Structural requirements for Caenorhabditis elegans DcpS substrates based on fluorescence and HPLC enzyme kinetic studies

Article abstract of DOI:10.1111/j.1742-4658.2010.07709.x

The activity of the Caenorhabditis elegans scavenger decapping enzyme (DcpS) on its natural substrates and dinucleotide cap analogs, modified with regard to the nucleoside 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 (K_m, V_max) were determined using fluorescence and HPLC methods, as complementary approaches for the kinetic studies of C. elegans DcpS. From the kinetic data, we determined which parts of the cap structure are crucial for DcpS binding and hydrolysis. We showed that m₃ ^2,2,7GpppG and m₃^2,2,7GpppA are cleaved with higher rates than their monomethylated counterparts. However, the higher specificity of C. elegans DcpS for monomethylguanosine caps is illustrated by the lower K_m values. Modifications of the first transcribed nucleotide did not affect the activity, regardless of the type of purine base. Our findings suggest C. elegans DcpS flexibility in the first transcribed nucleoside-binding pocket. Moreover, although C. elegans DcpS accommodates bulkier groups in the N7 position (ethyl or benzyl) of the cap, both 2′-O- and 3′-O-methylations of 7-methylguanosine result in a reduction in hydrolysis by two orders of magnitude.

Full text of DOI:10.1111/j.1742-4658.2010.07709.x

Structural requirements for Caenorhabditis elegans DcpS
substrates based on fluorescence and HPLC enzyme
kinetic studies
Anna Wypijewska1, Elzbieta Bojarska1, Janusz Stepinski1, Marzena Jankowska-Anyszka2,
Jacek Jemielity1, Richard E. Davis3 and Edward Darzynkiewicz1
1 Division of Biophysics, Institute of Experimental Physics, Faculty of Physics, University of Warsaw, Poland
2 Department of Chemistry, University of Warsaw, Poland
3 Department of Biochemistry and Molecular Genetics, University of Colorado, School of Medicine, Aurora, CO, USA
Keywords
The activity of the Caenorhabditis elegans scavenger decapping enzyme
(DcpS) on its natural substrates and dinucleotide cap analogs, modified with
regard to the nucleoside base or ribose moiety, has been examined. All
enzyme kinetics; fluorescence
spectroscopy; mRNA cap analogs; mRNA
degradation; scavenger decapping enzymes
tested dinucleotides were specifically cleaved between b- and c-phosphate
groups in the triphosphate chain. The kinetic parameters of enzymatic
Correspondence
hydrolysis (Km, Vmax) were determined using fluorescence and HPLC meth-
ods, as complementary approaches for the kinetic studies of C. elegans
DcpS. From the kinetic data, we determined which parts of the cap struc-
ture are crucial for DcpS binding and hydrolysis. We showed that
m32,2,7GpppG and m32,2,7GpppA are cleaved with higher rates than their
monomethylated counterparts. However, the higher specificity of C. elegans
DcpS for monomethylguanosine caps is illustrated by the lower Km values.
Modifications of the first transcribed nucleotide did not affect the activity,
regardless of the type of purine base. Our findings suggest C. elegans DcpS
flexibility in the first transcribed nucleoside-binding pocket. Moreover,
although C. elegans DcpS accommodates bulkier groups in the N7 position
(ethyl or benzyl) of the cap, both 2¢-O- and 3¢-O-methylations of 7-methyl-
guanosine result in a reduction in hydrolysis by two orders of magnitude.
E. Bojarska, Division of Biophysics, Institute
of Experimental Physics, Faculty of Physics,
University of Warsaw, 93 Zwirki & Wigury
Ave., 02-089 Warsaw, Poland
Fax: +48 22 554 0771
Tel: +48 22 554 0779
E-mail: elab@biogeo.uw.edu.pl
(Received 25 February 2010, revised 8 May
2010, accepted 12 May 2010)
doi:10.1111/j.1742-4658.2010.07709.x
Introduction
mRNA turnover is a critical determinant in the regula-
tion of gene expression [1–3]. The degradation of nor-
mal transcripts in eukaryotes occurs along two major
pathways, 5¢ fi 3¢ and 3¢ fi 5¢ decay, both initiated
by shortening of the poly(A) tail [4,5]. In the 5¢ fi 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¢ fi 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¢ fi 5¢ until it reaches the cap-containing
dinucleotide or a short capped oligonucleotide [8,9].
The residual cap structure m7GpppN (7-meth-
ylGpppN) is further hydrolyzed by the scavenger
decapping enzyme (DcpS) [10]. Capped dinucleotides
or oligonucleotides accumulated in cells could bind to
cap-binding proteins, such as eIF4E, and inhibit trans-
lation [11]. The hydrolysis of cap dinucleotides in
this context is thought to be important. However,
Abbreviations
ARCA (anti-reverse cap analog), m27,2¢-OGpppG and m27,3¢-OGpppG; bn7GpppG, 7-benzylGpppG; BODIPY, 4,4-difluoro-4-bora-3a,4a-diaza-
s-indacene; DcpS, scavenger decapping enzyme; et7GpppG, 7-ethylGpppG; HIT, histidine triad; m32,2,7GpppG, trimethylguanosine cap;
m7GpppN, 7-methylGpppN; m7Guo, 7-methylguanosine; MMG and TMG cap, monomethylguanosine and trimethylguanosine cap.
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A. Wypijewska et al.
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 histidine triad (HIT) family of pyrophos-
phatases, with decapping activity as the main, well-
defined biological function [16,17]. All of these
enzymes exhibit high specificity for cap structure and
limited activity towards nonmethylated dinucleotides
(e.g. ApppA and GpppG). Decapping scavengers uti-
lize an evolutionary conserved HIT motif 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 HIT with three
histidines separated by hydrophobic residues (His-u-
His-u-His-u). Structural analysis has revealed that
HIT proteins exist as homodimers containing nucleo-
tide-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 differ-
ent scavengers supports the functional significance of
this domain in decapping activity. Substitution muta-
genesis of the central histidine in human and nematode
decapping scavengers inactivates their hydrolytic prop-
erties, demonstrating that the central HIT motif is
critical for catalysis [14,20]. This histidine is involved
in the formation of a covalent nucleotidyl phosphohist-
idyl intermediate, the nucleophilic agent for the
c-phosphate group of dinucleoside triphosphate sub-
strates [19,20].
oligonucleotides up to four nucleotides were hydro-
lyzed [14].
Previous data have suggested that the substrate spec-
ificity of C. elegans DcpS differs from that of its
human and yeast orthologs [3,14,25,26]. However, nei-
ther detailed kinetic analysis of enzymatic cleavage nor
mechanisms of substrate recognition have been investi-
gated on C. elegans DcpS. In this article, we have
studied the substrate specificity and kinetic analysis of
recombinant C. elegans DcpS. Various dinucleotide
cap analogs, natural and chemically modified within
the 7-methylgunosine moiety or the first transcribed
nucleoside, have been investigated as potential sub-
strates. Kinetic parameters (Km, Vmax and Vmax ⁄ Km)
were determined to characterize the hydrolytic activity
of C. elegans DcpS.
Results
Decapping products of reactions catalyzed by
C. elegans DcpS
To identify the DcpS hydrolysis products of all investi-
gated dinucleotides presented in Fig. 1, high-perfor-
mance liquid chromatograms were analyzed. As an
example, chromatographic analysis for the cleavage of
monomethylguanosine (MMG) cap, trimethylguano-
sine (TMG) cap and GpppG are shown in Fig. 2. For
m32,2,7GpppG, the peak corresponding to the substrate
disappeared after 10 min of reaction (Fig. 2A). MMG
was almost completely hydrolyzed over 20 min
(Fig. 2B). The hydrolysis of GpppG was much slower
– after 120 min a considerable amount of the substrate
was still observed in the reaction mixture (Fig. 2C).
The analysis of the hydrolysis products (Table 1)
demonstrates that the cleavage of cap analogs occurs
exclusively between b- and c-phosphate groups within
the triphosphate bridge. These data confirm the earlier
observations that nematode DcpS utilizes the same
mechanism of catalysis as proposed for other HIT
pyrophosphatases cleaving the cap structure, and the
highly conserved HIT motif is involved in the binding
of the substrates and catalysis [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. Approxi-
mately 70% of nematode mRNAs possess a trimethyl-
guanosine cap (m32,2,7GpppG), whereas approximately
30% have a typical cap structure (m7GpppG) [21].
Both types of mRNA interact with polysomes and
undergo translation [12,22]. The presence of two popu-
lations of mRNAs has profound implications for pro-
teins that recognize specifically each mRNA [23]. The
eIF4E protein in C. elegans exists in five different iso-
forms, 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
Specificity of C. elegans DcpS towards MMG and
TMG caps
Initial studies on the substrate specificity of recombi-
nant C. elegans DcpS suggested that the protein was
specific for 7-methylguanosine (m7Guo) nucleotides.
The first quantitative experiments characterizing this
enzyme were reported by Kwasnicka et al. [25]. How-
ever, the specificity of C. elegans DcpS was defined
3004
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A. Wypijewska et al.
Caenorhabditis elegans DcpS kinetic studies
O
R2
OR5
OH
–
+
N
N
O
P
O
P
O
P
O
B
R1
O
O
O
O
N
N
O
–
–
–
O
O
O
OR4 OR3
—————————————————————————————————————
Cap
Reference
analogue
to synthesis
Structure
—————————————————————————————————————
m7GpppG
m32,2,7GpppG
m7GpppA
m32,2,7GpppA
m27,2’-OGpppG
m27,3’-OGpppG
bn7GpppG
33
33
33
33
28
27
38
38
34
R1 = NH2, R2 = CH3, R3 = R4 = H, R5 = OH, B = guanine
R1 = N(CH3)2, R2 = CH3, R3 = R4 = H, R5 = OH, B = guanine
R1 = NH2, R2 = CH3, R3 = R4 = H, R5 = OH, B = adenine
R1 = N(CH3)2, R2 = CH3, R3 = R4 = H, R5 = OH, B = adenine
R1 = NH2, R2 = CH3, R3 = CH3, R4 = H, R5 = OH, B = guanine
R1 = NH2, R2 = CH3, R3 = H, R4 = CH3, R5 = OH, B = guanine
R1 = NH2, R2 = CH2C6H5, R3 = R4 = H, R5 = OH, B = guanine
R1 = NH2, R2 = CH2CH3, R3 = R4 = H, R5 = OH, B = guanine
R1 = NH2, R2 = CH3, R3 = R4 = H, R5 = OH, B = 7-methyl-
guanine
et7GpppG
m7Gpppm7G
m7Gppp2’dG
35
35
35
R1 = NH2, R2 = CH3, R3 = R4 = R5 = H, B = guanine
R1 = NH2, R2 = CH3, R3 = R4 = H, R5 = OCH3, B = guanine
R1 = NH2, R2 = CH3, R3 = R4 = H, R5 = OH, B = N6-methyl-
adenine
m7Gpppm2’-O
m7Gpppm6A
G
—————————————————————————————————————
Fig. 1. Structures of the investigated cap analogs and references to their synthesis.
with m7GpppBODIPY, GpppBODIPY and ApppBO-
DIPY (BODIPY, 4,4-difluoro-4-bora-3a,4a-diaza-s-
indacene), but not with natural caps m7GpppG or
m32,2,7GpppG. Methylated mono- and dinucleotides
(m7GDP, m7GTP, m7GpppG, m32,2,7GpppG) have
only been examined as inhibitors of C. elegans scaven-
ger in the hydrolysis process of m7GpppBODIPY. The
inhibition constant calculated for m32,2,7GpppG
make a detailed comparison of C. elegans DcpS activ-
ity 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 caps were hyperbolic in the investi-
gated concentration ranges: 0.5–86 lm for m7GpppG
and 0.5–97 lm for m32,2,7GpppG. The kinetic parame-
ters 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 three times
higher for the TMG cap than for the MMG cap,
whereas the Vmax ⁄ Km values are almost the same. This
indicates that C. elegans DcpS has slightly different
substrate specificities for these natural compounds,
(Ki = 28.1
m7GpppG (Ki = 3.47
2.5 lm), eight-fold higher than for
0.84 lm), indicated less effi-
cient inhibitory properties of the trimethylated cap in
comparison with its monomethylated counterpart. On
the basis of these findings, it was concluded that the
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 reaction with the
substrates were not determined quantitatively [14]. To
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A. Wypijewska et al.
A
B
C
Fig. 2. HPLC profiles for the hydrolysis of m32,2,7GpppG (A), m7GpppG (B) and GpppG (C) catalyzed by Caenorhabditis elegans DcpS. The ini-
tial concentration of each substrate was 10 lM and the reactions were carried out with the same amount of enzyme: 1 lg. Absorbance was
measured at 260 nm (AU, arbitrary units). The chromatographic peaks were identified by comparison with the retention times of reference
samples.
with a preference for m7GpppG, as suggested previ-
ously [25,26]. However, the rate of hydrolysis catalyzed
by C. elegans DcpS is higher for the TMG cap.
values for these four compounds are similar to that
obtained for the 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 the catalytic mechanism of C. elegans
DcpS.
Kinetics of cap analogs modified in the first
transcribed nucleoside
To further examine the substrate specificity of C. ele-
gans DcpS, the hydrolysis of several other dinucleotide
cap analogs was examined. Substitution of adenine for
guanine as the second nucleotide in MMG and TMG
caps did not change significantly the substrate proper-
ties of m7GpppA and m32,2,7GpppA for DcpS catalysis
when compared with 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,
Kinetics of cap analogs modified in m7Guo
The next interesting part of our studies concerning the
substrate requirements for C. elegans DcpS revealed
that the enzyme tolerates differently sized substituents
at the N7 position of m7Guo. The kinetic data (Km,
Vmax and Vmax ⁄ Km) calculated for m7GpppG (7-methyl
GpppG), et7GpppG (7-ethylGpppG) and bn7GpppG
(7-benzylGpppG) clearly showed that all three com-
pounds are similarly recognized as substrates by the
nematode scavenger (Table 1). These findings suggest
m
2¢-OG) were all good DcpS substrates, as illustrated by
the kinetic data (Fig. 3, Table 1). The Km and Vmax
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A. Wypijewska et al.
Caenorhabditis elegans DcpS kinetic studies
Table 1. Comparison of the substrate specificity of cap analogs towards Caenorhabditis elegans DcpS, obtained by the initial velocity
method at 20 ꢀC in 50 mM Tris ⁄ HCl buffer containing 30 mM (NH4)2SO4 and 20 mM MgCl2 (pH 7.2).
Cap analog
Products of hydrolysis
Km (lM)
Vmax (UÆmg)1
)
Vmax ⁄ Km (min)1Æmg)1
)
Fluorescence method
m7GpppG
m7GpppA
m7Gpppm6A
m7Gpppm7G
m7GMP + GDP
m7GMP + ADP
1.17 0.14
0.60 0.11
1.03 0.16
1.12 0.14
1.23 0.13
1.36 0.41
42.13 3.91
15.39 2.08
0.61 0.18
1.83 0.15
3.85 0.41
2.36 0.16
1.53 0.11
1.09 0.11
1.33 0.12
0.91 0.10
1.66 0.12
2.00 0.26
3.28 0.19
0.51 0.11
3.12 1.45
3.06 0.12
4.65 0.26
2.06 0.10
1.30 0.18
1.83 0.38
1.30 0.23
0.81 0.13
1.35 0.17
1.47 0.48
0.08 0.01
0.03 0.01
5.09 2.80
1.67 0.15
1.21 0.15
0.87 0.07
m7GMP + m6ADP
m7GMP + m7GDP
m7GMP + m2¢-OGDP
m7GMP + 2¢dGDP
m27,2¢-OGMP + GDP
m27,3¢-OGMP + GDP
et7GMP + GDP
bn7GMP + GDP
m32,2,7GMP + GDP
m32,2,7GMP + ADP
m7Gpppm2¢-O
G
m7Gppp2¢dG
m27,2¢-OGpppG
m27,3¢-OGpppG
et7GpppG
bn7GpppG
m32,2,7GpppG
m32,2,7GpppA
HPLC method
m27,2¢-OGpppG
m27,3¢-OGpppG
m32,2,7GpppG
m27,2¢-OGMP + GDP
m27,3¢-OGMP + GDP
m32,2,7GMP + GDP
39.77 3.07
13.87 0.48
3.91 0.82
2.45 0.11
0.31 0.10
3.14 0.32
0.06 0.01
0.02 0.01
0.80 0.19
plasticity within the C. elegans DcpS cap-binding
pocket.
found that cleavage of the cap structure occurs exclu-
sively between b- and c-phosphate groups in the
triphosphate chain. We examined the ability of the
enzyme to act on various cap analogs in a quantitative
manner, employing two independent methods (fluores-
cence and HPLC) to determine the kinetic data.
We also examined m27,2¢-OGpppG and m27,3¢-OGpppG
(bearing additional methylation at the 2¢ or 3¢ oxygen
of m7Guo) as C. elegans DcpS substrates (Fig. 3). Km
values determined by the fluorimetric and HPLC meth-
ods for both compounds are significantly higher than
7,3¢-
for m7GpppG (Table 1). Furthermore, for m2
Monomethylated and trimethylated natural
substrates
OGpppG, the rate of hydrolysis is drastically reduced.
This compound has been studied previously as an
effective inhibitor of m32,2,7GpppA hydrolysis cata-
lyzed by C. elegans DcpS, with Ki = 1 lm [26], signifi-
cantly lower than the Km value (ꢀ 14 lm) determined
in this study (Table 1). Such a low Ki value indicates
tight binding of m27,3¢-OGpppG with DcpS, whereas
Km involves a contribution from the 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 [26].
The kinetic parameters obtained for m27,2¢-OGpppG
and m27,3¢-OGpppG indicate that the 2¢-OH and 3¢-OH
positions in the ribose ring of the m7Guo moiety play
a significant role in the catalytic activity of C. elegans
DcpS.
Among the different scavengers investigated (human,
nematode, yeast), C. elegans DcpS has a unique prop-
erty, i.e. the possibility to hydrolyze both monomethy-
lated (m7GpppG and m7GpppA) and trimethylated
(m32,2,7GpppG and m32,2,7GpppA) cap structures. Our
kinetic data demonstrate that trimethylated caps are
cleaved with higher rates than their monomethylated
counterparts (Table 1). However, MMG caps are
recognized with higher specificity, indicating that the
two additional methyl groups at the N2 position in
TMG caps account for the differences in Km for these
substrates.
Substrates with an alkyl group at the N7 position
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 the N7
position, introduced by a methyl or any alkyl group.
Discussion
A series of modified dinucleotide cap analogs studied
in this work defined several structural requirements for
substrate specificity towards C. elegans DcpS. We
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A. Wypijewska et al.
m7 GpppG
m7 Gpppm6A
m32,2,7 GpppG
different cap structures. This flexibility may explain
why significantly large groups, such as ethyl or benzyl,
can interact with nematode scavenger and be hydro-
lyzed with comparable rates.
A 6
5
4
3
2
1
0
Substrates modified in the first transcribed
nucleoside
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
analogs modified in the first transcribed nucleoside.
We established that modifications introduced into the
first transcribed nucleoside do not influence signifi-
cantly nematode DcpS kinetic parameters. The substi-
tution of adenine for guanine in m7GpppG or
m32,2,7GpppG does not affect the Km values. Other cap
analogs bearing modifications of Guo, such as m6A,
m7G, m2¢-OG and 2¢dG, have similar kinetic parame-
ters to m7GpppG, indicating that modifications of the
base or ribose moiety within the first transcribed nucle-
otide are not crucial for substrate recognition or the
rate of hydrolysis. Moreover, the Km value for
0
10 20 30 40 50 60 70 80 90 100 110 120
co [µM]
3
2
1
0
B
m7 GpppG
m7,2'–O GpppG
m7,3'–O GpppG
m7GpppG (1.17
the Km value reported for m7GpppBODIPY
(1.21 0.05 lm), containing an artificial fluorescent
0.14 lm) is remarkably similar to
probe BODIPY instead of guanine [25]. Caenorhabd-
itis elegans DcpS thus can accept different, even nonbi-
ological substituents instead of the first transcribed
nucleotide, which do not affect the substrate specificity
or hydrolysis rate.
0
10 20 30 40 50 60 70 80 90 100 110 120
co [µM]
Fig. 3. Caenorhabditis elegans DcpS hydrolysis kinetics with cap
analogs. (A) Comparison of the kinetic curves of C. elegans DcpS
natural substrates (m7GpppG, m32,2,7GpppG) and a cap analog with
a modification in the first transcribed nucleoside (m7Gpppm6A). (B)
Comparison of the kinetic curves of m7GpppG and anti-reverse cap
analogs (m27,2¢-OGpppG and m27,3¢-OGpppG). The initial velocity data
vo(co) were obtained from fluorescence studies.
A similar effect was observed for human DcpS.
Mutagenesis of the human DcpS amino acids responsi-
ble for the contacts with the first transcribed nucleoside
had little effect on enzyme activity, suggesting that the
structure of the binding pocket recognizing the first
transcribed nucleoside is more flexible than that of the
cap-binding pocket [20]. As shown in Fig. 4, the amino
acids recognizing the first transcribed nucleoside are
not conserved in DcpS homologs, indicating that inter-
action with this nucleoside is not very important for
decapping activity. We thus propose that DcpS pro-
teins exhibit structural plasticity for the first transcribed
nucleoside, which has no affect on enzyme hydrolysis.
Differently sized substituents (methyl, ethyl, benzyl)
introduce positive charge into the base moiety, which
is a key feature for the recognition of the cap struc-
ture. The amino acids involved in the stacking interac-
tions with the methylated base are not conserved in
different organisms (Fig. 4), and thus apparently are
not crucial for hydrolytic activity, as indicated by the
mutation L206A retaining over 90% of the wild-type
activity of human DcpS [20]. The substrate properties
of N7 alkylated dinucleotides (m7GpppG, et7GpppG,
bn7GpppG) do not differ significantly, as indicated by
the 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
Substrates modified by additional methylation at
the 2¢ or 3¢ oxygen of m7Guo
The kinetic parameters obtained for m27,2¢-OGpppG
and m27,3¢-OGpppG demonstrated the crucial role of
the 2¢-OH and 3¢-OH groups of the m7Guo moiety for
C. elegans DcpS hydrolysis. The 2¢-O-Me and 3¢-O-Me
3008
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A. Wypijewska et al.
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A
B
Fig. 4. Multiple sequence alignment of DcpS from different organisms generated using the CLUSTAL 2.0.12 program. The nematodes (Ancy-
lostoma duodenale, Ascaris suum, Brugia malayi, Heterodera glycines, Meloidogyne hapla, Caenorhabditis briggsae, Caenorhabditis elegans)
are framed. All the nematodes and the first three organisms (Schistosoma japonicum, Ciona intestinalis, Hydra magnipapillata) show trans-
splicing, suggesting that they would probably be able to hydrolyze the TMG cap. The remaining orthologs are from Homo sapiens,
Sus scrofa, Mus musculus, Drosophila melanogaster and Saccharomyces cerevisiae. The amino acids of each organism are numbered on
the right. Human DcpS (hDcpS) amino acids making vicinal or van der Waals’ contacts with m7GpppG are marked by arrows. The parts of
m7GpppG involved in these interactions and the percentage of m7GpppG hydrolysis catalyzed by hDcpS with mutation of these amino acids
to Ala are given above (n.d., not determined) [20]. Among the indicated amino acids, those identical to those in C. elegans DcpS are boxed
in black. (A) Alignment of the amino acids involved in the interactions with the first transcribed nucleoside (Guo) in the hDcpS–m7GpppG
complex. These amino acids are not conserved in the other DcpS proteins illustrated. Mutation of the indicated amino acids in hDcpS to Ala
only decreases slightly the enzymatic activity of the human scavenger [20]. (B) Alignment of the amino acids involved in the interactions with
the cap structure (m7Guo) in the hDcpS–m7GpppG complex. The majority of these amino acids are highly conserved within the presented
organisms. Mutations of these amino acids in human DcpS significantly or even completely inactivate the human enzyme [20].
analogs are so-called ARCA (anti-reverse cap analogs)
which are commercially available and used as sub-
strates for in vitro transcription reactions [27,28]. Such
analogs prevent their reverse incorporation into
mRNAs, thus producing 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
FEBS Journal 277 (2010) 3003–3013 ª 2010 The Authors Journal compilation ª 2010 FEBS
3009

Caenorhabditis elegans DcpS kinetic studies
A. Wypijewska et al.
mimic well natural transcripts, e.g. in the initiation of
translation (the methylation of ribose of m7Guo does
not disturb the interaction with eIF4E) [28]. We estab-
lished that, in some studies, ARCA-prepared tran-
scripts may not be a good mimic of natural transcripts
(this DcpS study is a good example). As indicated by
the high Km values and very low Vmax ⁄ Km values, both
of these compounds are poor substrates for C. elegans
DcpS. Interestingly, 2¢-O- and 3¢-O-methylations pro-
duce various susceptibilities of the cap to enzymatic
hydrolysis. Despite the fact that the efficiencies of
hydrolysis are reduced by two orders of magnitude
compared with the natural substrates, the kinetic
parameters (Km and Vmax) are significantly different.
Although the leaving group is the same as in the
MMG cap (GDP), the rate of hydrolysis observed for
m27,3¢-OGpppG is significantly lower, suggesting that
slow dissociation of the enzyme–product complex
might be a controlling 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 provide
the first evidence indicating that 2¢-O- and 3¢-O-methy-
lations of m7Guo may influence the action of cap-
binding proteins in a different manner. Our new
finding could be a good starting point for the elucidation
of the detailed mechanism of action on a molecular
level, for the study of inhibition and for the design of
effective inhibitors (in particular, human DcpS has
been selected as a therapeutic target for spinal muscu-
lar atrophy treatment [29]). Moreover, the differences
between the hydrolytic activities of m27,2¢-OGpppG and
m27,3¢-OGpppG may be crucial for their biotechnologi-
cal application.
scavengers recognize only monomethylated cap analogs
as substrates, whereas C. elegans DcpS is capable of
efficient cleavage of both MMG and TMG caps.
Kinetic data for the enzymatic hydrolysis of m7GpppG
catalyzed by S. cerevisiae Dcs1 (Km = 0.14 lm) [30]
and C. elegans DcpS (Km = 1.3 lm) (Table 1) illus-
trate their high specificity for the MMG cap. From
such low Km values, it can be concluded that DcpS
enzymes are capable of maintaining high specific hydro-
lytic activity down to submicromolar intracellular con-
centrations of capped dinucleotides and short mRNA
fragments. It therefore seems to be appropriately
adapted to clear various capped species from the cells.
Despite their well-known decapping function in
cytoplasmic mRNA turnover, yeast and human scav-
engers have been detected predominantly in the
nucleus [13]. This may suggest that yeast and mamma-
lian DcpSs are involved primarily in the nuclear degra-
dation of the cap structure. Their high specificity for
the MMG cap is crucial for the rapid removal of
methylated nucleotides from the nucleus, preventing
their misincorporation into the RNA chain during
transcription [30]. In contrast, nematode DcpS is pre-
dominantly a cytoplasmic protein [15]. Although some
regions of more intense DcpS labeling have been
observed, DcpS scavengers are not components of spe-
cific degradation foci–processing bodies. The fact that
C. elegans mRNAs are, in the majority (ꢀ70%), trime-
thylated may explain why most of the detectable DcpS
protein is observed in the cytoplasm [15] and the
higher hydrolytic activity towards the TMG cap deter-
mined in this study (Table 1). Dual activity of
C. elegans DcpS is required for efficient degradation of
mono- and trimethylated species, which may interact
with eIF4E proteins during translation.
The crucial role of the region associated with the
binding of the ribose moiety also arises from a
sequence alignment of different DcpS proteins (Fig. 4).
The amino acids interacting with m7Guo in human
DcpS (Asn110, Trp175, Glu185, Asp205, Lys207) are
highly conserved in the illustrated organisms. Muta-
tions of these crucial amino acids resulted in enzyme
inactivation or a significant decrease in activity [20].
Two amino acids, Asp205 and Lys207, are involved in
interactions with the 2¢-O- and 3¢-O-positions of the
ribose moiety of m7Guo in the human protein.
The ability of DcpS proteins to compete with eIF4E
for the cap structure supports the idea that DcpSs may
play modulatory roles at different levels of mRNA
metabolism (cap-dependent translation, miRNA-
guided translation repression, 5¢ fi 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 pro-
cesses [30]. It has also been suggested that decapping
activity in C. elegans and S. cerevisiae is required for
responses to heat shock and genotoxic stress [25,31].
The kinetic studies presented in this article provide
insight into the mechanism of interaction of MMG
and TMG caps with the binding pocket of C. elegans
DcpS. The detailed characteristics of the DcpS scaven-
ger presented in this study are essential to understand
the key step in mRNA turnover, and may enable the
design and synthesis of new cap analogs that are
Biological aspects
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
3010
FEBS Journal 277 (2010) 3003–3013 ª 2010 The Authors Journal compilation ª 2010 FEBS

A. Wypijewska et al.
Caenorhabditis elegans DcpS kinetic studies
of Biophysics, Institute of Experimental Physics, Faculty of
Physics, University of Warsaw, Poland, unpublished data);
selective inhibitors for parasitic nematode DcpSs, with-
out affecting their mammalian counterparts.
e
255(et7GpppG) = 21 900 m)1Æcm)1
17 800 m)1Æcm)1 258(m32,2,7GpppG) = 26 300 m)1Æcm)1
[36]. The coefficient for (e260
m32,2,7GpppA
;
e
256(bn7GpppG)
=
;
e
=
Materials and methods
28 900 m)1Æcm)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,
Waltham, MA, USA) at 20 ꢀC.
Materials
Recombinant C. elegans DcpS in pET16b [14] was grown
in Escherichia coli Rosetta (DE3) cells (Novagen, Madison,
WI, USA) at 37 ꢀC until an absorbance at 600 nm (A600) of
0.5 was reached. Protein production was induced by the
addition of 0.4 mm isopropyl thio-b-d-galactoside (IPTG)
and by shaking the bacterial culture for 16 h at 20 ꢀC. The
culture was centrifuged and the bacterial pellets were resus-
pended in ice-cold lysis buffer (20 mm Hepes, pH 7.5,
300 mm NaCl, 300 mm urea, 10% glycerol, 1% Triton
X-100, 10 mm imidazole); lysozyme was added to a final
concentration of 1 mgÆmL)1, the suspension was incubated
on ice for 30 min, and then sonicated on ice (15 · 30 s
every 1 min). The 6 · His-tagged DcpS was bound to
Ni2 + - nitrilotriacetic acid (NTA)-agarose (Novagen) for
60 min at 4 ꢀC, and unbound proteins were removed with
washing buffer (20 mm Tris ⁄ HCl, pH 7.5, 300 mm NaCl).
The bound protein was eluted with 2 mL portions of
elution buffer (20 mm Tris ⁄ HCl, pH 7.5, 300 mm NaCl)
containing increasing concentrations of imidazole (20–
300 mm). Fractions containing DcpS activity were dialyzed
against 20 mm Tris ⁄ HCl, pH 7.6, 50 mm KCl, 0.2 mm
EDTA, 20% glycerol and 1 mm dithiothreitol, 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 [32] and spectrophotometrically
The hydrolytic activity of the recombinant C. elegans
DcpS was assayed at 20 ꢀC in 50 mm Tris buffer containing
20 mm MgCl2 and 30 mm (NH4)2SO4 (final pH 7.2). DcpSs
have been reported to share a neutral pH range (pH 7–8) as
the optimum reaction medium for their activity [14,25,26].
We have demonstrated previously that the kinetic parame-
ters of enzymatic hydrolysis catalyzed by C. elegans DcpS
do not change significantly in this pH range [26]. However,
the fluorescence intensity and stacking interactions of
dinucleotide cap analogs are strongly dependent on pH.
The cationic (N1 protonated) form of the 7-alkylated resi-
due exhibits a higher fluorescence quantum yield and more
efficient stacking than its zwitterionic counterpart [38–40].
A lower pH is thus more favorable for the observation
of the fluorescence increase during the cleavage of the pyro-
phosphate bridge. Consequently, pH 7.2 was adopted for
the enzymatic hydrolysis assays monitored by the fluori-
metric method, as well as for the HPLC measurements.
The initial substrate concentration ranged from 0.5 to
120 lm, depending on the analyzed compound. DcpS cleav-
age assays were carried out with 0.11–1.98 lg of the recom-
binant protein. The products of enzymatic hydrolysis were
examined by analytical HPLC (Agilent Technologies 1200
Series, Santa Clara, CA, USA) using a reverse-phase Supe-
lcosil LC-18-T column (4.6 mm · 250 mm, 5 lm) and a
UV ⁄ VIS and fluorescence detector. After 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 mLÆmin)1. The
fluorescence at 337 nm (excitation at 280 nm) and absor-
bance at 260 nm were continuously monitored during the
analysis.
from
its
molar
absorption
coefficient
e280 =
38 900 m)1Æcm)1 (calculated from the amino acid composi-
tion of a monomer using an algorithm on the ExPASy
Server).
The cap analogs 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].
For all investigated dinucleotides, the spectrofluorimetric
method was used to determine the kinetic parameters. The
fluorescence measurements were performed on an LS 55
Analysis of hydrolysis kinetics
spectrofluorometer (Perkin-Elmer) in
a quartz cuvette
Dinucleotide cap analogs and their hydrolysis products
were identified using absorption and emission spectros-
copy and HPLC analysis. The concentrations of the investi-
gated substrates were determined on the basis of their
(Hellma, Mullheim, Germany) with an optical path length
of 4 mm for absorption and 10 mm for emission. The fluo-
rescence intensity was observed at 380 nm (excitation at
294–318 nm, depending on the cap analog) and corrected
for the inner filter effect. Hydrolysis was followed over
10 min by recording the time-dependent increase in fluores-
cence intensity caused by the removal of intramolecular
stacking as a result of enzymatic cleavage of the triphos-
phate bridge. The substrate concentration (c) at the time of
hydrolysis (t) was calculated as:
absorption coefficients: e255(m7GpppG) = 22 600 m)1Æcm)1
259(m7GpppA) = 21 300 m)1Æcm)1 262(m7Gpppm6A) =
21 100 m)1Æcm)1 259(m7Gpppm7G) = 16 000 m)1Æcm)1
255(m7Gpppm2¢-OG) = 19 600 m)1Æcm)1; e255(m7Gppp2¢dG) =
19 300 m)1Æcm)1 [37]; e255(m27,2¢-OGpppG) = 20 800 m)1Æcm)1
255(m27,3¢-OGpppG) = 22 000 m)1Æcm)1 (J. Zuberek, Division
;
e
;
e
;
e
;
e
;
e
FEBS Journal 277 (2010) 3003–3013 ª 2010 The Authors Journal compilation ª 2010 FEBS
3011

Caenorhabditis elegans DcpS kinetic studies
c ¼ coðItꢁIeÞ=ðIoꢁIeÞ
A. Wypijewska et al.
2 Cougot N, Babajko S & Seraphin B (2004) Cap-tabo-
lism. Trends Biochem Sci 29, 436–444.
where co is the initial concentration of the substrate, and It,
Io and Ie are the fluorescence intensities at time t, at the begin-
ning and at the end of the reaction, respectively. The initial
velocity (vo) of each reaction was calculated by the linear
regression of the substrate concentration versus time.
3 Parker R & Song H (2004) The enzymes and control of
eukaryotic mRNA turnover. Nat Struct Mol Biol 11,
121–127.
4 van Dijk E, Hir L & Seraphin B (2003) DcpS can act
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12086.
In order to confirm the fluorimetric data, the kinetic
7,3¢-O
parameters for m32,2,7GpppG, m27,2¢-OGpppG and m2
GpppG were also obtained by HPLC. Other cap analogs
could not be studied using chromatographic analysis,
because the sensitivity of the HPLC system was not ade-
quate to detect the very low substrate concentrations (0.2–
10 lm) necessary to determine Km values of ꢀ 1 lm. HPLC
analysis is more effective for kinetic studies of compounds
characterized by higher Km values (> 10 lm). In the HPLC
procedure, buffer solutions containing the respective dinu-
cleotides were incubated at 20 ꢀC for 10 min. The hydroly-
sis process was started by the addition of DcpS. At 3 or
5 min time intervals, 150 lL aliquots of the reaction mix-
ture were withdrawn and the reaction was 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 the examined compounds
during the course of hydrolysis was determined from the
area under the chromatographic peaks, using the following
formula:
5 Wilusz CJ, Gao M, Jones CL, Wilusz J & Peltz SW
(2001) Poly(A) binding protein regulates both deadeny-
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RNA 7, 1416–1424.
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(2002) The human Lsm1–7 proteins colocalize with the
mRNA degrading enzymes Dcp1 ⁄ 2 and Xrn1 in distinct
cytoplasmic foci. RNA 8, 1489–1501.
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lease xrn-1 is essential for ventral epithelial enclosure
during C. elegans embryogenesis. RNA 10, 59–65.
8 Wang Z & Kiledijan M (2001) Functional link between
the mammalian exosome and mRNA decapping
enzyme. Cell 107, 751–762.
9 Coller J & Parker R (2004) Eukaryotic mRNA decap-
ping. Annu Rev Biochem 73, 861–890.
10 Meyer S, Temme C & Wahle E (2004) Messenger RNA
turnover in eukaryotes: pathways and enzymes. Crit
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c ¼ coð1ꢁxÞ
11 Malys N, Carrol K, Miyan J, Tollervey D & McCarthy
JG (2004) The scavenger m7GpppX pyrophosphatase
activity of Dcs1 modulates nutrient-induced responses
in yeast. Nucleic Acids Res 32, 3590–3600.
where c is the substrate concentration at the time of hydro-
lysis (t), co is the initial substrate concentration and x is the
extent of decapping measured as the percentage of hydro-
lyzed substrate.
12 Lall S, Friedman C, Jankowska-Anyszka M, Stepinski
J, Darzynkiewicz E & Davis RE (2004) Contribution of
trans-splicing, 5¢-leader length, cap-poly(A) synergism,
and initiation factor to nematode translation in an
Ascaris suum embryo cell-free system. J Biol Chem 279,
45573–45585.
The initial velocity method was used to calculate the
kinetic parameters for both the fluorimetric and HPLC
methods. The initial velocity (vo) of each reaction was cal-
culated by the linear regression of the substrate concentra-
tion versus time. The Km and Vmax values were determined
from hyperbolic fits to the Michaelis–Menten equation by
nonlinear regression using originpro 7.0 (Microcal Soft-
ware, Northampton, MA, USA).
13 Decker CJ & Parker R (2002) mRNA decay enzymes:
decappers conserved between yeast and mammals. Proc
Natl Acad Sci USA 99, 12512–12514.
14 Cohen LS, Mikhli C, Friedman C, Jankowska-Anyszka
M, Ste˛pinski J, Darzynkiewicz E & Davis RE (2004)
´
_
Acknowledgements
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C. elegans scavenger DcpS. RNA 10, 1609–1624.
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This work was supported by the National Science Sup-
port Project 2008-1010 No. PBZ-MniSW-07 ⁄ I⁄ 2007
and National Institutes of Health Grant AI049558 to
R.E.D. E.D. is a Howard Hughes Medical Institute
International Scholar (Grant No. 55005604).
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M (2004) Functional analysis of mRNA scavenger
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