Tunable Artificial Enzyme-Cofactor Complex for Selective Hydrolysis of Acetals

Article abstract of DOI:10.1021/acs.joc.0c02519

Enzymes frequently use unimpressive functional groups such as weak carboxylic acids for efficient, highly selective catalysis including hydrolysis of acetals and even amides. Much stronger acids generally have to be used for such purposes in synthetic systems. We report here a method to position an acidic group near the acetal oxygen of 2-(4-nitrophenyl)-1,3-dioxolane bound by an artificial enzyme. The hydrolytic activity of the resulting artificial enzyme-cofactor complex was tuned by the number and depth of the active site as well as the hydrophobicity and acidity of the cofactor. The selectivity of the complex was controlled by the size and shape of the active site and enabled less reactive acetals to be hydrolyzed over more reactive ones.

Full text of DOI:10.1021/acs.joc.0c02519

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Article
Tunable Artificial Enzyme−Cofactor Complex for Selective
Hydrolysis of Acetals
Ishani Bose, Shixin Fa, and Yan Zhao*
Cite This: J. Org. Chem. 2021, 86, 1701−1711
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ABSTRACT: Enzymes frequently use unimpressive functional
groups such as weak carboxylic acids for efficient, highly selective
catalysis including hydrolysis of acetals and even amides. Much
stronger acids generally have to be used for such purposes in
synthetic systems. We report here a method to position an acidic
group near the acetal oxygen of 2-(4-nitrophenyl)-1,3-dioxolane
bound by an artificial enzyme. The hydrolytic activity of the
resulting artificial enzyme−cofactor complex was tuned by the
number and depth of the active site as well as the hydrophobicity
and acidity of the cofactor. The selectivity of the complex was
controlled by the size and shape of the active site and enabled less reactive acetals to be hydrolyzed over more reactive ones.
Our group has been interested in creating enzyme-mimetic
catalysts using different platforms including foldamers and
cross-linked micelles/reverse micelles.10−13 More recently, we
developed a method to perform molecular imprinting14,15 in
surface−core doubly cross-linked micelles.16 Molecular
imprinting is a powerful technique to construct guest-tailored
binding sites through templated polymerization. Molecularly
imprinted polymers are useful in many applications including
catalysis.17−30 When performed in the nanospace of a micelle,
molecular imprinting yields receptors capable of distinguishing
the shift of a single methyl group or addition/deletion of a
single methylene in the guest.31 These so-called molecularly
imprinted nanoparticles (MINPs) resemble enzymes in their
water solubility, the nanodimension, and a hydrophilic/
hydrophobic surface/core morphology. Complex biomolecules
including oligosaccharides32,33 and peptides34 can be used as
templates with appropriate functional monomers included in
the MINP preparation.
INTRODUCTION
■
Enzymes are fascinating catalysts from nearly every perspec-
tive.1 They catalyze some of the most challenging reactions in
nature with astounding efficiency under mild conditions. They
possess exquisite selectivity, which are able to pick a particular
substrate to react when numerous other ones are present in the
same mixture. Even more amazing is that they use
“unimpressive” functional groups to accomplish these extra-
ordinary tasks. Serine protease, as an example, relies on a
serine−histidine−aspartic acid catalytic triad to hydrolyze
amide bonds that would otherwise require concentrated
hydroxide or strong acids at elevated temperatures.2 Both
aspartic protease3 and glycosidase4,5 have a pair of carboxylic
acids in their active sites for the hydrolysis of amides and
acetals (in glycosides), respectively.
Central to the catalytic performance of an enzyme is its
molecular recognition. When only the substrate can enter the
catalytic active site or induce the necessary conformational
change to turn on catalysis, other structural analogues will stay
intact even if they have the same reactive group as the
substrate. In recent years, increasing attention has been paid
toward synthetic catalysts with similar molecular recognition
features. Some exciting developments emerged from these
studies, including selective activation of the C−H bond,6
Diels−Alder reaction at unreactive sites of anthracene,7 acidic
catalysis under basic conditions,8 and phosphorylation of a
particular hydroxyl among numerous others with similar
reactivity.9 Despite the tremendous progress made, however,
these enzyme mimics are rather rudimentary compared to their
natural counterparts and construction of multifunctionalized
active sites with accurately positioned catalytic groups remains
difficult.
In this paper, we report an MINP-based artificial enzyme−
cofactor complex for selective hydrolysis of acetals. The MINP
was able to bind both the substrate and the acid catalyst in its
imprinted site and help position the acidic group near the
acetal oxygen of the substrate. Close proximity of the reactive
group and the catalytic group allowed weak carboxylic acids to
work as well as much stronger sulfonic acids in certain
Received: October 22, 2020
Published: January 5, 2021
https://dx.doi.org/10.1021/acs.joc.0c02519
© 2021 American Chemical Society
J. Org. Chem. 2021, 86, 1701−1711
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Scheme 1. Preparation of Noncovalent MINP with a Schematic Representation of the Cross-Linked Structure
Scheme 2. Synthesis of Compounds 1a−d
constructs. A key feature of the system is the facile tuning of
catalytic activity by easy “plug-and-play” switching of the acid
cofactor, which could not be achieved if the acid groups are
covalently attached to the active site.35 In addition, both the
number of acid cofactors and the depth of the active site
impacted the catalysis strongly. The artificial enzyme−cofactor
complex was able to distinguish subtle structural features of the
substrates and even hydrolyze less reactive acetals over more
reactive onesa feature frequently seen in enzymatic catalysis
but rare in chemical catalysis.
was to imprint a construct with a substratelike and a catalyst-
like moiety (Scheme 1). Template 1 is color-coded to illustrate
the purpose of its different substructures: the green moiety is
the space holder for a noncovalently bound catalyst 2, the
magenta resembles the substrate 2-(4-nitrophenyl)-1,3-dioxo-
lane (3a), and the yellow is for the carboxylate group that
serves as a hydrophilic anchor. The para-nitro-substituted
acetal was chosen for its low background activity. The anionic
anchor was used to keep the template molecule and, in turn,
the imprinted site close to the surface of the micelle. Such a
location is expected to facilitate the removal of the template
after imprinting and also make the imprinted site readily
accessible to the substrate during the catalysis. If a binding site
could be faithfully imprinted from the template, the MINP
would be able to bring substrate 3a and acid catalysts (2a−d)
together in a way similar to what an enzyme does in an
RESULTS AND DISCUSSION
■
Design and Synthesis of MINP Catalysts. To make
weakly acidic carboxylic acids into effective catalysts, enzymes
have to position the acids in close proximity to the reactive
functionality. Our strategy to enforce such a spatial relationship
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a
Table 1. Binding Data for MINPs Determined by ITC and Fluorescence Titration
b
entry
host
guest (sodium salt of)
Ka (×106 M−1
)
−ΔG (kcal/mol) −ΔH (kcal/mol) TΔS (kcal/mol)
N
1
2
3
4
5
6
7
8
MINP(1a)
MINP(1a)
MINP(1a)
MINP(1a)
MINP(1b)
MINP(1b)
MINP(1b)
MINP(1b)
1a
2a
2b
2c
1b
2a
2c
2d
16.4 4.2 (10.5 7.44)
3.4 0.3 (3.15 0.35)
2.1 0.2
9.83
8.90
8.61
6.59
24.1 0.45
74.2 1.47
23.1 0.40
1.6 0.02
−14.27
−65.30
−14.48
4.99
0.9 0.01
0.7 0.01
1.4 0.02
c
0.068 0.002 (0.070 0.006)
2.5 0.3 (2.43 0.9)
8.72
c
45.9 0.77
c
−37.18
1.2 0.01
c
d
c
<0.0004 ( )
0.9 0.15 (1.03 0.65)
0.7 0.04
8.12
7.97
1.0 0.03
3.6 0.08
7.12
4.36
1.2 0.02
1.1 0.02
a
All the MINPs were prepared with a DVB/surfactant ratio of 0.5:1. Titrations were performed in duplicate using sodium salts of the templates and
acid cofactors in 10 mM HEPES buffer (pH 7.0) with 2% DMSO. The errors between the runs were <10%. The binding constants in parentheses
b
c
were obtained from fluorescence titration. N is the number of binding sites per MINP determined by ITC. Binding was extremely weak, and the
binding constant was estimated from ITC. Binding in the fluorescence titration was extremely weak, and the binding constant could not be
d
obtained.
enzyme−cofactor complex, with the acid group pointing at the
acetal oxygen for catalytic hydrolysis.
process is directly proportional to the hydrophobic surface area
buried upon binding.38−40 The entire template was certainly
larger than the acid cofactor which could only occupy a portion
of the imprinted site.
MINP(1a) bound sodium 2-naphthalenecarboxylate (the
conjugate base of 2c) much more weakly than sodium 1-
pyrenecarboxylate (the conjugate base of 2a), by about 50
times (Table 1, entries 2 and 4). The result further supported
successful imprinting, as the smaller naphthalene would not be
able to fit fully inside the larger imprinted site created for the
pyrenyl group of 1a. Filling the unoccupied space of the
hydrophobic imprinted site with water molecules was
unfavorable.
When the binding between an MINP and its own template
was compared, the Ka value was noticeably lower for
MINP(1b) than for MINP(1a) (compare entries 1 and 5 of
Table 1). This result was also expected from the hydrophobic
binding that depended on the surface area of the host−guest
interaction. Once again, the carboxylate salt of 2c was bound
more strongly than the sulfonate salt of 2d, but the difference
was not large (entries 7 and 8). Note that a mismatched but
smaller cofactor (i.e., 2c) could still enter the imprinted site of
MINP(1a) (entry 4). For MINP(1b), the mismatched
cofactor (i.e., 2a) was not bound at all, apparently because
of its larger size than the imprinted pocket (entry 6).
Validation of the Catalytic Design Hypothesis. Our
ITC and fluorescence titration could only measure the binding
of guests with some water solubility, including the templates
and the sodium salts of the acid cofactors. The targeted
substrate, 2-(4-nitrophenyl)-1,3-dioxolane (3a), was insoluble
in water. Its binding by the MINP, nonetheless, was evident
from the catalytic hydrolysis by the MINP-cofactor complex
(Figure 1a). In the absence of MINP, 2a was rather inactive
under our experimental conditions (85 °C in D2O) even at a
concentration of 200 μM or 20 mol % of the substrate. With
the matching MINP in the solution, both 1-pyrenecarboxylic
acid (2a) and 2-naphthalenecarboxylic acid (2c) became active
and could afford 45−61% yield under otherwise identical
reaction conditions. The hydrolytic yield stabilized above a 1:1
cofactor/MINP ratio. For most of our experiments, we used a
2:1 ratio to ensure that the binding site of the MINP was
saturated with the acid cofactor.
The preparation of the MINP started with spontaneous
incorporation of the largely hydrophobic 1 into the micelle of
surfactant 4, together with 2,2-dimethoxy-2-phenyl acetophe-
none (DMPA, a photoinitiator) and various amounts of
divinylbenzene (DVB). The tripropargylammonium head-
group of the surfactant allowed us to cross-link the surface
of the micelle by diazide 5 and functionalize it with monoazide
6 by the Cu(I)-catalyzed click reaction. Molecular imprinting
took place in the micellar core, when free-radical polymer-
ization cross-linked the vinyl groups of DVB and the
methacrylate of 4, essentially to “solidify” the core around
the template. Template removal happened during precipitation
of the MINP from acetone and repeated solvent washing.16
The synthesis of our templates 1a−d is shown in Scheme 2.
Characterization of MINP Catalysts. The surface- and
1
core-cross-linking of the micelle were monitored by H NMR
spectroscopy (Figure S1 in the Supporting Information). The
size of the cross-linked micelle was determined by dynamic
light scattering (DLS, Figures S2−S4), confirmed by trans-
mission electron microscopy.36 The formation of the imprinted
site was studied by isothermal titration calorimetry (ITC), one
of the most reliable methods to study intermolecular
interactions.37 A particular benefit of ITC is the simultaneous
determination of binding enthalpy and binding free energy as
well as the number of binding sites per nanoparticle (N) that
could help us estimate the yield of micellar imprinting. For
selected cases, we also confirmed the binding by fluorescence
titration and found that the values obtained (shown in
parentheses in Table 1) agreed well with those from ITC.
As shown in Table 1 (entry 1), MINP(1a) bound its
template 1a with a binding constant (Ka) of 16.4 × 106 M−1,
which translates to nearly 10 kcal/mol in binding free energy.
In our preparation, a 50:1 surfactant/template ratio was used,
and DLS indicated ∼50 cross-linked surfactants per nano-
particle (Supporting Information). The ideal yield of the
imprinted site was thus an average of 1 binding site per
nanoparticle. The number of binding sites determined by ITC
was ∼0.9, suggesting a high yield in the formation and vacation
of the imprinted sites.
MINP(1a) bound the sodium salts of pyrene acids 2a and
2b more weakly than its template 1a by about 5−8 times
(Table 1, entries 2−3)the sodium salts had to be used
because the acids were insoluble in water. The result was
reasonable because the driving force in a hydrophobic binding
Figure 1b shows the hydrolysis of 3a under different catalytic
conditions as a function of solution pH. In the absence of
MINP(1a), neither 1-pyrenecarboxylic acid (2a) nor 1-
pyrenesulfonic acid (2b) showed any catalytic effect beyond
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two acid cofactors in Figure 1b (the blue and red lines)
suggests a small advantage of the sulfonic acid over carboxylic
acid at lower pH values but the opposite at higher pH values.
According to the pH profiles, the reversal was a result of the
more gradual transition seen in 2a.
Encouraged by the abilities of the MINP to help the
hydrolysis of 3a, we performed additional structure−activity
studies to understand the catalysis better and summarize the
results in Table 2. A strong dependence of the catalysis on the
Table 2. Hydrolysis of 3a catalyzed by MINP-Acid Cofactor
Complexes
a
Figure 1. (a) Yield of hydrolysis for 3a as a function of acid cofactor
in D2O at 85 °C under different conditions. Acid 2a and 2c were used
with MINP(1a) and MINP(1b), respectively. The reaction without
the MINP was performed with 2a. (b) Hydrolysis of 3a as a function
of solution pH after 6 h at 85 °C under different catalytic conditions.
Reactions were performed in duplicate with the yields determined by
1H NMR spectroscopy using 1,4-dibromobenzene as an internal
standard. [3a] = 1.0 mM. [MINP] = 50 μM. [2a] = [2b] = 100 μM.
The MINPs were prepared with a DVB/surfactant ratio of 0.5:1.
entry
MINP
DVB/4
acid cofactor
yield (%)
1
2
3
4
5
6
7
8
MINP(1a)
MINP(1a)
MINP(1a)
MINP(1a)
MINP(1a)
MINP(1a)
MINP(1a)
MINP(1a)
MINP(1b)
MINP(1b)
MINP(1b)
MINP(1b)
MINP(1a−b)
none
0
0.5
1
0
0.5
1
0.5
0.5
0.5
0.5
0.5
0.5
0.5
-
2a
2a
2a
2b
2b
2b
2c
2d
2a
2b
2c
2d
none
2a−d
none
2a−d
2a
23
59
42
33
71
57
29
37
3
1
2
2
2
3
1
2
3
1
2
2
3
that of the buffer and hydrolysis of the acetal was only
observed below pH 7, consistent with the acid-catalyzed
hydrolysis.41 Once MINP(1a) was added, both 2a and 2b
became quite effective in the hydrolysis. Even though the
reaction slowed down at higher pH values, it was very
impressive that the MINP-cofactor complex could perform an
acid-catalyzed hydrolysis under basic conditions (pH 9−10), a
feature rarely seen in synthetic catalysts.8 By itself, MINP(1a)
was also inactive, giving the same yields as the buffers.
The large drop in the hydrolytic yield over pH 6−8 suggests
that a deprotonation occurred with the acid cofactor at higher
pH values. The transition, however, was much larger than the
pKa of sulfonic acid (−7) or carboxylic acid (4−5).42 It is
known that the microenvironment around an acid or base can
strongly impact its acidity/basicity, whether in the active site of
an enzyme43 or in a synthetic host.44−46 For example, the
ammonium side chain of a lysine has a pKa of 10.6 in water but
shifted to 5.6 in the active site of acetoacetate decarboxylase.
The two most common strategies to shift the pKa of a
functional group are through hydrophobic interactions47 and
ionic interactions.48 The former takes advantage of the change
of solvation during protonation or deprotonation. Because an
ionic group is better solvated by polar solvents such as water, it
becomes more difficult to protonate an amine or deprotonate a
carboxylic acid when the functional group migrates into a
hydrophobic microenvironment. A pKa shift can also occur as a
result of electrostatic interactions: vicinal positive charges
generally make protonation more difficult and deprotonation
easier.
Once these points are made clear, it is not a surprise to see
that 1-pyrenecarboxylic acid (2a) and 1-pyrenesulfonic acid
(2b) had their pKa increased to ∼7 in Figure 1b. With a large
hydrophobic group, these acid cofactors have a strong driving
force to enter the matching hydrophobic binding site within
MINP(1a). This hydrophobic effect (on the pKa) apparently
exceeded the electrostatic interactions of the ammonium head
groups of the cross-linked surfactants that otherwise would
decrease the pKa of the acids.49
If the pKa of 1-pyrenecarboxylic acid (2a) and 1-
pyrenesulfonic acid (2b) indeed shifted to ∼7, the small
degree of hydrolysis under basic conditions (up to pH 9−10)
should be caused by the small amount of the protonated
cofactor in the MINP binding site. The crossover between the
9
10
11
12
13
14
15
16
17
4
48
64
4
3
3
1
2
1
1
NINP
NINP
CTAB
0.5
0.5
-
5
<5
a
Reactions were performed in duplicate with the yields determined by
1H NMR spectroscopy using 1,4-dibromobenzene as an internal
standard. [3a] = 1.0 mM. [2a−d] = 100 μM. [MINP] = [NINP] = 50
μM. [CTAB] = 2.5 mM.
amount of DVB used in the MINP preparation was observed
(entries 1−6). The micelle of 4 could solubilize up to 1 equiv
of DVB relative to the surfactant. MINP(1a) prepared with
DVB/surfactant = 0.5:1 was found to give a better yield than
those with either no DVB or the maximum amount of DVB.
The rigidity of a cross-linked polymer such as MINP is
determined by the cross-linking density. A large amount of
DVB (the core cross-linker) is expected to increase the
integrity of the polymer network and prevent the imprinted
site from collapsing after the removal of the template. A smaller
amount of DVB, on the other hand, should make the polymer
network more flexible and can facilitate the binding and release
of the substrate and product. The medium level of DVB
apparently represented a good balance between the two.
We also performed several control experiments (Table 2,
entries 13−16). MINP in the absence of an acid cofactor, the
acid cofactors alone, the nonimprinted nanoparticle (NINP)
by itself, or the cofactors in the presence of NINP showed
negligible activity. These results demonstrate that the acid
cofactor had to work together with the MINP to cause the
hydrolysis, supporting our design hypothesis. The acid cofactor
(2a) was not active in “normal” cationic micelles of
cetyltrimethylammonium bromide (CTAB) (Table 2, entry
17). Apparently, the non-cross-linked micelle could not
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position the acid in the same way as our MINP that had the
specific binding pockets for both the substrate and the acid
cofactor.
3a at 85 °C after 6 h (Table 2, entry 2). The yield increased to
77% when MINP(1d) was used (Table 3, entry 3). The deeper
pocket also helped 1-pyrenesulfonic acid (2b), increasing the
yield from 71% to 86% (compare entry 5 of Table 2 and entry
4 of Table 3).
Another support for the binding-induced catalysis was the
proper matching between the imprinted pocket and the acid
cofactor required for the hydrolysis. MINP(1a) had a pyrene-
shaped pocket. With this MINP, the pyrenyl-based acids (2a or
2b) gave a significantly higher yield in the hydrolysis than the
smaller naphthyl-based acids (2c or 2d) (Table 2, entries 2 and
5 vs 7 and 8). The results were in line with the binding data in
Table 1, which showed a weaker but measurable binding for
the smaller (sodium salt of) 2c.
When MINP(1b) was used, the trend reversed, with the
naphthyl-based acids (2c or 2d) giving much higher yields
than the pyrenyl-based acids (2a or 2b) (Table 2, entries 9−
12). In fact, the mismatched 2a or 2b was completely inactive,
affording a yield similar to our negative controls. These results
once again were in agreement with the ITC data that showed
negligible binding between MINP(1b) and 2a (Table 1)
because of the larger size of the acids.
For both MINP(1a) and MINP(1b), sulfonic acids (2b and
2d) were found to be more active than the corresponding
carboxylic acids (2a and 2c). Because the reaction was
performed under neutral conditions, the trend was in line with
the observation in Figure 1b.
Fine-Tuning of the Catalytic Active Site. The facile
preparation of the MINP allowed us to tune the catalytic active
site in multiple ways using readily synthesized template
molecules. Template 1c had a linear space holder for the
acid cofactor. Our initial idea was to fine-tune the distance
between the acid group and the acetal oxygen through acid
cofactors 2e and 2f with different chain lengths. The chain
length, however, made little difference in the hydrolysis (Table
3, entries 1 and 2).
How could a deeper active site help the hydrolysis? Because
binding was essential to the catalysis based on our earlier
studies, we measured the binding constants of MINP(1d) for
the acid cofactors (2a and 2b). The numbers (Table 4, entries
2−3) turned out more than doubled those by MINP(1a)
(Table 1, entries 2−3). We attributed the enhanced binding to
a more hydrophobic binding pocket. The interfacial area of a
micelle is significantly more polar than the nonpolar core
because of increased water exposure. Moving the binding
pocket deeper into the core should enhance its hydrophobicity.
In addition to changing the depth of the active site, we could
also tune the number of acid cofactors using templates 1e and
1f, whose syntheses are shown in Schemes 3 and 4,
respectively. We did not use a double-pyrenyl design here
because too large a template could be trapped permanently
inside a cross-linked micelle.50 Of the two templates, 1e
matched substrate 3a in structure better than 1f, which had a
1,3-phenylene where the 1,3-dioxolane of the substrate was
expected to reside.
Our ITC binding data indicated that micellar imprinting
worked well for these templates as well. Both 1e and 1f were
bound by their MINPs with a large binding constant, >107 M−1
(Table 4, entries 4 and 7). In addition, the binding data for the
(sodium salts of the) acid cofactors fit well to a binding model
with two sequential binding sites. The two bindings seemed
independent as the binding constant was quite similar.
Most interestingly, even though the double-acid design
improved the hydrolysis consistently, the improvement was
much more pronounced with the weaker acid. The end result
was that carboxylic acid 2c became equally effective for the
catalysis as sulfonic acid 2d, with the two giving essentially the
same yield (compare entries 5−8 of Table 3 with entries 11−
12 of Table 2).
Table 3. Hydrolysis of 3a catalyzed by MINP-Acid Cofactor
a
Complexes
How could a carboxylic acid catalyze acetal hydrolysis as
well as a much stronger sulfonic acid? Because this only
happened in the double-acid catalyst, a simple answer could be
a higher probability for both the carboxylic acids to stay
protonated in the MINP-cofactor complex. When the acid
becomes highly acidic, it might be quite difficult for the
sulfonic acid cofactors to stay both protonated. If this is indeed
the case, the benefit of having two catalytic groups would be
lost.
Table 3 (entries 5−8) also showed that although 1e seemed
to give a slightly better yield than 1f, the difference was
statistically insignificant. This was good news because a perfect
matching between the space holder and the substrate was not
necessary.
These artificial enzyme−cofactor complexes displayed
enzymelike kinetics. Figure 2 shows the Michaelis−Menten
plot for MINP(1e) + 2c, one of our most active catalysts.
Nonlinear least-squares curve fitting yielded a Vmax value of
0.45 0.02 μM/min, a Km of 389 52 μM, and a kcat of 22.6
× 10−3 min−1. The catalytic efficiency (kcat/Km) was 57.9 M−1
min−1.
Substrate Selectivity of MINP Catalysts. With a
specifically shaped active site, our MINP should hydrolyze
acetals in a selective manner. For this purpose, we used
MINP(1d), with a deeper substrate binding site, to hydrolyze a
entry
MINP
DVB/4
acid cofactor
yield (%)
1
2
3
4
5
6
7
8
MINP(1c)
MINP(1c)
MINP(1d)
MINP(1d)
MINP(1e)
MINP(1e)
MINP(1f)
MINP(1f)
0.5
0.5
0.5
0.5
0.5
0.5
0.5
0.5
2e
2f
68
65
77
86
83
84
80
81
2
2
2
1
2
2
3
1
2a
2b
2c
2d
2c
2d
a
Reactions were performed in duplicate with the yields determined by
1H NMR spectroscopy using 1,4-dibromobenzene as an internal
standard. [3a] = 1.0 mM. [2a−f] = 100 μM. [MINP] = 50 μM.
More interesting results were obtained when we prepared an
MINP using template 1d. This compound had a biphenyl-
derived spacer holder for the substrate. Its linear framework,
longer length in comparison to 1a, and surface anchoring by
the carboxylate were expected to produce a deeper active site,
with the acid catalyst to be bound at the far end.
To our delight, the hydrolysis improved significantly as the
active site became deeper. For example, 1-pyrenecarboxylic
acid (2a) with MINP(1a) gave a 59% yield in the hydrolysis of
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a
Table 4. Binding Data for MINPs Determined by ITC and Fluorescence Titration
guest (sodium salt
of)
−ΔG
b
entry
host
Ka (× 106 M−1
)
(kcal/mol)
−ΔH (kcal/mol)
TΔS (kcal/mol)
N
1
2
3
4
5
6
7
8
9
MINP(1d)
MINP(1d)
MINP(1d)
MINP(1e)
MINP(1e)
MINP(1e)
MINP(1f)
MINP(1f)
MINP(1f)
1d
2a
2b
1e
2c
2d
1f
105 25 (108 60)
8.8 0.45 (9.08 0.22)
6.0 0.25
10.93
9.47
9.24
9.61
7.13 & 7.65
6.87 & 7.28
10.06
7.91 & 7.78
7.73 & 7.66
26.9 0.27
−15.97
−180.14
−30.66
−87.19
0.8 0.003
0.7 0.004
1.4 0.004
1.2 0.003
189.6 1.35
39.9 0.16
96.8 0.45
39.8 2.4 and 6.90 2.58 −32.7 and 0.75
44.1 2.0 and 64.2 2.6
49.0 0.8
50.8 1.4 and 62.7 2.0
54.8 4.0 and 4.35 4.68 −47.07 and 3.32
11.2 0.6
c
0.17 0.02 and 0.41 0.04
0.11 0.01 and 0.22 0.01
24.1 5.2
0.64 0.09 and 0.51 0.04
0.47 0.12 and 0.42 0.09
2
2
c
−37.2 and −56.9
−38.94
−42.9 and −54.9
0.9 0.01
c
2c
2d
2
2
c
a
All the MINPs were prepared with a DVB/surfactant ratio of 0.5:1. Titrations were performed in duplicate using sodium salts of the templates and
acid cofactors in 10 mM HEPES buffer (pH 7.0) with 2% DMSO. The errors between the runs were <10%. The binding constants in parentheses
b
c
were obtained from fluorescence titration. N is the number of binding sites per MINP determined by ITC. Titration data were fitted to a binding
model with two sequential binding sites.
Scheme 3. Synthesis of Compound 1e
number of acetal analogues to understand the selectivity.
Although the double-acid MINPs showed comparable
activities (Table 3), we did not use them because some of
the substrates (e.g., 3e) were expected to enter the
naphthalene-shaped pockets for the acid cofactors. Also, the
binding energy between acid cofactor 2a and MINP(1d) was
extremely large (Table 4, entry 2 for the sodium salt). This was
another important factor to consider. Hydrophobic inter-
actions are nonspecific by itself, and the binding selectivity
comes from the size/shape complementarity. When a large
pyrene cofactor was present, it would be difficult for the
substrates (3a−g) to compete with the cofactor for the pyrene-
shaped binding site.
For the selectivity experiments, because these acetals had
different intrinsic reactivity, we kept the reaction time at 6 h
but varied the temperature so that the background hydrolysis
stayed relatively slow during the reaction time. As shown in
Table 2, the background hydrolysis could be estimated from
the hydrolytic yields obtained in the presence of MINP or the
acid cofactor alone.
Table 5 shows that MINP(1d) + 2a could hydrolyze 3b and
3c quite well. If the ratio of the catalyzed yield over that of the
background was used to estimate the catalytic efficiency, the
order was 3a > 3b > 3c. Thus, all the para-substituted
benzaldehyde acetals were reasonable substrates for the MINP
catalyst, and the one with the lowest intrinsic activity benefited
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Scheme 4. Synthesis of Compound 1f
molecules to pass through. In this case, why did not 3f or 3g
do the same to get in and be converted? In our opinion, this
indeed could happen given the overall flexibility of the cross-
linked structure, as depicted in Scheme 1. Nonetheless, once
2a occupies the catalytic site, even if 3f or 3g could get into the
active site, their residence time, position, and/or orientation
within the active site must not be right for the catalysis to
happen.
CONCLUSIONS
■
Enzymes have inspired generations of chemists with their
fascinating performance. Difficulty in the construction of
complex-shaped active sites with accurately positioned catalytic
groups continues to hamper our design and synthesis of
enzyme mimics. This work demonstrates that with micellar
imprinting, one can quickly construct a substrate-tailored
nanospace in a water-soluble protein-sized organic nano-
particle, with catalytic groups positioned near the reactive
functionality. The size/shape selectivity of the imprinted site
allowed the substrate and the acid cofactor each to occupy its
designed positionan extremely important feature of our
system.
Nature uses unimpressive functional groups for challenging
catalysis, but chemists often have to resort to strong acids and
expensive metals not available in a biological system. The
stronger performance of a deeper active site in our catalytic
hydrolysis was a useful learning, which could provide guidance
to the design of future artificial enzymes. MINP(1e/f)
demonstrated that with a proper design of the active site,
weak carboxylic acids could become as effective as stronger
sulfonic acids.
Figure 2. Michaelis−Menten plot for the hydrolysis of 3a by
MINP(1e) + 2c. The reaction rates were measured in D2O at 85 °C.
[MINP(1e)] = 20 μM. [2c] = 40 μM. Reaction progress was
monitored by 1H NMR spectroscopy using 1,4-dibromobenzene as an
internal standard. The MINPs were prepared with a DVB/surfactant
ratio of 0.5:1.
the most from the catalysis. For the two naphthyl acetals 3d
and 3e, the 2-naphthyl derivative was a much better substrate
than the 1-naphthyl one, even though their background
reactivity was similar. The most likely reason for the selectivity
should be their different shape, with 3e bearing a higher
resemblance to 3a, the substrate that the catalyst was designed
for.
The most interesting selectivity was found in 3f and 3g.
Diethyl acetal 3f is known to hydrolyze faster than 1,3-
dioxolane acetal 3a by over 20 times in solution.41 However,
when catalyzed by MINP(1d) + 2a, 3f hydrolyzed very little
(Table 5), while 3a hydrolyzed easily under identical
conditions (Table 3). The long and narrow pocket generated
from 1d thus seemed to have difficulty accommodating the
acyclic acetals 3f and 3g or at least was poorly suited for
catalysis.
Overall, our method made modulation of the catalytic
activity readily achievable through systematic tuning of the
active site. The resulting artificial enzyme was able to override
intrinsic reactivity of substrates, a feature frequently found in
enzymes but difficult to obtain with synthetic catalysts. As a
cross-linked polymeric nanoparticle, MINP tolerates high
temperature, organic solvents,51 and extreme pH.52 These
features plus the facile construction and modification of the
One might wonder that, if large guests such as 2a or 2b
could get into the far end of the imprinted site within
MINP(1d) according to ITC (Table 4), the active site for the
substrate must be able to “breathe” to let relatively large
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a
Table 5. Hydrolytic Yields of Acetal Analogues by MINP Catalysts
a
[acetal] = 1.0 mM. [2a] = 100 μM. [MINP(1d)] = 50 μM. The MINPs were prepared with a DVB/surfactant ratio of 0.5:1.
128.7, 128.6, 127.2, 127.0, 63.4, 60.7, 52.2. HRMS (ESI) m/z: [M +
H]+ calcd for C17H17NO3, 284.1281; found, 284.1274.
active site make it a very versatile platform for artificial
enzymes.
General Procedure for the Synthesis of Compounds 9−12.
Pyridine (10 mmol, 2.0 equiv) was added to a solution of the
appropriate acid chloride (3.85 mmol, 0.77 equiv) and compound 7
or 8 (5 mmol, 1.0 equiv) in ethyl acetate (40 mL). The reaction
mixture was stirred at 50 °C in an oil bath for 6 h. After the mixture
was cooled to room temperature, the solid precipitate was removed by
filtration and the solvents were removed by rotary evaporation. The
residue was purified by flash chromatography over silica gel using 15:1
dichloromethane/ethyl acetate as the eluent to afford the final
product.
EXPERIMENTAL SECTION
■
General Experimental Methods. All organic solvents and
reagents were of ACS-certified grade or higher grade and were
purchased from commercial suppliers. Chemicals shifts are reported in
parts per million (ppm) relative to residual solvent peaks. Coupling
constants are reported in hertz. Electrospray ionization−high-
resolution mass spectrometry (ESI-HRMS) was performed on an
Agilent QTOF 6540 mass spectrometer with a QTOF detector. Milli-
Q water (18.2 MU; Millipore Co., USA) was used for MINP
preparation and all buffers. DLS data were obtained on a Malvern
Zetasizer Nano ZS at 25 °C. ITC was performed using a MicroCal
VP-ITC Microcalorimeter with Origin 7 software and VPViewer2000
(GE Healthcare, Northampton, MA). Fluorescence spectra were
recorded at ambient temperature on a Varian Cary Eclipse
Fluorescence spectrophotometer.
Compound 9. A white powder (0.68 g, 41%). 1H NMR (400 MHz,
CDCl3): δ 8.19−7.42 (bm, 13H), 6.82−6.71 (m, 1H), 4.48−3.24 (m,
7H). 13C{1H} NMR (100 MHz, CDCl3): δ 169.5, 166.7, 143.8, 132.1,
130.9, 130.5, 130.5, 129.9, 129.1, 128.5, 128.4, 127.4, 126.9, 126.8,
126.4, 126.2, 125.9, 125.7, 124.5, 124.2, 124.1, 123.5, 88.5, 66.5, 52.1,
47.2. HRMS (ESI) m/z: [M + H]+ calcd for C28H21NO4, 436.1543;
found, 436.1542.
Syntheses of substrates 3a−c/f,35 3d,53 surfactant 4,54 cross-linker
5,16 surface ligand 6,16 7,55 and 1335 were reported previously.
Syntheses of substrates 3e56 and 3g57 were adapted from the known
procedures.
1
Compound 10. A white powder (1.18 g, 85%). H NMR (400
MHz, CDCl3): δ 8.12−7.36 (m, 11H), 6.85−6.72 (m, 1H), 4.40−
3.34 (m, 7H). 13C{1H} NMR (100 MHz, CDCl3): δ 168.9, 166.7,
143.7, 133.8, 130.4, 130.0, 129.8, 129.3, 129.2, 128.5, 127.3, 126.7,
126.5, 126.5, 124.9, 124.5, 88.3, 66.5, 52.1, 47.1. HRMS (ESI) m/z:
[M + H]+ calcd for C22H19NO4, 362.1387; found, 362.1391.
Compound 8. A solution of methyl 4′-formyl-[1,1′-biphenyl]-4-
carboxylate (1.2 g, 5 mmol, 1.0 equiv) and 2-aminoethanol (0.32 mL,
5.25 mmol, 1.05 equiv) in toluene (80 mL) was heated to reflux in an
oil bath with a Dean−Stark apparatus to remove water for 7 h. After
the solvent was removed by rotary evaporation, the residue was used
directly in the next step without further purification. A white powder
1
Compound 11. A white powder (1.37 g, 70%). H NMR (400
MHz, CDCl3): δ 8.26−7.70 (m, 15H), 6.85−6.67 (m, 3H), 4.55−
3.38 (m, 7H). 13C{1H} NMR (100 MHz, CDCl3): δ 169.5, 167.0,
145.2, 140.6, 138.9, 132.3, 131.2, 130.8, 130.2, 129.8, 129.2, 128.7,
128.4, 127.6, 127.5, 127.2, 127.0, 126.7, 126.5, 126.3, 126.1, 125.8,
124.7, 124.3, 124.2, 123.8, 89.1, 66.6, 52.2, 47.5. HRMS (ESI) m/z:
[M + H]+ calcd for C34H25NO4, 512.1856; found, 512.1858.
1
(1.10 g, 81%). H NMR (400 MHz, DMSO-d6): δ 8.37 (s, 1H),
8.05−8.03 (m, 2H), 7.88−7.85 (m, 4H), 7.83−7.81 (m, 2H), 4.63 (t,
J = 4 Hz, 1H), 3.86 (s, 3H), 3.68−3.66 (m, 4H) ppm. 13C{1H} NMR
(100 MHz, DMSO-d6): δ 166.0, 161.2, 143.9, 140.6, 136.1, 129.9,
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1
Compound 12. A white powder (0.56 g, 33%). H NMR (400
MHz, CDCl3): δ 7.97 (d, J = 8.4 Hz, 2H), 7.51 (d, J = 8.4 Hz, 2H),
6.23 (s, 1H), 4.07−3.92 (m, 2H), 3.85 (s, 3H), 3.81−3.75 (m, 1H),
3.34−3.29 (m, 1H), 2.88−2.84 (m, 2H), 1.76−1.71 (m, 2H), 1.31−
1.17 (m, 18H), 0.80 (t, J = 6.4 Hz, 3H). 13C{1H} NMR (100 MHz,
CDCl3): δ 166.6, 142.9, 130.7, 129.8, 126.7, 90.1, 66.0, 52.2, 51.4,
46.2, 31.9, 29.6, 29.5, 29.4, 29.3, 29.3, 29.1, 28.4, 23.0, 22.7, 14.2.
HRMS (ESI) m/z: [M + H]+ calcd for C23H37NO5S, 440.2465;
found, 440.2465.
insoluble solid was removed by filtration and the solvents were
removed by rotary evaporation. The residue was purified by flash
chromatography over silica gel using 1:5 methanol/dichloromethane
1
as the eluent to afford an orange powder (0.18 g, 91%). H NMR
(400 MHz, DMSO-d6): δ 8.53−8.42 (m, 2H), 8.38−8.21 (m, 2H),
8.19−7.57 (m, 9H), 7.54−7.03 (m, 6H), 3.49−3.17 (m, 4H).
13C{1H} NMR (100 MHz, 323 K, DMSO-d6): δ 170.6, 166.9, 142.0,
138.2, 137.6, 133.2, 132.5, 128.8, 128.4, 128.1, 127.2, 126.5, 126.3,
125.7, 125.5, 125.4, 125.0, 45.3. HRMS (ESI) m/z: [M − H]− calcd
for C32H24N2O4, 499.1663; found, 499.1655.
General Procedure for the Synthesis of Compounds 1a−d.
The appropriate ester (9−12, 0.5 mmol, 1.0 equiv) was dissolved in a
2:1 methanol/ tetrahydrofuran (THF) mixture (2 mL), to which 1
mL of aqueous NaOH solution (2 N) was added. The reaction
mixture was stirred at room temperature for 5 h, followed by the
addition of 2 mmol of sodium bicarbonate. After 10 min, the insoluble
solid was removed by filtration and the solvents were removed by
rotary evaporation. The residue was purified by flash chromatography
over silica gel using 1:5 methanol/dichloromethane as the eluent to
afford the final product.
Compound 15. A mixture of (4-(tert-butoxycarbonyl)phenyl)-
boronic acid (0.15 g, 0.70 mmol, 1.67 equiv), 2-iodo-1,3-
dimethoxybenzene (0.11 g, 0.42 mmol, 1.0 equiv), and bis-
(triphenylphosphine)palladium(II) chloride (0.05 g, 0.07 mmol,
0.17 equiv) in dry THF (20 mL) and saturated NaHCO3 solution
(25 mL) was heated at 70 °C in an oil bath under nitrogen overnight.
The organic solvent was removed by rotary evaporation, and the
aqueous solution was extracted with dichloromethane (3 × 30 mL).
The combined organic solution was washed with brine (3 × 20 mL)
and dried over Na2SO4 and filtered. Dichloromethane was removed
by rotary evaporation, and the residue was purified by column
chromatography over silica gel using 1:5 ethyl acetate/hexane as the
1
Compound 1a. A white powder (0.21 g, 93%). H NMR (400
MHz, DMSO-d6): δ 13.05 (br s, 1H), 8.50−7.40 (bm, 13H), 6.87−
6.60 (m, 1H), 4.35−3.30 (m, 4H). 13C{1H} NMR (100 MHz,
DMSO-d6): δ 167.9, 167.2, 143.9, 139.4, 131.5, 131.1, 130.7, 130.1,
129.8, 129.5, 128.8, 128.4, 127.8, 127.1, 126.9, 126.4, 126.1, 125.8,
124.8, 124.5, 123.8, 123.7, 88.7, 66.3, 47.4. HRMS (ESI) m/z: [M +
H]+ calcd for C27H19NO4, 422.1387; found, 422.1391.
1
eluent to give a white powder (0.08 g, 62%). H NMR (400 MHz,
CDCl3): δ 8.06 (d, J = 8.0 Hz, 2H), 7.42 (d, J = 8.0 Hz, 2H), 7.32 (t, J
= 8.0 Hz, 1H), 6.67 (d, J = 8.0 Hz, 2H), 3.74 (s, 6H), 1.63 (s, 9H).
13C{1H} NMR (100 MHz, CDCl3): δ 165.9, 157.6, 139.0, 131.0,
130.4, 129.3, 128.9, 118.8, 104.3, 80.7, 55.9, 28.4. HRMS (ESI) m/z:
[M + H]+ calcd for C19H22O4, 315.1591; found, 315.1571.
1
Compound 1b. A white powder (0.18 g, 97%). H NMR (400
MHz, DMSO-d6): δ 13.00 (br s, 1H), 8.17−7.15 (m, 10H), 6.92−
6.90 (m, 1H), 6.51 (s, 1H), 4.21−3.57 (m, 4H). 13C{1H} NMR (100
MHz, DMSO-d6): δ 167.6, 167.1, 143.9, 134.1, 133.1, 131.1, 129.7,
129.5, 128.8, 128.5, 127.3, 127.2, 126.9, 126.5, 125.3, 124.6, 88.3,
66.4, 47.2. HRMS (ESI) m/z: [M + H]+ calcd for C21H17NO4,
348.1230; found, 348.1229.
Compound 16. 1 mL of BBr3 in dry dichloromethane (1 M) was
slowly added to a solution of 15 (0.10 g, 0.32 mmol, 1.0 equiv) in dry
dichloromethane (30 mL) at −78 °C. The reaction mixture was
warmed to room temperature and stirred for 15 h. After the reaction
was complete, the mixture was cooled to 0 °C and quenched by the
dropwise addition of water, followed by acidification with HCl (2 N),
and the organic phase was separated. The aqueous phase was
extracted with EtOAc (3 × 20 mL). The combined organic layers
were dried over Na2SO4, filtered, and removed by rotary evaporation.
The residue was purified over silica gel using 2:5 ethyl acetate/hexane
1
Compound 1d. A white powder (0.21 g, 80%). H NMR (400
MHz, DMSO-d6): δ 12.98 (br s, 1H), 8.38−7.71 (m, 15H), 7.00−
6.58 (m, 3H), 4.35−3.32 (m, 4H). 13C{1H} NMR (100 MHz,
DMSO-d6): δ 167.9, 167.2, 143.9, 139.5, 131.6, 131.2, 130.7, 130.2,
130.1, 129.5, 128.8, 128.5, 127.8, 127.3, 127.1, 126.9, 126.9, 126.8,
126.5, 126.1, 125.9, 125.6, 124.9, 124.5, 123.8, 123.7, 88.8, 66.3, 47.5.
HRMS (ESI) m/z: [M + H]+ calcd for C33H23NO4, 498.1700; found,
498.1701.
1
as the eluent to give a white powder (0.05 g, 68%). H NMR (400
MHz, CD3OD): δ 8.02 (d, J = 8.0 Hz, 2H), 7.49 (d, J = 8.0 Hz, 2H),
6.97 (t, J = 8.0 Hz, 1H), 6.42 (d, J = 8.0 Hz, 2H). 13C{1H} NMR
(100 MHz, CD3OD): δ 170.2, 156.6, 141.7, 132.4, 130.1, 129.9,
129.6, 117.0, 108.2. HRMS (ESI) m/z: [M − H]− calcd for
C13H10O4, 229.0506; found, 229.0503.
1
Compound 1c. A white powder (0.08 g, 34%). H NMR (400
MHz, CDCl3): δ 8.13 (d, J = 8.4 Hz, 2H), 7.63 (d, J = 8.4 Hz, 2H),
6.33 (s, 1H), 4.16−4.00 (m, 2H), 3.90−3.81 (m, 1H), 3.44−3.35 (m,
1H), 2.99−2.90 (m, 2H), 1.87−1.77 (m, 2H), 1.45−1.15 (m, 18H),
0.87 (t, J = 6.8 Hz, 3H). 13C{1H} NMR (100 MHz, CDCl3): δ 171.2,
144.0, 130.6, 129.9, 126.9, 110.1, 90.2, 66.2, 51.6, 46.3, 32.1, 29.7,
29.6, 29.5, 29.4, 29.2, 28.5, 23.1, 22.8, 14.3. HRMS (ESI) m/z: [M −
H]− calcd for C22H35NO5S, 424.2163; found, 424.2172.
Compound 14. 2-Naphthoyl chloride (1.08 g, 5.68 mmol, 4.0
equiv) in dry dichloromethane (15 mL) was slowly added to a
solution of 13 (0.50 g, 1.42 mmol, 1.0 equiv) and triethylamine (0.79
mL, 5.68 mmol, 4.0 equiv) in dry dichloromethane (15 mL) at 0 °C
under nitrogen. The reaction mixture was warmed to room
temperature and stirred for 12 h. After the mixture was washed
with saturated NaHCO3, brine, and dried (Na2SO4), the solvent was
removed by rotary evaporation. The residue was purified by column
chromatography over silica gel using 1:1 hexane/ethyl acetate as the
eluent to give an orange powder (0.55 g, 76%). 1H NMR (600 MHz,
DMSO-d6): δ 9.00−8.78 (m, 2H), 8.53−8.45 (m, 2H), 8.02−7.95
(m, 10H), 7.75 (d, J = 6.0 Hz, 1H), 7.63−7.57 (m, 4H), 3.87−3.51
(m, 7H). 13C{1H} NMR (100 Hz, DMSO-d6): δ 166.6, 165.8, 137.3,
134.2, 134.1, 131.9, 131.2, 130.3, 129.9, 129.4, 128.8, 127.8, 127.6,
127.5, 126.7, 124.2, 52.3, 49.5, 46.4. HRMS (ESI) m/z: [M + H]+
calcd for C33H26N2O4, 515.1971; found, 515.1959.
Compound 17. 2-Naphthoyl chloride (0.18 g, 0.95 mmol, 2.2
equiv) in dry dichloromethane (10 mL) was slowly added to a
solution of 16 (0.10 g, 0.43 mmol, 1.0 equiv) and triethylamine (0.13
mL, 0.95 mmol, 2.2 equiv) in dry dichloromethane (10 mL) at 0 °C
under nitrogen. The reaction mixture was warmed to room
temperature and stirred for 16 h. After the mixture was washed
with saturated NaHCO3, brine, and dried (Na2SO4), the solvent was
removed by rotary evaporation. The residue was purified by column
chromatography over silica gel using 5:1 hexane/ethyl acetate as the
eluent to give an orange powder (0.14 g, 61%). 1H NMR (600 MHz,
CDCl3): δ 8.46 (s, 2H), 7.91−7.82 (m, 9H), 7.60−7.53 (m, 8H), 7.37
(d, J = 8.0 Hz, 2H). 13C{1H} NMR (150 MHz, CDCl3): δ 166.8,
164.9, 149.4, 137.4, 135.9, 132.5, 132.1, 131.2, 130.1, 129.6, 129.5,
129.4, 128.8, 128.5, 127.9, 126.9, 126.8, 126.2, 125.3, 120.9. HRMS
(ESI) m/z: [M − H]− calcd for C35H22O6, 537.1344; found,
537.1337.
Compound 1f. Compound 17 (0.10 g, 0.18 mmol, 1.0 equiv) and
sodium bicarbonate (0.017 g, 0.20 mmol, 1.1 equiv) were dissolved in
10 mL of methanol, and the reaction mixture was stirred overnight at
room temperature. After the reaction mixture was concentrated by
rotary evaporation, the residual white powder (0.09 g, 89%) was used
in MINP preparation without further purification.
Preparation of MINP. A typical procedure is as follows: to a
micellar solution of surfactant 4 (10.2 mg, 0.02 mmol) in H2O (2.0
mL) were added DVB (2.8 μL, 0.02 mmol), DMPA in DMSO (10 μL
of a 12.8 mg/mL solution, 0.0005 mmol), and the relative template in
Compound 1e. Compound 14 (0.20 g, 0.39 mmol, 1.0 equiv) was
dissolved in a 2:1 methanol/THF mixture (2 mL), to which 1 mL of
aqueous NaOH solution (2 N) was added. The reaction mixture was
stirred at room temperature for 5 h, followed by the addition of
sodium bicarbonate (0.17 g, 2.0 mmol, 5.13 equiv). After 10 min, the
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DMSO (10 μL of a 0.04 M solution, 0.0004 mmol). The mixture was
subjected to ultrasonication for 10 min before cross-linker 5 (4.1 mg,
0.024 mmol), CuCl2 in H2O (10 μL of 6.7 mg/mL solution, 0.0005
mmol), and sodium ascorbate in H2O (10 μL of 99 mg/mL solution,
0.005 mmol) were added. After the reaction mixture was stirred
slowly at room temperature for 12 h, compound 6 (15.9 mg, 0.06
mmol), CuCl2 in H2O (10 μL of 6.7 mg/mL solution, 0.0005 mmol),
and sodium ascorbate in H2O (10 μL of 99 mg/mL solution, 0.005
mmol) were added. The mixture was stirred at room temperature for
another 6 h, purged with nitrogen for 15 min, sealed with a rubber
stopper, and irradiated in a Rayonet reactor for 12 h. The reaction
mixture was then poured into acetone (8 mL). The precipitate was
collected by centrifugation and washed with a mixture of acetone/
water (5 mL/1 mL) three times, followed by methanol/acetic acid (5
mL/0.1 mL) three times. The solid was then rinsed twice with
acetone (5 mL) and dried in air to afford the final MINPs as an off-
white powder. Typical yields were >80%.
Determination of Binding Constants by ITC. In general, a
solution of an appropriate guest in 10 mM N-(2-hydroxyethyl)-
piperazine-N′-ethanesulfonic acid (HEPES) buffer (pH 7.0) with 2%
DMSO at 298 K (DMSO was added to help the solubility of the
appropriate guest) was injected in equal steps into 1.43 mL of the
corresponding MINP in the same solution. The top panel shows the
raw calorimetric data. The area under each peak represents the
amount of heat generated at each ejection and is plotted against the
molar ratio of the MINP to the guest. The solid line is the best fit of
the experimental data to the sequential binding of N equal and
independent binding sites on the MINP. The heat of dilution for the
substrate, obtained by adding the substrate to the buffer, was
subtracted from the heat released during the binding. Binding
parameters were autogenerated after curve fitting using Microcal
Origin 7. All titrations were performed in duplicate, and the errors
between the runs were <10% using sodium salts of the templates and
acid cofactors.
AUTHOR INFORMATION
■
Corresponding Author
Yan Zhao − Department of Chemistry, Iowa State University,
Ames, Iowa 50011-3111, United States; orcid.org/0000-
0003-1215-2565; Email: zhaoy@iastate.edu
Authors
Ishani Bose − Department of Chemistry, Iowa State
University, Ames, Iowa 50011-3111, United States
Shixin Fa − Department of Chemistry, Iowa State University,
Ames, Iowa 50011-3111, United States
Complete contact information is available at:
https://pubs.acs.org/10.1021/acs.joc.0c02519
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
■
We thank the NIGMS (R01GM138427) for financial support
of this research.
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ASSOCIATED CONTENT
■
sı
* Supporting Information
The Supporting Information is available free of charge at
https://pubs.acs.org/doi/10.1021/acs.joc.0c02519.
Characterization of MINPs, ITC and fluorescence
titration curves, and NMR spectra of key compounds
(PDF)
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