer Co.) in a quartz cuvette (Hellma) with optical path length of 4 mm for absorption and 10 mm for emission. Fluorescence intensity was observed at 380 nm (excitation at 294 – 318 nm, depending on the cap analogue) and corrected for the inner filter effect. Hydrolysis was followed over 10 minutes by recording the time-dependent increase of fluorescence intensity, caused by removal of intramolecular stacking as a result of enzymatic cleavage of triphosphate bridge. The substrate concentration (c) at the time of hydrolysis (t) was calculated as:
where co is the initial concentration of the substrate, It, Io, Ie are fluorescence intensities at the time (t), at the beginning and at the end of the reaction, respectively. The initial velocity (vo) of each reaction was calculated by the linear regression of substrate concentration versus time.
In order to confirm fluorimetric data, the kinetic parameters for m32,2,7GpppG, m27,2′-OGpppG and m27,3′-OGpppG were also obtained by means of HPLC. Other cap analogues could not be studied using chromatographic analysis, because the sensitivity of the HPLC system was not adequate to detect very low substrate concentration (0.2 - 10 μM) necessary to determine Km values ~1 μM. HPLC analysis is more effective for kinetic studies of compounds characterized by higher Km values (> 10 μM). In HPLC procedure, buffer solutions containing respective dinucleotides were incubated at 20 °C for 10 min. The hydrolysis process was started by the addition of DcpS. At 3 or 5 min time intervals 150 μl aliquots of the reaction mixture were withdrawn and the reaction terminated by heat inactivation of the enzyme (2.5 min at 100 °C). The samples were then subjected to HPLC analysis as described above. The concentration of examined compounds during the course of hydrolysis was determined from the area under the chromatographic peaks, using the following formula:
where c is the substrate concentration at the time of hydrolysis (t), co is the initial substrate concentration, x is the extent of decapping measured as the % of hydrolyzed substrate.
The initial velocity method was used to calculate the kinetic parameters for both fluorimetric and HPLC method. The initial velocity (vo) of each reaction was calculated by the linear regression of substrate concentration versus time. The Km and Vmax values were determined from hyperbolic fits to the Michaelis-Menten equation by non-linear regression using OriginPro 7.0 (Microcal Software).
This work was supported by the National Science Support Project 2008-1010 No. PBZ-MNiSW-07/I/2007 and NIH Grant AI049558 to R.E.D. E. Darzynkiewicz is a Howard Hughes Medical Institute International Scholar (Grant No.55005604).