Molecularly Imprinted Synthetic Glucosidase for the Hydrolysis of Cellulose in Aqueous and Nonaqueous Solutions

Article abstract of DOI:10.1021/jacs.1c01352

Molecular imprinting is a powerful and yet simple method to create multifunctional binding sites within a cross-linked polymer network. We report a new class of synthetic glucosidase prepared through molecular imprinting and postfunctionalization of cross-linked surfactant micelles. These catalysts are protein-sized water-soluble nanoparticles that can be modified in multiple ways. As their natural counterparts, they bind a glucose-containing oligo- or polysaccharide. They contain acidic groups near the glycosidic bond to be cleaved, with the number and distance of the acid groups tuned systematically. Hydrolysis of cellulose in a key step in biomass conversion but is hampered by the incalcitrance of the highly crystalline cellulose fibers. The synthetic glucosidases are shown to hydrolyze cellobiose and cellulose under a variety of conditions. The best catalyst, with a biomimetic double acid catalytic motif, can hydrolyze cellulose with one-fifth of the activity of commercial cellulases in aqueous buffer. As a highly cross-linked polymeric nanoparticle, the synthetic catalyst is stable at elevated temperatures in both aqueous and nonaqueous solvents. In a polar aprotic solvent/ionic liquid mixture, it hydrolyzes cellulose several times faster than commercial cellulases in aqueous buffer. When deposited on magnetic nanoparticles, it retains 75% of its activity after 10 cycles of usage.

Full text of DOI:10.1021/jacs.1c01352

pubs.acs.org/JACS
Article
Molecularly Imprinted Synthetic Glucosidase for the Hydrolysis of
Cellulose in Aqueous and Nonaqueous Solutions
Xiaowei Li, Milad Zangiabadi, and Yan Zhao*
Cite This: J. Am. Chem. Soc. 2021, 143, 5172−5181
Read Online
sı
*
Metrics & More
Article Recommendations
Supporting Information
ACCESS
ABSTRACT: Molecular imprinting is a powerful and yet simple
method to create multifunctional binding sites within a cross-
linked polymer network. We report a new class of synthetic
glucosidase prepared through molecular imprinting and post-
functionalization of cross-linked surfactant micelles. These
catalysts are protein-sized water-soluble nanoparticles that can be
modified in multiple ways. As their natural counterparts, they bind
a glucose-containing oligo- or polysaccharide. They contain acidic groups near the glycosidic bond to be cleaved, with the number
and distance of the acid groups tuned systematically. Hydrolysis of cellulose in a key step in biomass conversion but is hampered by
the incalcitrance of the highly crystalline cellulose fibers. The synthetic glucosidases are shown to hydrolyze cellobiose and cellulose
under a variety of conditions. The best catalyst, with a biomimetic double acid catalytic motif, can hydrolyze cellulose with one-fifth
of the activity of commercial cellulases in aqueous buffer. As a highly cross-linked polymeric nanoparticle, the synthetic catalyst is
stable at elevated temperatures in both aqueous and nonaqueous solvents. In a polar aprotic solvent/ionic liquid mixture, it
hydrolyzes cellulose several times faster than commercial cellulases in aqueous buffer. When deposited on magnetic nanoparticles, it
retains 75% of its activity after 10 cycles of usage.
introduce catalytic groups that could not be introduced directly
through functional monomers during the initial imprinting. In
recent years, our group has developed a method to molecularly
imprint surfactant micelles.17 The nanoconfinement of the
polymerization and cross-linking within the surface-cross-linked
micelle yields an extraordinary templating effect,18 with the
imprint/nonimprint ratio in binding (i.e., imprinting factor)
frequently reaching hundreds19 and sometimes 10000.20 The
addition,18 removal,18 and shift21 of a single methyl (or
methylene) group in the guest can be distinguished by the so-
called molecularly imprinted nanoparticles (MINPs). Catalytic
groups can be installed inside the imprinted pockets to afford
highly selective, enzyme-mimetic catalysts.22−25
In this work, we report MINP-based synthetic glucosidases for
the hydrolysis of cellulose, taking advantage of the high fidelity
of micellar imprinting and facile postfunctionalization enabled
by the accessibility of the imprinted sites. Cellulose constitutes
35−50% of lignocellulosic biomass and is an abundant carbon-
neutral natural resource.26−28 Whether for chemical or fuel
production, it first has to be depolymerized into soluble
monomeric and oligomeric sugars, usually through catalytic
INTRODUCTION
■
The extraordinary catalytic efficiency and selectivity of enzymes
have motivated generations of scientists to develop synthetic
mimics with similar capabilities.1−3 Although chemists rightfully
argue that enzymes are “not different, just better” than synthetic
catalysts,4 artificial enzymes created on purely synthetic or
seminatural platforms generally do not match their natural
counterparts in performance.1−3 The deficiency of artificial
enzymes has many reasons. Apart from a detailed understanding
of enzymatic activity, a large roadblock is the lack of suitable
synthetic strategies to construct multifunctional active sites with
accurately positioned catalytic groups for substrates with
complex structures and shapes.
A powerful method to create multifunctional binding sites for
molecules of many different sizes is molecular imprinting.5−7 In
its traditional embodiment, template molecules are mixed with a
large amount of cross-linkers and functional monomers (FMs)
that can interact with the templates by noncovalent or reversible
covalent bonds. Free radical polymerization, followed by
template removal, yields a highly cross-linked polymeric
network with embedded imprinted sites complementary to the
templates ideally in size, shape, and distribution of functional
groups. Owing to simplicity of the preparation and broad utility
to molecules of different types, molecularly imprinted polymers
(MIPs) have found numerous applications in biomedical
research8,9 and catalysis.10−16
Received: February 3, 2021
Published: March 24, 2021
To create a catalytic active site similar to those found in
enzymes, one needs not only high fidelity in the imprinting
process but also abilities to postmodify the imprinted site to
https://doi.org/10.1021/jacs.1c01352
J. Am. Chem. Soc. 2021, 143, 5172−5181
© 2021 American Chemical Society
5172

Journal of the American Chemical Society
pubs.acs.org/JACS
Article
Scheme 1. General Procedure for the MINP Preparation from Mixed Micelle of 1 and 2 Containing DVB and DMPA
hydrolysis. Although natural enzymes (i.e., cellulases) exist for
this process,29 the easily denatured protein structure limits the
operating window of these enzymes and makes their recycling
difficult. In recent years, great efforts have been devoted toward
improving the stability of cellulases30,31 but high temperatures
and nonaqueous solvents are inherently difficult for any enzyme-
based catalysts. In contrast, our polymer-based synthetic
glucosidases were shown to tolerate elevated temperatures,
nonaqueous solution, and extreme pH conditions. Additionally,
they could be easily “clicked” onto magnetic nanoparticles to be
converted into a heterogeneous catalyst that retained 75% of its
activity after 10 cycles of usage.
template, and a second round click reaction between 3 and the
residual alkynes on the micelle installs a layer of hydrophilic
ligand on the surface. The surface ligands enable the resulting
MINPs to be purified by simple precipitation and washing with
organic solvents, with the template molecules removed during
the process. A surfactant/template ratio of 50 is commonly used
to give an average of one binding site per nanoparticle, since
MINP contains ∼50 cross-linked surfactants.17
The most challenging aspect of building a synthetic enzyme is
probably accurate positioning of multiple catalytic groups near
the bonds to be transformed.1−3 For this purpose, we designed
template 5, consisting of a glucose and an aglycon containing a
reversible imine bond (Scheme 2). It reacted with vinyl-
phenylboroxole 4 in situ in the micellar solution to afford
amphiphilic, anionic template−FM complex 6, which was
stabilized by the cationic micelle.36,38−40 The resulting MINP-
(5) had the template covalently polymerized into the micellar
core due to the vinyl group on the template. The imine bound
was hydrolyzed by 6 N hydrochloric acid at 95 °C to give MINP-
CHO(5). The aldehyde group in the active site was further
derivatized with amino acids 8a−i via reductive amination
following previously established protocols39,41,42 to afford
MINP (5+8a−i) as our synthetic glucosidase. The cross-linking
and formation of MINPs generally can be characterized by a
combination of 1H NMR spectroscopy (Figures S1−2),
dynamic light scattering (Figures S3−4), and transmission
electron microscopy (Figure S5).
RESULTS AND DISCUSSION
■
Preparation of Synthetic Glucosidase and Its Hydrol-
ysis of Cellobiose. Carbohydrates can be bound by boronic
acids32−35 or boroxole36−39 via reversible boronate bonds
formed with specific 1,2- and 1,3-diols on the sugar. The
glycosidic linkages in cellulose are essentially acid-sensitive
acetals. Thus, to hydrolyze cellulose, a synthetic cellulase needs
to bind a portion of the polymer chain and have an acidic group
right next to the glycosidic bond to be cleaved.
Our construction of such a catalyst was based on molecular
imprinting5−7 in cross-linked micelles (Scheme 1).17,35,39 This
method starts with spontaneous formation of mixed micelles of
1 and 2 containing a template molecule, divinylbenzene (DVB, a
free radical cross-linker), and 2,2-dimethoxy-2-phenylacetophe-
none (DMPA, a photoinitiator). Cu(I) catalysts are used to
cross-link the surface of the micelle by the highly efficient
alkyne−azide “click” reaction, facilitated additionally by the
proximity of the reactive groups. UV-induced free-radical
polymerization then cross-links the micelle core around the
As shown in Scheme 2, MINP (5+8a−i) has a boroxole group
in the active site to bind the terminal glucose of cellobiose (or
cellulose), with an acid positioned near the glycosidic bond to
catalyze the hydrolysis. Boronate 6 has two free hydroxyl groups,
and its amphiphilicity helps itself to be anchored near the surface
5173
https://doi.org/10.1021/jacs.1c01352
J. Am. Chem. Soc. 2021, 143, 5172−5181

Journal of the American Chemical Society
pubs.acs.org/JACS
Article
Scheme 2. Preparation of MINP-Based Synthetic Glucosidase, with a Schematic Representation of the Active Site Functionalized
a
with 8i and a Cellobiose Bound by the Boroxole Group Introduced through FM 4
a
The surface ligands are omitted for clarity.
of the micelle: a feature important to the removal of the template
to vacate the imprinted site after micellar imprinting.17 In the
final catalyst, it is also important to keep the active site close to
the surface so that the unbound glucose residues of the substrate
can stay in solution while the terminal glucose engages in
reversible boronate formation with the boroxole inside the active
site.
Amino acids 8a−i allowed the introduction of different acids
in the active site, with its flexibility and distance to the glycosidic
bond tuned systematically. To our delight, the resulting
MINP(5+8a−i) hydrolyzed cellobiose, our model substrate, in
a 21−64% yield at 60 °C and in pH 6 buffer (Table 1, entries 1−
7). It is interesting that 8a, which was similar in dimension to the
part of the aryl aglycon of 5 removed by imine hydrolysis,
yielded a less active catalyst than 8d. A tightly fit active site thus
seems less efficient than one with some flexibility. When the
number of methylene groups (n = 3) stayed the same with 8d,
8h, and 8i, the most acidic sulfonic acid gave the fastest
hydrolysis.
The above method is not limited to a monoacidic design. A
huge number of glycosidases exist in nature to cleave the
glycosidic bonds of oligo- and polysaccharides.43 A highly
conserved feature of these enzymes is a pair of carboxylic acids in
the active site, a remarkably simple catalytic motif. The
hydrolytic mechanism involves one carboxylic acid as a general
acid to protonate the glycosidic oxygen. For inverting
glycosidases, the other carboxyl 7−11 Å apart, in the
deprotonated form, is a general base to deprotonate the
Table 1. Hydrolysis of Cellobiose Catalyzed by MINPs in 10
mM MES Buffer (pH 6)
a
Entry
catalysts
temp. (°C)
yield (%)
1
2
3
4
5
6
7
8
MINP(5+8a)
MINP(5+8b)
MINP(5+8c)
MINP(5+8d)
MINP(5+8e)
MINP(5+8h)
MINP(5+8i)
MINP(5+8i)
MINP(7+8c)
MINP(7+8d)
MINP(7+8e)
MINP(7+8f)
MINP(7+8g)
MINP(7+8f)
NINP with 8f
8i
60
60
60
60
60
60
60
90
60
60
60
60
60
90
60
60
60
60
42
22
25
51
21
57
64
74
0
17
54
94
77
98
0
6
4
4
7
4
4
8
6
9
10
11
12
13
14
15
16
17
18
2
6
4
8
1
b
∼2
∼1
0
8f
none
a
Reactions were performed with 0.2 mM of cellobiose and 20 μM of
catalysts in 1.0 mL MES buffer (10 mM, pH = 6.0) for 24 h. Yields
were determined by LC-MS using standard curves generated from
b
authentic samples. NINP was nonimprinted nanoparticle prepared
with 1 equiv FM 4 but without any template.
5174
https://doi.org/10.1021/jacs.1c01352
J. Am. Chem. Soc. 2021, 143, 5172−5181

Journal of the American Chemical Society
pubs.acs.org/JACS
Article
a
Table 2. ITC Binding Data for Sugar Guests by MINPs
Entry
MINP
guest
pH
Ka(×103 M−1
)
ΔG (kcal/mol)
N
1
2
3
4
5
6
7
8
9
MINP-CHO(5)
MINP(5+8i)
MINP-CHO(7)
MINP(7+8f)
MINP(5+8i)
MINP(5+8i)
MINP(7+8f)
MINP(7+8f)
glucose
glucose
glucose
glucose
glucose
cellobiose
glucose
cellobiose
glucose
7.4
7.4
7.4
7.4
6.0
6.0
6.0
6.0
7.4
8.85 0.68
9.07 0.86
−5.38
−5.40
−5.78
−5.94
−5.28
−5.12
−5.72
1.03 0.03
0.83 0.09
1.01 0.03
0.91 0.03
0.95 0.07
0.78 0.08
0.79 0.09
17.40 1.92
22.60 1.07
7.52 0.79
5.71 0.51
15.70 1.32
6.98 1.52
−5.24
1.06 0.17
b
b
b
b
NINP
<0.05
a
The FM/template ratio in the MINP synthesis was 1:1 unless otherwise indicated. The cross-linkable surfactants were a 3:2 mixture of 1 and 2.
The titrations were performed in 10 mM HEPES buffer at pH 7.4 or 10 mM MES buffer at pH 6.0 at 298 K. The ITC titration curves are reported
in the Figures S6−S14, including the binding enthalpy and entropy. N is the average number of binding site per nanoparticle measured by ITC.
b
The particle was prepared with 1.0 equiv. FM 4 but without any template. Because the binding constant was estimated from ITC, -ΔG and N are
not listed.
attacking water nucleophile. For retaining glycosidases, the
other carboxylate (∼5 Å apart) is the attacking nucleophile.43
To create a similar feature in our synthetic glucosidase, we
synthesized template 7 with two imine bonds, which allowed
two acids to be installed in the active site, sandwiching the
glycosidic bond in the final structure (Scheme 2). Gratifyingly,
the optimized diacidic catalyst, i.e., MINP(7+8f), hydrolyzed
cellobiose significantly better (94% yield at 60 °C) than the best
monoacidic MINP(5+8i), despite the more acidic sulfonic acid
in the latter (Table 1). The activity, once again, was sensitive to
the distance of the acids. Control experiments indicated that
nonimprinted nanoparticles (NINPs) were ineffective and the
background hydrolysis was low with or without the acid
Figure 1. Effects of solution pH on the hydrolysis of cellobiose by
additives (entries 15−18). Note that MINP(5+8i) and
MINP(5 +8i) and MINP(7+8f). Reactions were performed with 0.2
MINP(7+8f) required different lengths of tethers to connect
mM of cellobiose and 20 μM of MINP in 1.0 mL of buffer (10 mM) at
the acidic groups. The longer tether (n = 5) required for
MINP(7+8f) is reasonable given the longer distance between
the imine group and the exocyclic glycosidic oxygen in the
template.
37 °C for 24 h. Yields were determined by LC-MS using standard curves
generated from authentic samples. NaOAc buffer was used for pH 4.0−
5.0, MES buffer for pH 5.5−6.5, and HEPES buffer for pH 7.0−7.4.
Both the intermediate aldehyde-containing MINP-CHOs and
the final catalysts bound glucose in aqueous buffer, with a
binding constant (Ka) of 7−23 × 103 M−1 determined from
isothermal titration calorimetry (ITC) (Table 2). The binding
affinity matched those for monosaccharides by natural lectins
(Ka = 103−104 M−1).44 Binding for cellobiose was slightly
weaker than that for glucose, and a decrease of pH from 7.4 to
6.0the pH for the cellobiose hydrolysis experiments in Table
1weakened the binding slightly. Molecular imprinting was key
to the binding, as the nonimprinted nanoparticles (NINPs)
prepared with 4 in the absence of template showed negligible
binding. The MINP/NINP binding ratio (i.e., the imprinting
factor) for glucose was >450 for MINP(7+8f).
solution46 but the microenvironment of an acid (or base) is
known to strongly influence its strength. For example, the
ammonium side chain of lysine has a pKa of 10.5 in water but the
value decreases to 5.6 in the active site of acetoacetate
decarboxylase.47 The shift often results from a combination of
hydrophobic effect48 and ionic interactions.49 The former
destabilizes ionic species and increases the pKa of a neutral acid
(e.g., sulfonic or carboxylic acid) but decreases the pKa of a
charged acid such as ammonium. As for the electrostatic
interactions, vicinal positive charges generally make protonation
of a base more difficult and deprotonation of an acid easier. A
recent work of ours shows that 1-pyrenesulfonic acid has a pKa of
ca. 7 inside a MINP pocket.50 In the case of MINP(5+8i), the
overall increase of hydrolytic activity over pH 7.4−4.5 suggests
that the pKa of its sulfonic acid should be lower than 7.
Second, in order for the MINP catalyst to hydrolyze
cellobiose efficiently, the boroxole group needs to bind the
terminal glucose from the nonreducing end. Benzoboroxole has
a pKa of 7.3,51 and the strongest binding (for fructose) occurs at
∼ pH 7.4, very close to the pKa value.40 Inside the MINP, the
anionic form of the boroxole−sugar complex (shown in Scheme
2) can be stabilized by the cationic headgroups of the cross-
linked surfactants,38 and the acid−base equilibrium of boroxole
also strongly depends on environmental polarity.40 Nonetheless,
our ITC data in Table 2 clearly shows that binding for guests
Catalytic Behavior of MINP Catalysts in the Hydrolysis
of Cellobiose. MINP(5+8i) and MINP(7+8f), our best mono-
and diacid catalysts, showed different pH profiles in their
hydrolysis of cellobiose (Figure 1). The monosulfonic acid
catalyst displayed an increase of activity upon decreasing
solution pH all the way to pH 4.5, where the peak activity was
reached. Its overall preference for acidic conditions can be
understood from the following two considerations.
First, to be useful in an acid-catalyzed hydrolysis of acetal,45
the sulfonic acid in the active site of MINP(5+8i) needs to be
able to donate a proton to the glycosidic oxygen and thus be
protonated prior to substrate binding. The pKa of the sulfonic
acid in 2-aminoethanesulfonic acid is 1.5 in an aqueous
5175
https://doi.org/10.1021/jacs.1c01352
J. Am. Chem. Soc. 2021, 143, 5172−5181

Journal of the American Chemical Society
pubs.acs.org/JACS
Article
becomes weaker at lower pH for both MINP(5+8i) and
MINP(7+8f).
A great many wild-type and engineered β-glucosidases have
been reported in the literature, showing a broad range of
catalytic activities in cellobiose hydrolysis.53,54 The catalytic
efficiency of the diacidic MINP(7+8f) (kcat/Km = 132 M−1
min−1) was roughly 1/14th of that of digestive β-glucosidase
GH1 (a relatively slow enzyme) from Spodoptera frugiperda
(kcat/Km = 1780 M−1 min−1, kcat = of 5.7 min−1, and Km = 3.2
mM).55 In comparison to the natural β-glucosidase, MINP-
(7+8f) had a lower catalytic turnover but significantly stronger
binding for the substrate.
Hydrolysis of Cellulose in Aqueous and Nonaqueous
Solution. Encouraged by the facile hydrolysis of cellobiose by
the synthetic glucosidases, we attempted their hydrolysis of
cellulose: a much more challenging substrate due to its highly
crystalline nature.
The initial tests involved hydrolysis in aqueous buffer,
following established procedures to measure enzyme activity.31
For comparison purposes, we used a commercial cellulase
isolated from Aspergillus niger, which is an endocellulase active
on both cellulose and related oligomers.56 A well-established
spectrophotometric assay was used to monitor the hydrolysis of
type 20 Sigmacell cellulose powder (Figure S17).31,57 The
method measures all reducing sugars formed even though liquid
chromatography−mass spectrometry (LC-MS) indicated that
glucose was the dominant product (Figure S18).
The overall pH profile of the MINP-catalyzed hydrolysis,
hence, should reflect a trade-off between the above two effects:
hydrolysis is favored by acidic conditions but binding of the
substrate prefers a neutral pH. Interestingly, although MINP-
(5+8i) and MINP(7+8f) both seemed to experience such a
trade-off, the optimal pH for the dicarboxylic acid catalyst was
significantly higher, at pH 6.0 instead of 4.5 (Figure 1). The
increase is reasonable given that (a) a carboxylic acid has a
higher pKa than a sulfonic acid, at least in an aqueous solution,
and (b) the cooperative catalysis of the two carboxylic acids in
natural glycosidases requires one acid to be protonated and
another one deprotonated.43 A similar mechanism should be
followed by MINP(7+8f) given its high catalytic activity and
sensitivity in the tether length (Table 1).
MINP(5+8i) and MINP(7+8f) displayed Michaelis−Menten
kinetics in the hydrolysis of cellobiose (Figures 2 and 3).
Table 3 compares the hydrolytic properties of MINP(7+8f)
and cellulase from Aspergillus niger under a variety of conditions.
For the natural enzyme, an increase in the catalyst’s
concentration lowered the observed enzyme activity from
0.200 to 0.107 μmol mg−1 h−1 (Table 3, entries 2, 4, and 6).
This result is reasonable, because many cellulases operate
through a processive mechanism in which the enzyme
hydrolyzes a cellulose chain while decrystallizing it from
cellulose crystals.29,58 To do this, the enzyme has a tunnel
(e.g., cellobiohydrolases) or deep cleft (e.g., endoglucanases)
that binds a cellulose chain sometimes with over 20 kcal/mol
binding energy.59 Often a trade-off between processivity and
hydrolytic rate is observed that derives from the binding.60 Since
only the cellulose chains on the surface of the crystals can react
with these enzymes, once the surface is saturated with a strongly
binding cellulase, additional enzymes will not increase the
hydrolytic rate and will decrease the observed average enzyme
activity measured from the formation of reducing sugars.
MINP(7+8f) clearly did not have such an ability. The same
increase of its concentration displayed an opposite trend and
steadily increased the observed enzyme activity, from 0.007 to
0.024 μmol mg−1 h−1 (Table 3, entries 1, 3, and 5). This is not
surprising because the binding free energy for glucose and
cellobiose by the synthetic glucosidase was only 5−6 kcal/mol
(Table 2), far lower than those of common cellulases. Under our
experimental conditions, the steady increase of the observed
enzyme activity suggests that the surface is far from being
saturated by the MINP catalyst, even at 1:1 weight ratio of
cellulose and the catalyst. Because the repeat unit in a cellulose
polymer has a molecular weight of 162 Da and the MINP (with
an average of one active site per nanoparticle) ∼ 50000, the
substrate/catalyst ratio was 300 to 1500 under our experimental
conditions.
Figure 2. Michaelis−Menten plot for the hydrolysis of cellobiose by
MINP(5+8i) in MES buffer (10 mM, pH = 6.0) at 60 °C. [MINP(5
+8i)] = 20.0 μM.
Figure 3. Michaelis−Menten plot for the hydrolysis of cellobiose by
MINP(7+8f) in MES buffer (10 mM, pH = 6.0) at 60 °C. [MINP(7
+8f)] = 10.0 μM.
MINP(5+8i) had a kcat value of 0.027 min−1 and Km of 0.74 mM
in pH 6 buffer at 60 °C. MINP(7+8f) had an experimentally
comparable Km value and a turnover 3.2-times faster than the
monoacidic MINP. The rate constant for cellobiose hydrolysis
in acidic water is estimated from a modified Saeman equation to
be ∼4.9 × 10−10 −1 at pH 6 and 60 °C.52 Thus, a rate
s
acceleration of ca. 9 × 105 and 3 × 106 was achieved by
MINP(5+8i) and MINP(7+8f), respectively, over acid-
catalyzed hydrolysis in aqueous solution at pH 6. Clearly,
positioning the acidic groups at the glycosidic bond is hugely
beneficial to the catalyzed hydrolysis.
Despite its lack of a processive mechanism, MINP(7+8f)
showed activities that actually can be compared with those of
natural cellulase: the enzyme/MINP ratio in Table 3 decreased
steadily from 29 all the way to 4.5 with an increase of catalyst
concentration (entries 1−6). Even if the activity of the enzyme
5176
https://doi.org/10.1021/jacs.1c01352
J. Am. Chem. Soc. 2021, 143, 5172−5181

Journal of the American Chemical Society
pubs.acs.org/JACS
Article
a
Table 3. Hydrolysis of Cellulose by Cellulases and MINP(7+8f)
catalyst conc
(mg/mL)
temp
(°C)
[reducing sugar]
enzyme activity
relative activity (enzyme/
MINP)
entry
catalysts
solvent
(mg/mL)
(μmol mg−1 h−1
)
1
2
3
4
5
6
7
8
MINP(7+8f)
cellulases
MINP(7+8f)
cellulases
MINP(7+8f)
cellulases
MINP(7+8f)
cellulases
MINP(7+8f)
cellulases
MINP(7+8f)
cellulases
1.0
1.0
2.0
2.0
5.0
5.0
5.0
5.0
5.0
5.0
5.0
5.0
pH 5 NaOAc buffer
pH 5 NaOAc buffer
pH 5 NaOAc buffer
pH 5 NaOAc buffer
pH 5 NaOAc buffer
pH 5 NaOAc buffer
pH 5 NaOAc buffer
pH 5 NaOAc buffer
pH 5 NaOAc buffer
pH 5 NaOAc buffer
83% H3PO4
37
37
37
37
37
37
60
60
90
90
37
37
0.015 0.002
0.43 0.03
0.07 0.01
0.68 0.35
0.25 0.04
1.15 0.21
0.43 0.08
0.24 0.03
0.79 0.13
0
0.007 0.001
0.200 0.011
0.017 0.007
0.158 0.021
0.024 0.004
0.107 0.019
0.039 0.008
0.022 0.003
0.073 0.012
0
29
9
4.5
0.56
0
9
10
11
12
0.71 0.33
0
0.067 0.031
0
0
83% H3PO4
a
The reactions were performed in duplicates with [cellulose] = 5.0 mg/mL in 1.0 mL of 10 mM NaOAc buffer (pH 5) for 12 h unless indicated
otherwise.
a
Table 4. Hydrolysis of Cellulose by MINP(5+8i) and MINP(7+8f) in Mixtures of Ionic Liquids and a Polar Aprotic Solvent
entry
catalyst
solvent
temp. (°C)
[reducing sugar] (mg/mL)
enzyme activity(μmol mg−1 h−1
)
1
2
3
4
5
6
7
8
MINP(5+8i)
MINP(5+8i)
MINP(5+8i)
MINP(5+8i)
MINP(5+8i)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
MINP(7+8f)
[C4mim]Cl
90
90
90
110
130
90
90
90
90
90
90
110
130
90
90
90
90
90
90
90
90
0.44 0.05
1.33 0.17
2.24 0.14
2.87 0.21
3.41 0.33
0.21 0.04
1.17 0.14
1.41 0.08
1.98 0.21
2.87 0.17
4.17 0.37
4.51 0.44
4.97 0.61
5.92 0.11
6.76 0.32
1.55 0.11
5.41 0.41
5.87 0.24
6.94 0.15
7.24 0.18
6.25 0.58
0.051 0.006
0.154 0.019
0.258 0.016
0.329 0.024
0.391 0.038
0.024 0.005
0.135 0.016
0.163 0.009
0.229 0.024
0.332 0.020
0.480 0.043
0.519 0.051
0.572 0.071
0.431 0.014
0.197 0.011
0.179 0.013
0.623 0.047
0.676 0.028
0.504 0.021
0.211 0.035
1:1 [C4mim]Cl/DMF
1:1 [C4mim]Cl/DMSO
1:1 [C4mim]Cl/DMSO
1:1 [C4mim]Cl/DMSO
[C4mim]Cl
9:1 [C4mim]Cl/DMSO
8:2 [C4mim]Cl/DMSO
7:3 [C4mim]Cl/DMSO
6:4 [C4mim]Cl/DMSO
5:5 [C4mim]Cl/DMSO
5:5 [C4mim]Cl/DMSO
5:5 [C4mim]Cl/DMSO
5:5 [C4mim]Cl/DMSO
5:5 [C4mim]Cl/DMSO
4:6 [C4mim]Cl/DMSO
9
10
11
12
13
14
15
16
17
18
19
20
21
b
c
d
5:5 [C4mim]Cl/DMSO+5% H2O
1:9 [C2mim]OAc/DMSO
1:9 [C2mim]OAc/DMSO
1:9 [C2mim]OAc/DMSO
2:8 [C2mim]OAc/DMSO+5% H2O
b
c
0.719 0.067
a
b
The reactions were performed in duplicates with [cellulose] = 8.0 mg/mL and [MINP] = 2.0 mg/mL in 0.5 mL of solvent for 24 h. [MINP] =
c
d
3.2 mg/mL. [MINP] = 8.0 mg/mL. Cellulose dissolved partially in this mixture.
at low concentration is compared with the activity of the MINP
at high concentration (entries 2 and 5), the difference was a
factor of 8.3.
lose62,63 and are known to facilitate the acid-catalyzed hydrolysis
of cellulose.64−66 Unfortunately, enzymes have poor stability
and low activity in ionic liquids, especially at elevated
temperatures.30 In contrast, the extraordinary stability of the
MINP-based artificial enzymes allowed the hydrolysis to be
performed under conditions totally impossible for natural
enzymes.
One notable strength of the synthetic glucosidase was its
stability, due to its highly cross-linked nature. While the natural
cellulase continued to lose activity as the reaction temperature
increased from 37 to 60 and then to 90 °C in the pH 5 buffer,
MINP(7+8f) became more efficient, with its activity at 90 °C
(0.073 μmol mg−1 h−1) reaching nearly 70% of that of cellulases
(0.107 μmol mg−1 h−1) at 37 °C at the same concentration of
catalyst. The robustness of the MINP catalysts was also shown
by its ability to hydrolyze cellulose in 83% phosphoric acid,
which dissolved cellulose completely61 but inactivated the
natural enzymes. The synthetic catalyst was nearly 3 times more
active in this extremely acidic condition than in pH 5 buffer at
the same temperature of 37 °C.
Table 4 shows that the activity of both MINP(5+8i) and
MINP(7+8f) improved in ionic liquids as the reaction mixture
became homogeneous. A polar aprotic solvent (dimethyl
sulfoxide or DMSO) could be added to further speed up the
hydrolysis as long as the reaction mixture stayed homoge-
neous.67 The solvent effect in the ionic liquids/DMSO mixtures
followed a trend opposite to that observed in cellulose hydrolysis
catalyzed by p-toluenesulfonic acid (p-TSA). Hydrolysis of
cellulose (and also cellobiose) is reported to become faster with
a higher fraction of ionic liquids in the binary solvent mixture for
the small acid catalyst, as a result of increased Hammett acidity
Ionic liquids such as 1-butyl-3-methylimidazolium chloride
([C4mim]Cl) have a tremendous ability to dissolve cellu-
5177
https://doi.org/10.1021/jacs.1c01352
J. Am. Chem. Soc. 2021, 143, 5172−5181

Journal of the American Chemical Society
pubs.acs.org/JACS
Article
of p-TSA by the ionic liquids.68 For our MINP catalysts, an
increase of acidity first is not expected to be useful, because the
more acidic, monosulfonic acid MINP(5+8i) was less active
than the less acidic, dicarboxylic acid MINP(7+8f) in aqueous
or nonaqueous solutions. Apparently, cooperative catalysis
between the carboxylate/carboxylic acid was far more powerful
than the “brute force” derived from a single stronger sulfonic
acid. Moreover, when the acid groups are located inside the
MINP active site, the benefit seen in the solution most likely
would not even occur. Higher ionic liquids in the binary mixture
substantially increases the viscosity of the solution.68 Diffusion
of the 5 nm-sized MINP catalyst thus would become
significantly faster as more DMSO is added to the mixture,
and many have helped the mass transfer and contributed to the
higher activity of MINP(7+8f).
Figure 4. Comparison of reducing sugar formed during hydrolysis of
cellulose by the synthetic MINP catalysts in 2:8 [C2mim]OAc/DMSO
with 5% H2O at 90 °C and natural cellulase in NaOAc buffer pH 5.0 at
37 °C. [cellulose] = 8 mg/mL, [catalyst] = 2 mg/mL.
In the 1:1 mixture of [C4mim]Cl/DMSO, the enzyme activity
of MINP(7+8f) reached 0.48 μmol mg− 1h−1 at 90 °C (Table 4,
entry 11), 2.4 times the best (0.20 μmol mg−1 h−1) from the
cellulase in aqueous buffer under our experimental conditions
(Table 3, entry 2). An increase of temperature to 130 °C
increased the activity of MINP(7+8f) further to 0.572 μmol
MINP(5+8i) and MINP(7+8f) maintained their activity very
well over the extended period of heating (Figure 4). Table 2
shows that both MINPs bound glucose slightly more strongly
than cellobiose, at least in aqueous solution. Thus, product
inhibition was a concern for these synthetic enzymes, which was
also a challenge for natural cellulases.29 To our delight, no
obvious slowdown of the hydrolysis was seen in Figure 4, when
the initial slope and those of the later reaction times were
compared. MINP contains multiple hydroxylated surface ligands
(Scheme 1). It is possible that in the ionic liquids/DMSO
mixture, a dissolved cellulose chain could interact with these
and/or other surface functionalities on the cross-linked micelle.
Small sugar products are not expected to benefit from such
interactions.
One advantage of MINP(7+8f) was its well-defined catalytic
site and improved mass transfer in the homogeneous reaction
mixture. Recovery of homogeneous catalysts is often an
associated problem. In the MINP preparation, the surface−
core doubly cross-linked micelles were typically covered with
monoazide 3 for enhanced hydrophilicity and facile purification
(Scheme 1). Without the termination, the alkyne-containing
MINPs could be easily “clicked” onto azide-functionalized
magnetic nanoparticles (MNPs) prepared via a literature
procedure70 (Scheme S3). The resulting MINP(7+8f)@MNP
composite (Figure 5) became a reusable heterogeneous catalyst
that could be recovered simply with a magnet after each
hydrolytic cycle. Figure 6 shows that the catalyst maintained
75% of its activity after 10 cycles of hydrolysis.
mg−1 −1 (Table 4, entry 13), underscoring the robustness of the
h
synthetic enzyme. A small amount of water could speed up the
reaction even further (Table 4, entry 17; Table S1).
1-Ethyl-3-methylimidazolium acetate ([C2mim]OAc) can
dissolve cellulose particularly well in the presence of other
organic solvents.67 In our hands, a 1:9 [C2mim]OAc/DMSO
mixture increased the hydrolytic activity of MINP(7+8f) to
0.676 μmol mg−1 h−1 (Table 4, entry 18). Notably, in these
homogeneous reaction mixtures, increasing the concentration of
the MINP catalyst was no longer beneficial and actually
decreased the observed enzyme activity (compare entries 11
with 14−15 and also entries 18−20), similar to the trend
observed for the cellulase in aqueous reaction. Thus, a high
concentration of the MINP catalyst was no longer needed in the
ionic liquids/DMSO mixture, possibly because the dissolved
polymer chains were already saturated with the MINP at the low
catalyst loading. Because the MINP could only bind the
nonreducing end of a cellulose chain through boronate bonds,
each polymer chain could only accommodate a single catalyst.
Also, boronate bonds are labile in water;40 stronger binding in
the more organic ionic liquids/DMSO mixture was not
surprising.
Addition of water to the [C2mim]OAc/DMSO mixture
required more ionic liquids to keep cellulose homogeneous in
the solution. The highest activity achieved by our synthetic
glucosidase was 0.719 μmol mg−1 h−1 in 2:8 [C2mim]OAc/
DMSO with 5% H2O, ∼ 3.6 times of the best for the cellulase in
aqueous buffer at 37 °C. Ionic liquids generally inactive
cellulases30,69 and indeed inactivated the cellulase from
Aspergillus niger under our harsh experimental conditions. The
best comparison for MINP(7+8f)a mimic of β-glucosidase
in the literature is the cationized β-glucosidase reported by
Hallet and co-workers, which displayed remarkable stability in
ionic liquids.31 Nonetheless, when the latter was used as the sole
catalyst to hydrolyze cellulose in [C2mim]OAc, the observed
enzyme activity was <0.1 μmol mg−1 h−1 at temperatures ranging
from 50 to 120 °C.
CONCLUSIONS
■
In summary, micellar imprinting using judiciously designed
templates and postmodification together provide a powerful way
to construct synthetic enzymes in a bottom-up fashion. What
was key to the construction was the strong templating effect
exemplified by the large imprint/nonimprint ratio (Table 2),
nanodimension of the imprinted micelle, good accessibility of
the imprinted pocket, and good solubility of MINP in solvents
such as DMF and DMSO. These features allowed a facile one-
pot synthesis of complex imprinted pockets from small-molecule
template molecules in the core of water-soluble organic
nanoparticles. In addition, they enabled chemical derivatization
of the imprinted pockets by standard chemical reactions to
convert them into synthetic enzymes with accurately positioned,
tunable catalytic groups.
Figure 4 shows the amount of reducing sugar formed over a
period of 24 h at 90 °C from the best monoacidic and diacidic
catalysts. We also included the reaction profile for the cellulase
from Aspergillus niger in aqueous buffer at 37 °C for comparison.
5178
https://doi.org/10.1021/jacs.1c01352
J. Am. Chem. Soc. 2021, 143, 5172−5181

Journal of the American Chemical Society
pubs.acs.org/JACS
Article
nonaqueous solutions where natural enzymes are unable to
operate. Not only can the operating window of cellulose
depolymerization be greatly expanded in such a way, the
excellent reusability of the synthetic enzymes also represents
another major advantage.
ASSOCIATED CONTENT
* Supporting Information
■
sı
The Supporting Information is available free of charge at
https://pubs.acs.org/doi/10.1021/jacs.1c01352.
Synthetic procedures, characterization of compounds and
materials, ITC binding curves, additional tables and
figures, and NMR data (PDF)
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
Xiaowei Li − Department of Chemistry, Iowa State University,
Ames, Iowa 50011-3111, United States; orcid.org/0000-
0001-9479-4857
Milad Zangiabadi − Department of Chemistry, Iowa State
University, Ames, Iowa 50011-3111, United States;
orcid.org/0000-0003-0886-1553
Complete contact information is available at:
https://pubs.acs.org/10.1021/jacs.1c01352
Figure 5. TEM images of (a, b) silica gel-coated Fe3O4 magnetic
nanoparticles, (c, d) NH2-MNP, (e−h) the final MINP(7+8f)@MNP.
Notes
The authors declare the following competing financial
interest(s): Iowa State University Research Foundation has
filed a patent application on the technology described in the
work.
ACKNOWLEDGMENTS
We thank NSF (CHE-1708526) for supporting the research.
■
REFERENCES
■
(1) Breslow, R. Artificial Enzymes; Wiley-VCH: Weinheim, 2005.
(2) Kirby, A. J.; Hollfelder, F. From Enzyme Models to Model Enzymes;
Royal Society of Chemistry: Cambridge, UK, 2009.
(3) Raynal, M.; Ballester, P.; Vidal-Ferran, A.; van Leeuwen, P. W. N.
M. Supramolecular Catalysis. Part 2: Artificial Enzyme Mimics. Chem.
Soc. Rev. 2014, 43, 1734−1787.
Figure 6. Recyclability of MINP(7+8f)@MNP for cellulose hydrolysis
in 2:8 [C2mim]OAc/DMSO with 5% H2O at 90 °C.
(4) Knowles, J. R. Enzyme Catalysis: Not Different, Just Better. Nature
1991, 350, 121−124.
(5) Wulff, G. Enzyme-Like Catalysis by Molecularly Imprinted
Polymers. Chem. Rev. 2002, 102, 1−28.
(6) Haupt, K.; Mosbach, K. Molecularly Imprinted Polymers and
Their Use in Biomimetic Sensors. Chem. Rev. 2000, 100, 2495−2504.
(7) Ye, L.; Mosbach, K. Molecular Imprinting: Synthetic Materials as
Substitutes for Biological Antibodies and Receptors. Chem. Mater.
2008, 20, 859−868.
(8) Pan, J.; Chen, W.; Ma, Y.; Pan, G. Molecularly Imprinted Polymers
as Receptor Mimics for Selective Cell Recognition. Chem. Soc. Rev.
2018, 47, 5574−5587.
(9) Zhang, H. Molecularly Imprinted Nanoparticles for Biomedical
Applications. Adv. Mater. 2020, 32, 1806328.
(10) Wulff, G.; Liu, J. Design of Biomimetic Catalysts by Molecular
Imprinting in Synthetic Polymers: The Role of Transition State
Stabilization. Acc. Chem. Res. 2012, 45, 239−247.
MINP(7+8f) came close to some natural β-glucosidases in its
ability to hydrolyze cellobiose in aqueous solution and could
function under conditions completely impossible for natural
enzymes such as 83% H3PO4 and ionic liquids/DMSO mixture
at 90 °C. An enzyme activity of 0.719 μmol mg−1 h−1 in 2:8 [C2
mim]OAc/DMSO with 5% H2O was unprecedented for a
synthetic glucosidase. A concoction of enzymes are often used
by nature and also in industrial processes to hydrolyze cellulose:
endocellulase to target the amorphous region of cellulose fibers,
exocellulase to depolymerize the chain from the nonreducing
end into glucose oligomers, and β-glucosidase to hydrolyze the
oligomers.29 It is envisioned that multiple synthetic mimics of
these enzymes can work synergistically likewise, in aqueous and
5179
https://doi.org/10.1021/jacs.1c01352
J. Am. Chem. Soc. 2021, 143, 5172−5181

Journal of the American Chemical Society
pubs.acs.org/JACS
Article
(11) Muratsugu, S.; Shirai, S.; Tada, M. Recent Progress in
Molecularly Imprinted Approach for Catalysis. Tetrahedron Lett.
2020, 61, 151603.
(33) James, T. D.; Phillips, M. D.; Shinkai, S. Boronic Acids in
Saccharide Recognition; RSC Publishing: Cambridge, 2006.
(34) Pal, A.; Bérubé, M.; Hall, D. G. Design, Synthesis, and Screening
of a Library of Peptidyl Bis(Boroxoles) as Oligosaccharide Receptors in
Water: Identification of a Receptor for the Tumor Marker TF-Antigen
Disaccharide. Angew. Chem., Int. Ed. 2010, 49, 1492−1495.
(35) Li, X.; Zhao, Y. Synthetic Glycosidase Distinguishing Glycan and
Glycosidic Linkage in Its Catalytic Hydrolysis. ACS Catal. 2020, 10,
13800−13808.
(36) Dowlut, M.; Hall, D. G. An Improved Class of Sugar-Binding
Boronic Acids, Soluble and Capable of Complexing Glycosides in
Neutral Water. J. Am. Chem. Soc. 2006, 128, 4226−4227.
(37) Kim, H.; Kang, Y. J.; Kang, S.; Kim, K. T. Monosaccharide-
Responsive Release of Insulin from Polymersomes of Polyboroxole
Block Copolymers at Neutral pH. J. Am. Chem. Soc. 2012, 134, 4030−
4033.
(38) Gunasekara, R. W.; Zhao, Y. A General Method for Selective
Recognition of Monosaccharides and Oligosaccharides in Water. J. Am.
Chem. Soc. 2017, 139, 829−835.
(39) Li, X.; Zhao, Y. Synthetic Glycosidases for the Precise Hydrolysis
of Oligosaccharides and Polysaccharides. Chem. Sci. 2021, 12, 374−
383.
(40) Bérubé, M.; Dowlut, M.; Hall, D. G. Benzoboroxoles as Efficient
Glycopyranoside-Binding Agents in Physiological Conditions: Struc-
ture and Selectivity of Complex Formation. J. Org. Chem. 2008, 73,
6471−6479.
(41) Xing, X.; Zhao, Y. Binding-Promoted Chemical Reaction in the
Nanospace of a Binding Site: Effects of Environmental Constriction.
Org. Biomol. Chem. 2018, 16, 2855−2859.
(42) Xing, X.; Zhao, Y. Fluorescent Nanoparticle Sensors with Tailor-
Made Recognition Units and Proximate Fluorescent Reporter Groups.
New J. Chem. 2018, 42, 9377−9380.
(43) Zechel, D. L.; Withers, S. G. Glycosidase Mechanisms: Anatomy
of a Finely Tuned Catalyst. Acc. Chem. Res. 2000, 33, 11−18.
(44) Toone, E. J. Structure and Energetics of Protein-Carbohydrate
Complexes. Curr. Opin. Struct. Biol. 1994, 4, 719−728.
(45) Fife, T. H.; Jao, L. K. Substituent Effects in Acetal Hydrolysis. J.
Org. Chem. 1965, 30, 1492−1495.
(46) CRC Handbook of Chemistry and Physics; 88th ed.; Lide, D. R., Ed.
CRC Press/Taylor & Francis Group: Boca Raton, FL, 2007; p 8−42.
(47) Westheimer, F. H. Coincidences, Decarboxylation, and Electro-
static Effects. Tetrahedron 1995, 51, 3−20.
(48) Matulis, D.; Bloomfield, V. A. Thermodynamics of the
Hydrophobic Effect. I. Coupling of Aggregation and pKa Shifts in
Solutions of Aliphatic Amines. Biophys. Chem. 2001, 93, 37−51.
(49) Henao, J. D.; Suh, Y.-W.; Lee, J.-K.; Kung, M. C.; Kung, H. H.
Striking Confinement Effect: Aucl4− Binding to Amines in a Nanocage
Cavity. J. Am. Chem. Soc. 2008, 130, 16142−16143.
(50) Bose, I.; Fa, S.; Zhao, Y. Tunable Artificial Enzyme-Cofactor
Complex for Selective Hydrolysis of Acetals. J. Org. Chem. 2021, 86,
1701−1711.
(51) Tomsho, J. W.; Pal, A.; Hall, D. G.; Benkovic, S. J. Ring Structure
and Aromatic Substituent Effects on the pKa of the Benzoxaborole
Pharmacophore. ACS Med. Chem. Lett. 2012, 3, 48−52.
(52) Mohd Shafie, Z.; Yu, Y.; Wu, H. Effect of Initial Ph on
Hydrothermal Decomposition of Cellobiose under Weakly Acidic
Conditions. Fuel 2015, 158, 315−321.
(53) Singhania, R. R.; Patel, A. K.; Pandey, A.; Ganansounou, E.
Genetic Modification: A Tool for Enhancing Beta-Glucosidase
Production for Biofuel Application. Bioresour. Technol. 2017, 245,
1352−1361.
(12) Kirsch, N.; Hedin-Dahlström, J.; Henschel, H.; Whitcombe, M.
J.; Wikman, S.; Nicholls, I. A. Molecularly Imprinted Polymer Catalysis
of a Diels-Alder Reaction. J. Mol. Catal. B: Enzym. 2009, 58, 110−117.
(13) Chen, Z. Y.; Xu, L.; Liang, Y.; Zhao, M. P. pH-Sensitive Water-
Soluble Nanospheric Imprinted Hydrogels Prepared as Horseradish
Peroxidase Mimetic Enzymes. Adv. Mater. 2010, 22, 1488−1492.
(14) Servant, A.; Haupt, K.; Resmini, M. Tuning Molecular
Recognition in Water-Soluble Nanogels with Enzyme-Like Activity
for the Kemp Elimination. Chem. - Eur. J. 2011, 17, 11052−11059.
(15) Shen, X.; Huang, C.; Shinde, S.; Jagadeesan, K. K.; Ekström, S.;
Fritz, E.; Sellergren, B. Catalytic Formation of Disulfide Bonds in
Peptides by Molecularly Imprinted Microgels at Oil/Water Interfaces.
ACS Appl. Mater. Interfaces 2016, 8, 30484−30491.
(16) Yuan, Y.; Yang, Y.; Faheem, M.; Zou, X.; Ma, X.; Wang, Z.; Meng,
Q.; Wang, L.; Zhao, S.; Zhu, G. Molecularly Imprinted Porous
Aromatic Frameworks Serving as Porous Artificial Enzymes. Adv.
Mater. 2018, 30, 1800069.
(17) Awino, J. K.; Zhao, Y. Protein-Mimetic, Molecularly Imprinted
Nanoparticles for Selective Binding of Bile Salt Derivatives in Water. J.
Am. Chem. Soc. 2013, 135, 12552−12555.
(18) Chen, K.; Zhao, Y. Effects of Nano-Confinement and
Conformational Mobility on Molecular Imprinting of Cross-Linked
Micelles. Org. Biomol. Chem. 2019, 17, 8611−8617.
(19) Duan, L.; Zangiabadi, M.; Zhao, Y. Synthetic Lectins for Selective
Binding of Glycoproteins in Water. Chem. Commun. 2020, 56, 10199−
10202.
(20) Zangiabadi, M.; Zhao, Y. Molecularly Imprinted Polymeric
Receptors with Interfacial Hydrogen Bonds for Peptide Recognition in
Water. ACS Appl. Polym. Mater. 2020, 2, 3171−3180.
(21) Awino, J. K.; Gunasekara, R. W.; Zhao, Y. Sequence-Selective
Binding of Oligopeptides in Water through Hydrophobic Coding. J.
Am. Chem. Soc. 2017, 139, 2188−2191.
(22) Arifuzzaman, M.; Zhao, Y. Artificial Zinc Enzymes with Fine-
Tuned Catalytic Active Sites for Highly Selective Hydrolysis of
Activated Esters. ACS Catal. 2018, 8, 8154−8161.
(23) Li, X.; Zhao, Y. Chiral Gating for Size- and Shape-Selective
Asymmetric Catalysis. J. Am. Chem. Soc. 2019, 141, 13749−13752.
(24) Hu, L.; Arifuzzaman, M. D.; Zhao, Y. Controlling Product
Inhibition through Substrate-Specific Active Sites in Nanoparticle-
Based Phosphodiesterase and Esterase. ACS Catal. 2019, 9, 5019−
5024.
(25) Bose, I.; Zhao, Y. pH-Controlled Nanoparticle Catalysts for
Highly Selective Tandem Henry Reaction from Mixtures. ACS Catal.
2020, 10, 13973−13977.
(26) Huber, G. W.; Iborra, S.; Corma, A. Synthesis of Transportation
Fuels from Biomass: Chemistry, Catalysts, and Engineering. Chem. Rev.
2006, 106, 4044−4098.
(27) Luterbacher, J. S.; Martin Alonso, D.; Dumesic, J. A. Targeted
Chemical Upgrading of Lignocellulosic Biomass to Platform Molecules.
Green Chem. 2014, 16, 4816−4838.
(28) Jing, Y.; Guo, Y.; Xia, Q.; Liu, X.; Wang, Y. Catalytic Production
of Value-Added Chemicals and Liquid Fuels from Lignocellulosic
Biomass. Chem. 2019, 5, 2520−2546.
(29) Payne, C. M.; Knott, B. C.; Mayes, H. B.; Hansson, H.; Himmel,
M. E.; Sandgren, M.; Ståhlberg, J.; Beckham, G. T. Fungal Cellulases.
Chem. Rev. 2015, 115, 1308−1448.
(30) Wahlström, R. M.; Suurnäkki, A. Enzymatic Hydrolysis of
Lignocellulosic Polysaccharides in the Presence of Ionic Liquids. Green
Chem. 2015, 17, 694−714.
(54) Sun, J.; Wang, W.; Ying, Y.; Hao, J. A Novel Glucose-Tolerant
Gh1 B-Glucosidase and Improvement of Its Glucose Tolerance Using
Site-Directed Mutation. Appl. Biochem. Biotechnol. 2020, 192, 999−
1015.
(31) Brogan, A. P. S.; Bui-Le, L.; Hallett, J. P. Non-Aqueous
Homogenous Biocatalytic Conversion of Polysaccharides in Ionic
Liquids Using Chemically Modified Glucosidase. Nat. Chem. 2018, 10,
859−865.
́
(55) Tamaki, F. K.; Araujo, E. M.; Rozenberg, R.; Marana, S. R. A
(32) Wulff, G.; Vesper, W. Enzyme-Analogue Built Polymers.8.
Preparation of Chromatographic Sorbents with Chiral Cavities for
Racemic-Resolution. J. Chromatogr. 1978, 167, 171−186.
Mutant B-Glucosidase Increases the Rate of the Cellulose Enzymatic
Hydrolysis. Biochem. Biophys. Rep. 2016, 7, 52−55.
5180
https://doi.org/10.1021/jacs.1c01352
J. Am. Chem. Soc. 2021, 143, 5172−5181

Journal of the American Chemical Society
pubs.acs.org/JACS
Article
(56) Okada, G. Cellulase of Aspergillus Niger. Methods Enzymol. 1988,
160, 259−264.
(57) Jurick II, W.M.; Vico, I.; Whitaker, B.D.; Gaskins, V.L.;
Janisiewic, W.J. Application of the 2-Cyanoacetamide Method for
Spectrophotometric Assay of Cellulase Enzyme Activity. Plant Pathol. J.
2012, 11, 38−41.
(58) Igarashi, K.; Uchihashi, T.; Koivula, A.; Wada, M.; Kimura, S.;
Okamoto, T.; Penttilä, M.; Ando, T.; Samejima, M. Traffic Jams Reduce
Hydrolytic Efficiency of Cellulase on Cellulose Surface. Science 2011,
333, 1279−1282.
(59) Payne, C. M.; Jiang, W.; Shirts, M. R.; Himmel, M. E.; Crowley,
M. F.; Beckham, G. T. Glycoside Hydrolase Processivity Is Directly
Related to Oligosaccharide Binding Free Energy. J. Am. Chem. Soc.
2013, 135, 18831−18839.
(60) Nakamura, A.; Watanabe, H.; Ishida, T.; Uchihashi, T.; Wada,
M.; Ando, T.; Igarashi, K.; Samejima, M. Trade-Off between
Processivity and Hydrolytic Velocity of Cellobiohydrolases at the
Surface of Crystalline Cellulose. J. Am. Chem. Soc. 2014, 136, 4584−
4592.
(61) Zhang, Y. H. P.; Cui, J.; Lynd, L. R.; Kuang, L. R. A Transition
from Cellulose Swelling to Cellulose Dissolution by O-Phosphoric
Acid: Evidence from Enzymatic Hydrolysis and Supramolecular
Structure. Biomacromolecules 2006, 7, 644−648.
(62) Wang, H.; Gurau, G.; Rogers, R. D. Ionic Liquid Processing of
Cellulose. Chem. Soc. Rev. 2012, 41, 1519−1537.
(63) Swatloski, R. P.; Spear, S. K.; Holbrey, J. D.; Rogers, R. D.
Dissolution of Cellose with Ionic Liquids. J. Am. Chem. Soc. 2002, 124,
4974−4975.
(64) Rinaldi, R.; Palkovits, R.; Schuth, F. Depolymerization of
Cellulose Using Solid Catalysts in Ionic Liquids. Angew. Chem., Int. Ed.
2008, 47, 8047−8050.
(65) Binder, J. B.; Raines, R. T. Fermentable Sugars by Chemical
Hydrolysis of Biomass. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 4516−
4521.
(66) Zhang, Z.; Song, J.; Han, B. Catalytic Transformation of
Lignocellulose into Chemicals and Fuel Products in Ionic Liquids.
Chem. Rev. 2017, 117, 6834−6880.
(67) Rinaldi, R. Instantaneous Dissolution of Cellulose in Organic
Electrolyte Solutions. Chem. Commun. 2011, 47, 511−513.
̀
(68) de Oliveira, H. F. N.; Fares, C.; Rinaldi, R. Beyond a Solvent: The
Roles of 1-Butyl-3-Methylimidazolium Chloride in the Acid-Catalysis
for Cellulose Depolymerisation. Chem. Sci. 2015, 6, 5215−5224.
(69) Turner, M. B.; Spear, S. K.; Huddleston, J. G.; Holbrey, J. D.;
Rogers, R. D. Ionic Liquid Salt-Induced Inactivation and Unfolding of
Cellulase from Trichoderma Reesei. Green Chem. 2003, 5, 443−447.
(70) Lin, P.-C.; Chou, P.-H.; Chen, S.-H.; Liao, H.-K.; Wang, K.-Y.;
Chen, Y.-J.; Lin, C.-C. Ethylene Glycol-Protected Magnetic Nano-
particles for a Multiplexed Immunoassay in Human Plasma. Small
2006, 2, 485−489.
5181
https://doi.org/10.1021/jacs.1c01352
J. Am. Chem. Soc. 2021, 143, 5172−5181