Kinetic insights of DNA/RNA segment salts catalyzed knoevenagel condensation reaction

Article abstract of DOI:10.1021/cs500882r

In this work, we demonstrated that (i) salts of RNA/DNA segments were capable of catalyzing Knoevenagel condensation at physiological pH 7.0 with efficiency comparable to one of the best enzymes, porcine pancreatic lipase (PPL); and (ii) a broad scope of

Full text of DOI:10.1021/cs500882r

Letter
pubs.acs.org/acscatalysis
Kinetic Insights of DNA/RNA Segment Salts Catalyzed Knoevenagel
Condensation Reaction
Weina Li,†,‡ Sergey N. Fedosov,† Tianwei Tan,*,‡ Xuebing Xu,† and Zheng Guo*,†
†Department of Engineering, Faculty of Science and Technology, Aarhus University, Gustav Wied Vej 10, Aarhus 8000, Denmark
‡College of Life Science and Technology, Beijing University of Chemical Technology, Beisanhuan East Road 15, Beijing 100029,
China
S
* Supporting Information
ABSTRACT: In this work, we demonstrated that (i) salts of
RNA/DNA segments were capable of catalyzing Knoevenagel
condensation at physiological pH 7.0 with efficiency compar-
able to one of the best enzymes, porcine pancreatic lipase
(PPL); and (ii) a broad scope of substrates could be
successfully used in this reaction. Velocity of catalysis was
positively correlated with the content of GC nucleosides in
DNA/RNA; while the 3D-organization in DNA segments
largely contributed to the elevated turnover number of
catalysis. An insight into the reaction mechanism (based on
quantitative analysis of kinetics) elucidated general similarities
between DNA, RNA, and PPL in the substrate binding
mechanisms.
KEYWORDS: Knoevenagel condensation, DNA segments, RNA, kinetic, lipase
ibonucleic acid (RNA) and deoxyribonucleic acid (DNA)1
that the catalytic activity of st-DNA in a Michael reaction could
Rare traditionally regarded as the carriers of information. be associated with the basic nature of their nucleotides.
Yet, they were also discovered as catalysts since Tarasow et al.2
Knoevenagel condensation is a nucleophilic addition with
great potential for synthesis of drug intermediates,16 where basic
found that RNA could catalyze a reaction of carbon−carbon
catalysts (such as piperidine,17 amino acids,18 and enzymes19)
bond formation. Jaschke’s group developed anthracene con-
̈
jugated ribozymes, which mediated the Diels−Alder reaction.3
Though chemically and structurally dissimilar to enzymes, the
ribozymes specifically interacted with their substrates and
products,4 changed active site conformations,5 and were
allosterically activated.6 Michaelase represents another example
of ribozymes reported to be capable of catalyzing the C−C bond
formation reaction.7 Similarly, DNA with either conjugated
anthracene or in combination with the copper complex
accelerated the Diels−Alder reaction.8,9 As reviewed else-
where,10 these self-assembled DNA-based asymmetric catalysts
have demonstrated high enantioselectivity in a variety of
carbon−carbon or carbon−heteroatom bond-forming reactions.
DNA as a catalyst has also been successfully applied to an aldol
reaction11 and Henry12 and Michael reactions.13 It was recently
demonstrated that salmon testes DNA (st-DNA) in complex
with copper catalyzed Michael addition reactions.14 The
mechanisms of DNA/RNA-mediated catalysis are apparently
not identical if considering different reactions performed under
different conditions. Yet, the nucleotides contain a variety of
functional groups including those of ribose/deoxyribose sugars,
phosphate, and organic bases. These groups are capable of
H-bonding, metal coordination, π−π interactions, etc., which
often endows the oligo-nucleotide molecules with catalytic or
enzyme-like activity.7−14 For example, Izquierdo et al.15 found
are widely used. We recently found that the condensation of
aromatic aldehydes and malonitrile can be also catalyzed by
pH-neutral amino acid salts (e.g., Lys•HCl).20 Interestingly,
there are also several reported examples of spontaneous synthesis
of ylidenemalonitriles and the Knoevenagel condensation of aryl
aldehydes with nitroacetonitrile in water in the absence of any
catalyst.21
To maintain the normal biological functions, thousands of
different reactions (affecting human health) take place in the
human body at physiological pH (7.0) and mild conditions. It is
known that malonaldehyde and alkanones are secondary
degradation compounds from fatty acid oxidation, which can
result in the generation of toxic compounds in the human body
via Knoevenagel condensation and other reactions at physio-
logical pH.22 In addition, the role of nucleotides and amino
acids as pre-enzyme catalysts (functional in nonaggressive
solvents at neutral pH) is of fundamental interest for
evolutionary biochemistry.1,2 Herein we explore the potential
of neutralized RNA/DNA as catalysts, which accelerate
Knoevenagel condensation. Ethanol/water mixtures were used
Received: November 20, 2013
Revised: August 8, 2014
Published: August 25, 2014
© 2014 American Chemical Society
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to mediate the reaction and increase the solubility of substrates.
A broad spectrum of substrates was analyzed, and the extensive
kinetic study allowed a more accurate quantitative comparison
of the catalysts. The analysis and characterization of products
were detailed in a recent publication.20 The experimental
procedures, theory, and analysis of the reactions and kinetic
study were described in the Supporting Information.
The commercial DNA/RNA in lyophilized acid forms
exhibited nearly no catalytic activity when dissolved in the
medium (yields of ≈3% in 4 h). When the nucleotides were
neutralized to pH 7.0 by either NaOH or basic amino acids
(L-lysine, L-arginine, or L-histidine) the reaction was dramati-
cally accelerated. The yields became comparable to those of
porcine pancreatic lipase (PPL, one of the best enzymes for
Knoevenagel condensation) and pH-neutral amino acid salts
Arg•HCl and Lys•HCl (Figure 1). Still, the catalytic efficiency
reaction rates from a linear plot.23 We show the Hammett plot
of the Knoevenagel condensation of para-substituted benzalde-
hydes with ethyl cyanoacetate catalyzed by DNA fragment
sodium salt and PPL (Figure 2). It is evident that neither
DNA nor PPL presented a good linear correlation between
σ-constants and the logarithmic ratios of rate constants
(log(k/k0)), as would be expected for a simple chemical
catalyst.24 This result indicates a complex mechanism of
catalysis performed by DNA/PPL, where the formation and
dissociation of temporary ligand-catalyst complexes apparently
occur. Despite some occasional deviations, there was still a clear
tendency that log(k/k0) of DNA/PPL mediated reactions
increases with increasing σ-constant (Figure 2). We observed
earlier a similar result in Lys and Lys•HCl catalyzed reactions.20
Yet, the order of efficient substrate combinations in DNA•-
NaOH-catalyzed reactions was different from the one
obtained with Lys, Lys•HCl, or PPL. Steric effects, associated
with the binding of substrates to the 3D-organized molecules
(DNA, PPL), might be the reason. Therefore, interpretation
of the Hammett plot cannot be straightforward. Electron-
withdrawing groups in the second substrate (methylene com-
pound Bx) increased velocity of condensation (Table 1B).20,25
The maximal rate was attained in the reaction A0 + B1
(Entry 1).
The intrinsic catalytic potential of a nucleotide apparently
depends on the composition of its base-pairs.26 In this context,
we proportionally varied the composition of the mixture of
the individual nucleotides from primarily A-dT (A-U) to
predominantly C-G. The reaction velocity increased as the CG
content increased (Figure 3).27 This acceleration is likely to be
caused by the electronic effect of amine/imine compounds
(better represented in C-G pairs). When analyzing the
individual nucleosides, cytidine exhibited the highest activity,
followed by thymidine deoxyriboside (Table S2). Within an
RNA structure, the properly positioned cytosines and
adenines can potentially serve as general acid or base catalysts
if their ring nitrogen atoms N1 and N3 are protonated
(Table S2(A)).28 The same general acid or base catalysis can
also take place in our study with the degraded DNA (crude
oligonucleotides, < 50 bp). The DNA salts were not expected
to be exclusively in the duplex form because of their partial
degradation, a pH-neutral environment and the elevated
temperatures (40 or 50 °C) of our catalytic experiments.
Unpaired bases are apparently less frequent in the long native
DNA molecules (700 bp), but the presence of “hairpin” and
“cruciform” structures provides various possibilities for the
unusual binding patterns between the DNA molecule and the
substrates. We might speculate whether the molecular basis of
catalysis is associated with the functional groups like R−NH2,
R−OH, and R−CO, which possibly participate in the forma-
tion of transition state complexes during the substrate binding.
Close positioning of the two substrates and/or destabilization
of their electronic densities might cause spontaneous con-
densation, not unusual in such reactions.
Figure 1. Catalytic activity of DNA, RNA, and amino acids-HCl. The
concentrations used: 10 g/L of DNA/RNA salts neutralized by Arg/
His/Lys/NaOH to pH 7.0; 0.0188 mol/L of Arg•HCl, 0.0608 mol/L
of His•HCl, 0.026 mol/L of Lys•HCl, 0.0212 mol/L of NaCl (the
same molar amount of Arg, His, Lys, and NaOH as the corresponding
DNA salts was maintained, Table S1 in the Supporting Information);
10 g/L of PPL. DNA was represented by short crude oligonucleotides
(<50 bp, degraded). Reaction condition: 0.2 M benzaldehyde, 0.3 M
ethyl cyanoacetate, ethanol medium with 4% water added, 40 °C,
500 rpm.
of DNA-Arg and DNA-Lys was somewhat slower than that
of Arg-HCl and Lys-HCl, if calculating it per single amino acid.
Increased activity of the neutralized oligo-nucleotides was not
associated with the ionic strength because no catalytic activity
was observed in the solutions of NaCl (Figure 1).
The catalytic versatility of neutralized DNA/RNA was
examined using the substrates with substituted groups, and
the results were compared to the analogous experiments with
PPL (Table 1). The aromatic aldehydes (Ax) with different
electron-withdrawing groups had a more active electrophilic
center of the carbonyl group and presented higher reaction
velocities in comparison to the substrates bearing the electron-
donating groups. The apparent rate coefficients are shown in
Table 1A, where the highest value was obtained in the reac-
tion A4 + B0 (Entry 4). On the contrary, the reaction A6 + B0
(Entry 6) yielded the lowest apparent rate coefficient. The
Hammett equation has been one of the most widely used
means for the study and interpretation of organic reactions. Its
σ-constants are obtained simply from the ionization properties
of organic acids in solution, and they can frequently predict the
Water is expected to play a pivotal role in achieving high
catalytic efficiency, because (i) the proper water/ethanol ratio
provides a solvent with intermediate polarity important for
the solubility of substrates (Figure S1(A));20 (ii) hydration of
the nucleotide bases affects their protonation patterns and 3D-
structure.29 Different optimal water contents were observed for
different catalysts (Figure S1(B)). A relatively sharp optimum
around 50% was found for short DNA/RNA fragments (<50
bp). The 700 bp DNA•NaOH fragments attained maximal
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Table 1. Condensation Reaction of Substituted Aromatic Aldehydes (A) and Active Methylene Compounds (B)
a
b
(A) Apparent Rate Coefficients Obtained in the Reactions Ax + B0 (Various A) Catalyzed by DNA-NaOH and PPL
(B) Coefficients for A0 + Bx (Various B)
entry(Bx)
kDNA min−1
kPPL min−1
0 R2 = −CN, R3 = −COOEt (pKa 13.1)
1 R2 = R3 = −CN (pKa 11.1)
2 R2 = R3 = −COOEt (pKa 16.4)
0.0038
0.0450
0.00003
0.0090
0.0799
0.00002
a
b
kapp = −log10(At/A0)/t, where t = time. Catalyst: 50 mg of DNA-NaOH or PPL in a 5 mL reaction mixture, 40 °C. *Entry 11: the substrate is
trans-cinnamaldehyde, not joined to the R1 group.
Figure 3. Dependency of the catalytic activity of the nucleoside
mixtures on CG concentration (stipulating different proportions of
AdT (AU) and CG). General reaction condition: 0.055 mmol bases
mixtures (as detailed in the Supporting Information), 0.24 M
benzaldehyde, 0.36 M ethyl cyanoacetate, medium of 4% water
added to ethanol, within 2 h, 40 °C, 500 rpm.
Figure 2. Hammett plot for the Knoevenagel condensation of para-
substituted benzaldehydes with ethyl cyanoacetate catalyzed by DNA
fragment sodium salt (circles) and PPL (squares). The reaction
conditions correspond to those in Table 1.
Spectroscopic analysis is known to be an effective tool to
examine the individual interaction between substrate and
catalyst.30,31 The UV−vis absorption curve of DNA•NaOH
+A0 was different obviously from the curve of DNA•NaOH
in intensity and peak shifting, indicating the formation of the
reaction velocity at a lower water content of 5%−25%.
A very broad optimum was found for the enzyme PPL
(5%−50%).
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c d
,
Table 2. Kinetic Constants of a General Velocity Equation of a Bisubstrate Reaction
a
700 bp DNA-NaOH
DNA-NaOH
DNA-Lys
RNA-NaOH
RNA-Lys
Lys-HCl
PPL
0.32
Vmax (M/min)
KsA (M)
0.182
≤0.03
≤0.04
1.02
0.042
0.037
0.060
0.463
0.750
0.028
1.500
0.934
0.034
0.039
0.049
0.317
0.397
0.021
1.619
0.817
0.030
0.111
0.177
0.239
0.380
0.027
1.119
0.898
0.035
0.124
0.110
0.374
0.334
0.021
1.658
0.824
0.225
0.364
0.271
1.93
≤0.07
≤0.05
0.707
KsB (M)
KmA (M)
KmB (M)
1.36
1.440
0.250
0.899
0.894
0.50
b
Catal. (M)
0.014
13.35
0.822
2.6 × 10−5
≈12300
k+ (min−1
Adj. R2
)
0.813
a
b
The fitting parameters for PPL are not precise, because variation of KsA within the range of 0.07−1 × 10−5 M gives nearly the same R2. The catalyst
concentration corresponds to (i) a single nucleotide in the chains of DNA and RNA, (ii) individual Lys in Lys-HCl, and (iii) the whole molecule of
c
d
PPL, see also discussion in the main text. Corresponding, for example, to a rapid-equilibrium random binding mechanism. Deoxyribonucleic acid
sodium salt was from herring testes (type XIV, ca. 700 bp), DNA was from herring sperm (degraded, crude oligonucleotides <50 bp), ribonucleic
acid was from torula yeast (Type VI) ; DNA without other notation refers to the degraded, crude oligonucleotides <50 bp. The relative standard
errors of parameters were on average as follows: 0.53·Vmax
,
2.3·KsA, 2.2·KsB, 0.84·KmA
.
0.84·KmB. Nomenclature: KsX describes the
dissociation of X from the catalyst (usually E+X↔EX), KmX describes the corresponding Michaels constant. All constants are related to each other
according to the ratio KsA/KmA = KsB/KmB in the case of a random binding mechanism. The values of KsB were assessed from this assumption. Adj. R2
stands for coefficient of determination adjusted for the number of parameters in the model. Reaction conditions: 50 °C and optimal water
concentration in ethanol solvent. Water concentration is, probably, a “hidden” component of all rate and dissociation constants.
DNA•NaOH-A0 complex; whereas the curves of DNA•NaOH
and DNA•NaOH+B0 are much similar, indicating a different
type of catalyst-substrate interactions (Supporting Information,
Page P8). The possibility of the independent interaction of
substrates A0 or B0 with DNA•NaOH or RNA•NaOH was
established by monitoring the intrinsic fluorescent response
of the catalysts toward addition of either A0 or B0. Detectable
changes of fluorescence took place in the approximate ranges of
concentration 0−0.6 mM for substrate A0 and 0−1 mM for
substrate B0 (Supporting Information, Pages S10−S11),
indicating the independent binding of A0 or B0 with a catalyst
could take place without the presence of the other. The
responses of fluorescence quenching as a function of quencher
concentrations at different temperatures were also analyzed by
the Stern−Volmer equation (Figures S3 and S4). For both
DNA•NaOH and RNA•NaOH, substrate A0 presented a
bigger quenching constant than B0 at the same temperature;
and the constant decreases as temperature increases, indicating
a ground-state complex formation between A0 and catalysts.
Substrate B0 demonstrated a different fluorescence quenching
behavior for DNA•NaOH (Ksv decreases as temperature
increases) and RNA•NaOH (Ksv increases as temperature
increases), suggesting the former is static quenching and
the latter belongs to dynamic quenching.30 This also indicates
the interaction may vary among different substrate-catalyst
combinations. The above observations are inconsistent with
UV−vis spectra analysis that substrate A0 and B0 may engender
different interactions with catalysts, which might be related to
their molecular structure differences.31
of a bisubstrate reaction, which includes the largest set of
parameters.32
Vmax
v =
;
KsA KmB
Vmax = k+·e0
KmA
a
KmB
b
1 +
+
+
a
(1)
b
This equation contains four parameters, where KmA and KmB
correspond to the Michaelis constants of the substrates A and
B, respectively; KsA is the substrate constant of A; and Vmax is
the maximal velocity. The latter parameter can be expressed
via e0 (the total concentration of catalyst E) and k+, which is
the turnover number (i.e., the maximal number of catalytic
conversions per time unit per active site).
The full equation (eq 1) usually corresponds to either the
mechanism of fast-equilibrium random binding (Scheme 1) or
Scheme 1. Proposed Kinetic Mechanism DNA/RNA-Salts
Appears To Progress via the Shown Catalytic Cycle
However, it should be emphasized that fluorescence analysis
is incapable of delineating the interaction between catalyst and
bisubstrates, cooperation and sequential collisions,30−32 because
the reaction may occur when catalyst/A0/B0 are simultaneously
presented. Meanwhile, the reaction velocity recorded at a
single combination of the two substrates does not provide
detailed insight into the functioning of a catalyst.32 Therefore,
we investigated the probable substrate-binding mechanisms
using various combinations of substrates A0 and B0, which is
the particular interest of this work. The DNA/RNA-catalyzed
reactions were compared to those in the presence of lipase
PPL and amino acid Lys•HCl (Table 2). Approximation of our
data was performed by the general velocity equation (eq 1)
the steady state ordered binding scheme. Very small values of
some constants might provide additional information about
possible cancelation of the elements in the denominator. In
such a way, alternative mechanisms can be hypothesized, e.g.
fast-equilibrium ordered binding, ping-pong, etc. The exper-
imental data were presented in 3D-coordinates (initial velocity
v as a function of A- and B-concentrations) and subjected
to computer fitting (Figure 4). The coefficients of best
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Figure 4. Dependence of the reaction velocity v on the substrate concentrations (a0, b0) for different catalysts: (A) DNA-NaOH fragments of 700
bp; (B) DNA-NaOH fragments of <50 bp; (C) Lys-HCl; (D) PPL.
approximation are shown in Table 2. The goodness of fit was
assessed via coefficient of determination R2 adjusted for the
number of parameters.
even though each base pair has the same catalytic potency. In
the absence of any additional information, we ascribed activity
to each individual nucleotide (amino acid) within the joined
chain, when making a comparison between the catalysis of
widely different masses. The turnover numbers of both DNA
and RNA salts were slightly higher than the k+ value of simple
catalyst Lys•HCl (see Table 2). Efficiency of each nucleotide
within 700 bp DNA•NaOH increased if compared to a shorter
chain of 50 bp DNA•NaOH. Substitution of NaOH by Lys
provided only marginal effects. The enzyme PPL had the highest
turnover number of ≈12300 min−1 (Table 2), if calculating k+
per the whole enzyme molecule composed of 450 amino acids.
Yet, k+ of PPL was reduced to 27 min−1 (if calculating its value
per single amino acid). This is only ≈2 times higher than k+ of a
nucleotide within 700 bp DNA•NaOH. Comparison of k+ for
the whole molecules of ≤50 bp DNA (<100 nucleotides) and
PPL (i.e., two molecular species of similar complexity and size)
gave the difference by 2 orders of magnitude (k+ = 12300 min−1
and 150 min−1, respectively). Calculation of k+ per 52 kDa of
mass (the mass of PPL, approximately equal to 80 bp DNA)
gives the values of 12300 min−1 (PPL), 120 min−1 (52 kDa
within DNA of 50 bp), and 1070 min−1 (52 kDa within DNA of
700 bp). Disregarding the evaluation method of k+, it is visible
that a higher potential for self-organization in the long DNA
All parameters of Table 2 had measurable values and did not
tend to zero (or negative values). We, therefore, hypothesize
the random binding to be a plausible model. It stipulates
assembly of the reactants into a ternary complex (e.g., A•E•B;
E stands for catalyst), where the sequence of ligand binding
is unimportant. In our case, the attachment of one substrate
apparently hinders the binding of the other (e.g., KsA < KmA),
probably, because A and B compete for nearly the same specific
site on the catalyst. The mechanism can be also interpreted as
a variant of the ordered binding with several steady state steps
but not the fast equilibrium ordered binding. In the current
work we did not pursue precise identification of the model but
used the kinetic analysis as a tool to quantify and compare the
main parameters of different catalysts.
An important characteristic of any complex catalyst is its
turnover number k+ or kcat (equal to Vmax/e0). The meaning
of k+ is not absolutely straightforward if the concentration
of the active site (e0) is not established. The usual assumption
of 1 catalytic site per whole molecule unnecessarily favors large
molecules. For example, k+ of 1 bp DNA becomes 10-fold less
than k+ of the identical 10 bp DNA because e0(1bp)/e0(10bp) = 10,
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Notes
fragments gives the higher catalytic potencies approaching those
of enzymes.
The authors declare no competing financial interest.
The presence of either Na+ or basic amino acids associated
with phosphates of DNA and RNA chains had apparently little
effect on the turnover number as long as nucleotides remained
neutralized. We have, therefore, tested if any conformational
changes take place in DNA/RNA upon the binding of amino
acids. We found that (i) all the DNA/RNA salts remained in
B-form according to CD spectra, while RNA had a little blue
shift; (ii) DNA•Lys, DNA•Arg increased fluorescence emission
and showed little sensitivity to solvent polarity when dissolved
ACKNOWLEDGMENTS
■
This work was supported by the National Basic Research Program
of China (973 program: 2013CB733600, 2012CB725200), the
National Nature Science Foundation of China (21390202), and
the National High-Tech R&D Program of China (2014AA022101,
2014AA021904). The support provided by China Scholar-
ship Council (No. +2011688018) during a visit to Aarhus is
acknowledged.
1
in 55% ethanol or water; (iii) H NMR and CD spectra of
DNA•NaOH and DNA•Lys verified that Na+ and Lys indeed
interacted electrostatically with phosphate groups (see details in
Spectrum Study in the Supporting Information).
REFERENCES
■
(1) (a) Kruger, K.; Grabowski, P. J.; Zaug, A. J.; Sands, J.;
Gottschling, D. E.; Cech, T. R. Cell 1982, 31, 147−157. (b) Guerrier-
Takada, C.; Gardiner, K.; Marsh, T.; Pace, N.; Altman, S. Cell 1983,
35, 849−857.
Combining the current knowledge with the generally
accepted Knoevenagel condensation mechanisms,14−19,28
we can propose the following pathway (Scheme 1). First, a
ternary complex A•E•B is formed via a fast random binding
or according to the ordered steady state mechanism (both
described by eq 1). The substrates are located very closely and
partially hinder the binding of each other to catalyst E (because
KmA > KsA). At the second step, a proton is transferred from
ethyl cyanoacetate to the catalyst (e.g., DNA/RNA-salt) and
enolate ion is formed. Then another substrate benzaldehyde
accepts the proton and simultaneously becomes connected
to enolate ion with formation of a carbon−carbon bond.
Finally, the Knoevenagel adduct is released from the catalyst.
An alternative kinetic scheme (also compliant with eq 1)
suggests a temporary binding of the active hydrogen compound
(B) to the catalyst followed by abstraction of proton. Finally,
a direct collision with benzaldehyde (with a very weak or no
complex formation) leads to the condensation. A comparison
of kinetics for simple and complex catalysts (planned in the
future) might shed some light on this subject.
In conclusion, we have demonstrated that RNA/DNA
oligonucleotides can effectively catalyze the Knoevenagel
condensation reaction at physiological pH. The best achieved
efficiency was comparable to the enzyme PPL. Velocity of
catalysis was positively correlated to the content of GC
nucleosides. A long strand of 700 bp DNA•NaOH exhibited
higher turnover number than 50 bp DNA•NaOH, DNA•Lys,
and Lys•HCl, which stresses the importance of higher 3D-
organization. The deoxyribo- or ribophosphate backbone was
not directly involved in catalysis. Quantitative characterization
of kinetics elucidated general similarities of DNA, RNA, lipase
PPL, and Lys in the substrate binding mechanisms (compliant
with general equation of a bisubstrate reaction). This work
demonstrated a novel group of catalysts and illustrated possible
mechanisms of catalysis relevant to evolutionary biochemistry
of simple molecules.
(2) Tarasow, T. M.; Tarasow, S. L.; Eaton, B. E. Nature 1997, 389,
54−57.
(3) (a) Seelig, B.; Jaschke, A. Tetrahedron Lett. 1997, 38, 7729−7732.
̈
(b) Seelig, B.; Jaschke, A. Chem. Biol. 1999, 6, 167−176. (c) Seelig, B.;
̈
Jaschke, A. Bioconjugate Chem. 1999, 10, 371−378. (d) Seelig, B.;
Keiper, S.; Stuhlmann, F.; Jaschke, A. Angew. Chem., Int. Ed. 2000, 39,
̈
̈
4576−4579.
(4) Stuhlmann, F.; Jaschke, A. J. Am. Chem. Soc. 2002, 124, 3238−
̈
3244.
(5) Wombacher, R.; Jaschke, A. J. Am. Chem. Soc. 2008, 130, 8594-−
̈
8595.
(6) Helm, M.; Petermeier, M.; Ge, B.; Fiammengo, R.; Jaschke, A. J.
̈
Am. Chem. Soc. 2005, 127, 10492−10493.
(7) Sengle, G.; Eisenfuhr, A.; Arora, P. S.; Nowick, J. S.; Famulok, M.
̈
Chem. Biol. 2001, 8, 459−473.
(8) Chandra, M.; Silverman, S. K. J. Am. Chem. Soc. 2008, 130, 2936−
2937.
(9) Boersma, A. J.; Klijn, J. E.; Feringa, B. L.; Roelfes, G. J. Am. Chem.
Soc. 2008, 130, 11783−11790.
(10) (a) Park, S.; Sugiyama, H. Molecules 2012, 17, 12792−12803.
(b) Park, S.; Sugiyama, H. Angew. Chem., Int. Ed. 2010, 49, 3870−
3878.
(11) Sun, G.; Fan, J.; Wang, Z.; Li, Y. Synlett. 2008, 16, 2491−2494.
(12) Fan, J.; Sun, G.; Wan, C.; Wang, Z.; Li, Y. Chem. Commun. 2008,
32, 3792−3794.
(13) De Rosa, M.; Di Marino, S.; D’Ursi, A. M.; Strianese, M.;
Soriente, A. Tetrahedron 2012, 68, 3086−3091.
̀
(14) (a) Coquiere, D.; Feringa, B. L.; Roelfes, G. Angew. Chem., Int.
Ed. 2007, 46, 9308−9311. (b) Li, Y.; Wang, C.; Jia, G.; Lu, S.; Li, C.
Tetrahedron 2013, 69, 6585−6590. (c) Megens, R. P.; Roelfes, G.
Chem. Commun. 2012, 48, 6366−6368. (d) Roelfes, G. Org. Biomol.
Chem. 2010, 817, 3868−3873.
(15) Izquierdo, C.; Luis-Barrera, J.; Fraile, A.; Aleman
Commun. 2014, 44, 10−14.
́
, J. Catal.
(16) Knoevenagel, E. Ber. Dtsch. Chem. Ges. 1894, 27, 2345−2346.
(17) Kuzemko, M. A.; Van Arnum, S. D.; Niemczyk, H. J. Org. Process
Res. Dev. 2007, 11, 470−476.
(18) Li, Y.; Chen, H.; Shi, C.; Shi, D.; Ji, S. J. Comb. Chem. 2010, 12,
231−237.
ASSOCIATED CONTENT
* Supporting Information
■
(19) (a) Lai, Y. F.; Zheng, H.; Chai, S. J.; Zhang, P. F.; Chen, X. Z.
Green Chem. 2010, 12, 1917−1918. (b) Kapoor, M.; Gupta, M. N.
Process Biochem. 2012, 47, 555−569. (c) Liu, Z.-Q.; Liu, B.-K.; Wu, Q.;
Lin, X.-F. Tetrahedron 2011, 67, 9736−9740.
S
Details on DNA/RNA salt preparation, pH influence
assumption, the kinetic and spectral characterization data for
catalysts. This material is available free of charge via the
Internet at http://pubs.acs.org.I
(20) Li, W.; Fedosov, S. N.; Tan, T.; Xu, X.; Guo, Z. Appl. Biochem.
Biotechnol. 2014, 173, 278−290.
(21) (a) Bigi, F.; Conforti, M. L.; Maggi, R.; Piccinno, A.; Sartori, G.
Green Chem. 2000, 2, 101−103. (b) Amantini, D.; Fringuelli, F.;
Piermatti, O.; Pizzo, F.; Vaccaro, L. Green Chem. 2001, 3, 229−232.
(22) (a) Janero, D. R. Free Radical Biol. Med. 1990, 9, 515−540.
(b) Esterbauer, H.; Schaur, R. J.; Zollner, H. Free Radical Biol. Med.
1991, 11, 81−128.
AUTHOR INFORMATION
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Corresponding Authors
*E-mail: guo@mb.au.dk.
*E-mail: twtan@mail.buct.edu.cn.
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dx.doi.org/10.1021/cs500882r | ACS Catal. 2014, 4, 3294−3300

ACS Catalysis
Letter
(23) (a) Hammett, L. P. J. Am. Chem. Soc. 1937, 59, 96−103.
(b) Mansch, C.; Leo, A.; Taft, R. W. Chem. Rev. 1991, 97, 165−195.
(24) Cabello, J. A.; Campelo, J. M.; Garcia, A.; Luna, D.; Marinas, J.
M. J. Org. Chem. 1984, 49, 5195−5197.
(25) Venkatanarayana, M.; Dubey, P. K. Synth. Commun. 2011, 42,
1746−1759.
(26) (a) Brackett, D. M.; Dieckmann, T. ChemBioChem 2006, 7,
839−843. (b) Hougland, J. L.; Sengupta, R. N.; Dai, Q.; Deb, S. K.;
Piccirilli, J. A. Biochemistry 2008, 47, 7684−7694. (c) Bevilacqua, P. C.;
Yajima, R. Curr. Opin. Chem. Biol. 2006, 10, 455−464. (d) Chandra,
M.; Silverman, S. K. J. Am. Chem. Soc. 2008, 130, 2936−2937.
(e) Park, S.; Ikehata, K.; Watabe, R.; Hidaka, Y.; Rajendran, A.;
Sugiyama, H. Chem. Commun. 2012, 48, 10398−10400.
(27) Rosati, F.; Boersma, A. J.; Klijn, J. E.; Meetsma, A.; Feringa, B.
L.; Roelfes, G. Chem.Eur. J. 2009, 15, 9596−9605.
(28) Nakano, S.-i.; Chadalavada, D. M.; Bevilacqua, P. C. Science
2000, 287, 1493−1497.
(29) (a) Rosenberg, J. M.; Seeman, N. C.; Day, R. O.; Rich, A. J. Mol.
Biol. 1976, 104, 145−167. (b) Saenger, W. Principles of nucleic acid
structure; Springer-Verlag: New York, 1984; Vol. 7, p 368.
(30) (a) Rahman, M. H.; Maruyama, T.; Okada, T.; Yamasaki, K.;
Otagiri, M. Biochem. Pharmacol. 1993, 46, 1721−1731. (b) Zhang, Y.-
Z.; Zhou, B.; Zhang, X.-P.; Huang, P.; Li, C.-H.; Li, Y. J. Hazard.
Mater. 2009, 163, 1345−1352.
(31) (a) Ross, P. D.; Subramanian, S. Biochemistry 1981, 20, 3096−
3102. (b) Zhang, Z.; Xiong, D.; Chen, L.; Kang, Q.; Zeng, Z.
Spectrochim. Acta, Part A 2012, 96, 132−138.
(32) (a) Alberty, R. A. J. Am. Chem. Soc. 1953, 75, 1928−1932.
(b) Fedosov, S. N.; Brask, J.; Pedersen, A. K.; Nordblad, M.; Woodley,
J. M.; Xu, X. J. Mol. Catal. B: Enzym. 2013, 85−86, 156−168.
NOTE ADDED AFTER ASAP PUBLICATION
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After this paper was published ASAP on August 27, 2014,
corrections were made to the Acknowledgments. The corrected
version was reposted September 5, 2014.
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