Dynamic Tuning in Synthetic Glycosidase for Selective Hydrolysis of Alkyl and Aryl Glycosides

Abstract

Enzymes use sophisticated conformational control to optimize the dynamics of their protein framework for efficient catalysis. Although it is difficult to employ a similar strategy to improve catalysis in a synthetic enzyme, we here report that modulation of the dynamics of the substrate in the active site is readily achievable in a complex between a molecularly imprinted nanoparticle (MINP) and its acid cofactor, through tuning of the size and shape of the imprinted site. As the alkyl glucoside substrate is bound with increasing strength and held in a more tightly fitted pocket, the acid-catalyzed glycan hydrolysis becomes more difficult. A larger, wider active site, although less able to bind the substrate, affords a higher catalytic activity, likely due to easier alignment of the substrate and the acid cofactor for a general acid catalysis. The substrate selectivity is controlled by both the tightness of the aglycon-binding site and the orientation of the glycan-binding boroxole group.

Graphical Abstract

Introduction

Dynamics of enzymes is critical to their catalysis. The importance of active-site dynamics is implied in the induced-fit model of enzymes, which states that a substrate of an enzyme causes “an appreciable change” of the active site to optimize the orientation of the catalytic groups for catalysis.¹ Dynamics, however, goes far beyond the active site, sometimes throughout the protein structure, to strongly affect catalysis. Lactate dehydrogenase (LDH), as an example, upon binding its substrate, undergoes a series of catalytically important conformational changes over microsecond to millisecond timescale before the substrate is shuttled to the active site.²

There is significant interest in developing synthetic catalysts with enzyme-like efficiency and selectivity. Traditionally, artificial enzymes have been constructed with macrocycles such as cyclodextrins.^3–5 Their well-formed hydrophobic pockets are used for substrate binding; installation of catalytic groups converts them into enzyme-like catalysts. Despite the great progress made, it is challenging to tune the shape of highly symmetrical macrocycles for complex-shaped substrates. Controlling the dynamics of the catalytic groups rationally is even more difficult.

Molecular imprinting is an alternative method to prepare synthetic receptors.^6–8 Facile polymerization and cross-linking around template molecules quickly affords a cross-linked polymer network with embedded imprinted sites ideally complementary to the templates in size, shape, and distribution of functional groups. Installation of catalytic groups in the imprinted site, either directly or through postmodification, enables molecularly imprinted polymers (MIPs) to perform enzyme-like catalysis.^9–15 Dynamics of a polymer network can be tuned in principle by its cross-linking density, which is ultimately controlled by the type and amount of cross-linker used in the polymerization.¹⁶ Nonetheless, a high cross-linking density is generally needed for MIPs to maintain the integrity of their imprinted sites, and altering this parameter tends to negatively impact the imprinting factors of MIPs.¹⁷

Our group in recent years developed a method to perform molecular imprinting in cross-linked surfactant micelles. A nanoconfined environment in micellar imprinting affords an extraordinary imprinting effect¹⁸ and the resulting molecularly imprinted nanoparticles (MINPs) often display imprinting factors in the hundreds or even 10,000.¹⁹ Specific functional groups may be introduced in the imprinted pockets to turn MINPs into artificial enzymes.^20–22

In this work, we report a synthetic glycosidase comprising an imprinted cross-linked micelle and an acid cofactor, for selective hydrolysis of alkyl and aryl glycosides. Molecular imprinting allowed us to tune the size and shape of the aglycon-binding pocket of the active site and, indirectly, dynamics of the substrate. Interestingly, mobility of the aglycon of the substrate in the active site was found critical to the catalyzed hydrolysis and the “most fitted” active site had the lowest catalytic activity. Selective hydrolysis of glycosides is highly challenging and only a few examples have been reported to hydrolyze activated aryl glycosides.^23–27 Alkyl glycosides are far more stable than the commonly used p-nitrophenyl glycosides and thus much more difficult to hydrolyze.²⁸ The half-life of methyl β-d-glucopyranoside, for example, is about 4.7 million years at 25 °C over pH 7–14.²⁹ Meanwhile, they represent an important class of biomolecules—i.e., glycolipids—and play vital roles in biological processes such as cell growth, cell trafficking, and cancer development.^30–33

Results and Discussion

Synthesis and Characterization of MINPs.

Our preparation of a synthetic alkyl glycosidase is described in detail in the Experimental Section, adapted from reported procedures.²⁰ Briefly, template molecule 1a–f, amphiphilic in nature, is solubilized in water by mixed micelles of cross-linkable surfactants 3a and 3b (Scheme 1). To help hydrophilic templates move into micelles for molecular imprinting, a hydrophobic functional monomer (FM) is generally used.^34,35 In this case, vinylphenylboroxole 2 forms boronate 1a–f·2 with the 4,6-diol of glucosides in situ,^36,37 and the resulting anionic boronate is stabilized by cationic micelles.^21,34 The highly efficient azide–alkyne click reaction is then used to cross-link the surface of the mixed micelle and functionalize them with monoazide 4. Free radical polymerization among the methacrylates of the cross-linkable surfactants and micelle-solubilized divinylbenzene (DVB) “solidifies” the micellar core around the template, creating the imprinted site. Meanwhile, the boroxole FM is turned into a binding group, covalently attached to the imprinted site by its polymerized vinyl group. Amphiphilicity of the template molecule helps to anchor the template and ultimately the imprinted site near the surface of the cross-linked micelle, facilitating template removal and binding of guest molecules afterwards.

Scheme 1.

Preparation of a MINP-cofactor complex to hydrolyze hexyl β-d-glucoside (6).

Templates 1a–f were synthesized in a one-step reaction from the corresponding alkoxyamine derivative (Scheme 2 in the Experimental Section).^38,39 The pyrenyl group is included in the template to produce a hydrophobic imprinted site in the final MINP(1a–f) for 1-pyrenecarboxylic acid (5), the acid cofactor to hydrolyze acetal.²⁰ Proper matching between the space-holder on the template for the cofactor and the final cofactor gives a strong driving force for 1-pyrenecarboxylic acid (5) to go inside the MINP and be positioned next to the glycosidic bond.⁴⁰ The cyan-colored moiety of the template is meant to mimic hexyl β-d-glucopyranoside (6), the substrate of hydrolysis, and had an aglycon varying in size and shape. Although it is difficult to control the dynamics of a cross-linked polymeric nanoparticle while maintaining integrity of the imprinted site, our idea here is to vary the size of the alkyl-binding pocket of the MINP to influence the dynamics of the bound substrate.

Scheme 2.

Synthesis of template 1a–f.

Formation of the imprinted site was characterized by isothermal titration calorimetry (ITC), one of the most reliable methods to study intermolecular interactions.⁴¹ As shown in Table 1, MINP(1a), i.e., the MINP prepared with 1a as the template, binds its template strongly, with a binding constant of K_a = 19.3 × 10⁵ M⁻¹ (entry 1). The strong binding—with nearly 9 kcal/mol of binding free energy—is derived from the boroxole FM that binds the 4,6-diol of a glucoside through reversible boronate bonds^21,34,36,42 and a large driving force for the pyrenyl group of the template to enter the pyrene-shaped imprinted site. Meanwhile, none of the other analogues (1b–f) shows any detectable binding to MINP(1a) (entries 2–6). Apparently, the larger/differently shaped hydrophobic groups on these structural analogues have difficulty fitting into the narrow, hexyl-shaped imprinted site from 1a. Exposure of a sizable hydrophobic group to water is known to be highly detrimental to binding,⁴³ even if the other parts of these analogues (i.e., glycan and pyrenyl) are identical to those of 1a.

Table 1.

ITC binding data of MINP(1a–f) for the templates and acid cofactor 5.^a

entry	MINP	guest	Ka (× 105 M−1)	ΔH (kcal/mol)	ΔS (cal/mol/K)	ΔG (kcal/mol)
1	MINP(1a)	1a	19.3 ± 2.41	−20.1 ± 0.67	−38.7	−8.56
2	MINP(1a)	1b	-b	-b	-b	-b
3	MINP(1a)	1c	-b	-b	-b	-b
4	MINP(1a)	1d	-b	-b	-b	-b
5	MINP(1a)	1e	-b	-b	-b	-b
6	MINP(1a)	1f	-b	-b	-b	-b
7	MINP(1b)	1b	28.0 ± 3.28	−10.7 ± 0.26	−6.23	−8.84
8	MINP(1c)	1c	33.2 ± 3.74	−22.5 ± 0.47	−45.8	−8.84
9	MINP(1d)	1d	34.5 ± 5.66	−43.1 ± 1.35	−114	−9.11
10	MINP(1e)	1e	41.3 ± 6.49	−35.0 ± 0.92	−87.0	−9.06
11	MINP(1f)	1a	9.54 ± 1.75	−13.5 ± 1.10	−17.8	−8.19
12	MINP(1f)	1b	13.6 ± 2.00	−62.7 ± 12.0	−182	−8.44
13	MINP(1f)	1c	5.73 ± 1.50	−27.7 ± 5.78	−66.6	−7.84
14	MINP(1f)	1d	5.24 ± 1.70	−14.7 ± 4.28	−23.2	−7.78
15	MINP(1f)	1e	-b	-b	-b	-b
16	MINP(1f)	1f	41.7 ± 5.44	−13.4 ± 0.29	−14.7	−9.02
17	MINP(1a)	5	10.6 ± 1.16	−2.61 ± 0.07	18.8	−8.22
18	MINP(1b)	5	11.4 ± 1.00	−2.95 ± 0.05	17.8	−8.26
19	MINP(1c)	5	13.4 ± 1.48	−1.89 ± 0.04	21.7	−8.36
20	MINP(1d)	5	10.7 ± 1.74	−22.0 ± 0.79	−46.3	−8.20
21	MINP(1e)	5	10.4 ± 1.49	−8.20 ± 0.25	0.04	−8.21
22	MINP(1f)	5	10.4 ± 1.21	−12.6 ± 0.30	−14.6	−8.25

^aTitrations were carried out in 10 mM citrate buffer pH = 5.0 at 298K. The errors between the runs were <10%.

^bBinding was extremely weak. The binding constant could not be determined accurately with ITC.

We also used MINP(1f)—the best co-catalyst (vide infra)—to further understand the binding selectivity of these materials. This particular MINP has a significantly wider pocket for the aglycon (of substrate 6) than MINP(1a). It is interesting that its 1-naphthyl-imprinted pocket was able to bind all the other templates (1a–1d) with an appreciable strength (K_a = 5–14 × 10⁵ M⁻¹) except 1e, which has a 2-naphtyl aglycon (Table 1, entries 11–15). Apparently, the linear hexyl of 1a by folding could occupy the imprinted site of MINP(1f),⁴⁴ in agreement with a previous study. The phenyl groups of 1b–1d also could fit in but the 2-naphthyl of 1e must have a wrong shape for MINP(1f), being relatively long and narrow.

Micellar imprinting, therefore, is extremely faithful in reproducing structural features of the template in the imprinted site. Table 1 shows that every MINP binds its own template strongly, with a binding energy of almost 9 kcal/mol (Table 1, entries 1, 7–10, and 16). Meanwhile, all the MINPs binds 1-pyrenecarboxylic acid (5) with a binding free energy of 8.2–8.3 kcal/mol (entries 17–22). Thus, the hydrophobic pyrenyl group contributes majorly to the binding of the templates. This feature is important because the substrate, hexyl β-d-glucopyranoside (6), also has an alkyl group. During the catalysis, the acid cofactor needs to compete effectively for its imprinted site, while the alkyl group stays in its designed place.

Correlation between Substrate Binding and Catalysis.

According to ITC, MINP(1a–f) binds the hydrolysis substrate, hexyl β-d-glucopyranoside (6), with a binding free energy of 5.7–6.4 kcal/mol in pH 5 citrate buffer (Table 2). The titrations were performed in the presence of 1 equivalent of 1-pyrenecarboxylic acid (5) in the solution, to mimic what would happen during the catalysis. Because catalytic hydrolysis requires much higher temperatures (vide infra), there was no concern for hydrolysis at room temperature used for the ITC titrations.

Table 2.

ITC binding data of MINP(1a–f) for substrate 6.^a

entry	MINP	guest	Ka (× 104 M−1)	ΔH (kcal/mol)	ΔS (cal/mol/K)	ΔG (kcal/mol)
1	MINP(1a)	6	4.60 ± 0.59	−0.38 ± 0.01	20.1	−6.37
2	MINP(1b)	6	2.45 ± 0.36	−0.82 ± 0.04	17.3	−5.98
3	MINP(1c)	6	2.11 ± 0.40	−0.55 ± 0.02	17.9	−5.89
4	MINP(1d)	6	2.19 ± 0.46	−1.15 ± 0.06	16.0	−5.92
5	MINP(1e)	6	1.69 ± 0.45	−1.64 ± 0.08	13.8	−5.75
6	MINP(1f)	6	1.58 ± 0.14	−0.51 ± 0.04	17.5	−5.73

^aTitrations were carried out in 10 mM citrate buffer pH = 5.0 at 298K, in the presence of 1 equivalent of 1-pyrenecarboxylic acid (5) in the solution. The errors between the runs were <10%.

Table 2 shows that the binding constant of the MINP for the substrate is the in the range of K_a = 1.6–4.6 × 10⁴ M⁻¹ and more or less decreases from MINP(1a) to MINP(1f). Hence, as the imprinted pocket for the aglycon becomes larger and more poorly matched with hexyl, a decrease in binding affinity is observed (Figure 1, red bars). This type of size exclusion is a sign of successful imprinting.^44,45 The results are also reasonable because an incompletely and poorly fitted hydrophobic pocket in water will inevitably have water molecules inside, creating unfavorable water–hydrocarbon contact as a result.^46,47

Figure 1.

Binding constants of different MINPs for hexyl β-d-glucopyranoside (6) at 25 °C in 10 mM citrate buffer (pH 5) and the hydrolytic yields of substrate 6 after 24 h at 95 °C in 10 mM citrate buffer (pH 5) catalyzed by the different MINPs and acid cofactor 5. [substrate] = 0.5 mM. [MINP(1f)] = 50 μM. [5] = 100 μM. Yields were determined by HPLC using calibration curves generated from authentic samples. The data points in yield are connected to guide the eye.

The absolute binding energy for the substrate by the six MINPs differed by up to 0.64 kcal/mol (Table 2). This difference was enough to change the hydrolysis of hexyl β-d-glucopyranoside (6) significantly. As shown in Figure 1 (blue curve), the hydrolytic yield for 6 went from 0% with MINP(1a) to 37% with MINP(1f) in the presence of 1-pyrenecarboxylic acid at 95 °C after 24 h. The change was dramatic given all the catalysts had a similar design and the binding for the acid-cofactor was practically the same (Table 1, entries 17–22).

There are at least three possible reasons for the different catalytic activities of the MINP-cofactor complexes. First, since the hexyl aglycon of the substrate is expected to fully occupy the imprinted pocket of MINP(1a), there is very little space for water in the active site, which is the nucleophile needed for the hydrolysis—the pyrene-shaped hydrophobic site of MINP(1a) is assumed to be fully occupied by the acid cofactor during catalysis due to its strong binding. Without a nearby water molecule(s), hydrolysis will not occur. This explanation, although reasonable for MINP(1a), cannot explain the consistently increased hydrolysis in the other MINPs. MINP(1e) and MINP(1f), for example, were both imprinted against an naphthyl derivative, with the only difference being the position of the substitution. Yet, the MINP prepared from the 1-substituted naphthyl template was clearly more active than the one prepared from the 2-substituted naphthyl template.

The second possible cause for the different activities in Figure 1 could be different rates in the binding of the substrate and release of the product. Differences in these rates, nonetheless, are unlikely to be an important factor in our case because the hydrolysis (24 h at 95 °C) is far slower than the reversible boronate bond formation and hydrophobically driven binding. During ITC titration, for example, the binding reached equilibrium within 120–180 s after each injection even at 25 °C (Figures S3–10).

We thus consider the most likely reason for the consistent increase of catalytic activity in Figure 1 is derived from higher mobility of the hexyl tail in the larger and wider alkyl-binding site. Since hydrolysis of an acetal by a nearby carboxylic acid happens through a general acid-catalyzed reaction,⁴⁸ the exocyclic oxygen needs to hydrogen-bond with the acid co-factor at certain point in the reaction. As the imprinted pocket for the alkyl tail gets larger and wider—e.g., from MINP(1b) to MINP(1c) to MINP(1d)—the hexyloxy group would enjoy more freedom in the alkyl-binding pocket, enabling its oxygen to better align itself with the carboxylic acid of the cofactor. If this was indeed the case, it is interesting that the hexyloxy group had different mobility in the two naphthyl-imprinted pockets of MINP(1e) and MINP(1f): once again, a wider pocket seems more amenable to the catalysis than the narrower and deeper one.

The inverse correlation between substrate binding/fitting and catalytic hydrolysis in Figure 1 was also observed when MINP(1b) and MINP(1f) were employed to hydrolyze benzyl β-d-glucopyranoside (7). In this case, the substrate was best matched with MINP(1b), which was imprinted from a benzyl-derived template. The substrate was bound strongly, with a K_a value of (1.03 ± 0.12) × 10⁵ M⁻¹ by MINP(1b) and (8.32 ± 1.26) × 10⁴ M⁻¹ by MINP(1f) (Figure S8). Yet, MINP(1b) was completely inactive in the hydrolysis of 7 while MINP(1f) gave a 10% yield under the same reaction conditions.

Selectivity in Hydrolysis.

The relatively large and wide active site of MINP(1f)–5 complex allowed it to hydrolyze a variety of alkyl glycosides (Figure 2). The lower activity of the benzyl derivative is in line with the mobility postulation mentioned above, although intrinsic reactivity could also be a factor. Methyl α- and β-d-glucosides (8 and 9) are both hydrolyzed, with little selectivity, consistent with the large active site that could easily accommodate the methyl group in either configuration. Methyl β-d-galactoside 10 differs from 9 by the inversion of a single hydroxyl. Its inactivity in the hydrolysis highlights the importance of the boroxole binding to the catalysis because the C4 hydroxyl used for the boronate bond formation is altered.

Figure 2.

Hydrolytic yields of substrates 6–10 after 24 h at 95 °C in 10 mM citrate buffer (pH 5) catalyzed by MINP(1f) and acid cofactor 5. [substrate] = 0.5 mM. [MINP(1f)] = 50 μM. [5] = 100 μM. Yields were determined by HPLC using calibration curves generated from authentic samples.

The convenience of molecular imprinting enabled us to tune the selectivity of our synthetic glycosidase readily. MINP(1e) has a relatively long and narrow pocket that should accommodate suitable aryl glycosides, whose hydrolysis can be easily monitored by UV-vis spectroscopy. Indeed, when the MINP(1e)–5 complex was employed to hydrolyze 11α/β–13α/β at 40 °C in pH 6.5 MES buffer, the most reactive substrates were the β-glucoside and β-mannoside, both having the same trans-4,6-diol as template 1e (Figure 3). The narrow pocket preferred the β-anomers roughly by a 3:1 ratio. The galactosides (13α and 13β) exhibited low activity, consistent with the importance of the C4 hydroxyl in the substrate binding. For the best substrate (11β), the MINP(1e)–5 complex gave a Michaelis constant of K_m = 168 μM and a catalytic turnover of k_cat = 3.67 × 10⁻³ s⁻¹ (Figure 4). The catalytic efficiency was thus k_cat/K_m = 22.4 M⁻¹s⁻¹.

Figure 3.

Pseudo first-order rate constants of substrate 11α/β–13α/β at 40 °C in 10 mM MES buffer (pH 6.5) catalyzed by MINP(1e) and acid cofactor 5. [substrate] = 0.10 mM. [MINP(1e)] = 5.0 μM. [5] = 10 μM. Hydrolysis was monitored by absorbance at 320 nm for the formation of the p-nitrophenol product.

Figure 4.

Michaelis-Menten plot for the hydrolysis of 11β by MINP(1e) and 5 in MES buffer (10 mM, pH = 6.5) at 40 °C. [MINP(1e)] = 5.0 μM. [5] = 10 μM.

Conclusions

Without sophisticated conformational control of enzymes, molecularly imprinted artificial enzymes cannot utilize dynamics of the active site directly to boost its catalysis. Tuning the dynamics of the polymer network through cross-linking density is also impractical, because the integrity of the imprinted site cannot be guaranteed. Molecular imprinting, however, affords a convenient way to tune the size and shape of the active site of a molecularly imprinted artificial enzyme and indirectly modulates the dynamics of the bound substrate. Our data shows that this approach could be a powerful strategy to enhance catalysis, able to revive an otherwise completely incompetent catalyst—i.e., MINP(1a)+5—to one that displayed moderate activity for a highly challenging alkyl glycoside (6). Given that a single noncovalently bound carboxylic acid is involved in the hydrolysis, the improvement in catalysis is significant. Enzymes use numerous strategies to achieve extraordinary catalytic efficiency and selectivity,^49–51 it is encouraging that some of these features can be mimicked in readily prepared synthetic materials. One noteworthy benefit of these molecularly imprinted nanoparticles is their excellent thermal, solvent, and pH stability, due to their highly cross-linked polymeric nature.

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 ppm relative to residual solvent peaks. Coupling constants are reported in hertz. ESI-HRMS were recorded on 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. Dynamic light scattering (DLS) data were collected 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). UV-vis spectra were recorded on a Cary 100 Bio UV-visible spectrophotometer.

Syntheses of FM 2,³⁴ surfactant 3a,³⁴ surfactant 3b,³⁴ surface ligand 4,³⁴ and O-(1-pyrenylmethyl)hydroxylamine (14)⁵² were reported previously.

Pyrenecarboxylic acid 5 (97% purity) was purchased from TCI America and substrate 6 (98% purity) was purchased from Sigma-Aldrich.

General procedure for the synthesis of compounds 15a–f.

The appropriate aldehyde (1 mmol, 0.5 M, 1.0 equiv) was added to a solution of compound 14 (0.25 g, 1 mmol, 1.0 equiv) in ethanol (2 mL). After the reaction mixture was stirred at room temperature overnight, solvent was removed by rotary evaporation. The residue was dissolved in 1 mL of dry THF and 1 mL of acetic acid, followed by the addition of borane-pyridine complex (200 μL, 2.2 mmol, 2.2 equiv). After the reaction mixture stirred at room temperature for 6 h, the reaction was quenched by the addition of 6 M HCl solution dropwise. The mixture was poured into 20 mL of water and extracted with dichloromethane (3 × 15 mL). The combined organic layer was dried over sodium sulfate and concentrated by rotary evaporation. The residue was purified by flash chromatography over silica gel to afford the final product.

Compound 15a.

The product was purified by flash chromatography over silica gel using 10:1 hexane/ethyl acetate as the eluent to afford the final product as a yellow oil (0.21 g, 62%). ¹H NMR (400 MHz, 298K, CDCl₃) δ 8.43 (d, J = 9.2 Hz, 1H), 8.15 (ddd, J = 15.2, 7.7, 5.8 Hz, 4H), 8.07 – 7.97 (m, 4H), 5.61 (s, 2H), 3.07 (t, J = 5.4 Hz, 2H), 1.63 (p, J = 7.4 Hz, 2H), 1.30 – 1.18 (m, 6H), 0.82 (t, J = 6.8 Hz, 3H). ¹³C{¹H} (100 MHz, 298K, CDCl₃) δ 131.7, 131.1, 130.7, 129.8, 128.2, 128.1, 127.98, 127.6, 127.3, 126.2, 125.9, 125.3, 124.8, 124.6, 124.5, 123.4, 74.6, 51.6, 31.5, 29.4, 26.7, 22.5, 13.9. HRMS (ESI) m/z: Calcd for C₂₃H₂₆NO m/z: [M+H]⁺ 332.2009; Found 332.2042.

Compound 15b.

The product was purified by flash chromatography over silica gel using 10:1 hexane/ethyl acetate as the eluent to afford the final product as a yellow powder (0.22 g, 64%). ¹H NMR (400 MHz, 298K, CDCl₃) δ 8.26 (d, J = 9.2 Hz, 1H), 8.23 – 8.17 (m, 2H), 8.14 (d, J = 7.7 Hz, 1H), 8.11 – 7.97 (m, 5H), 7.44 – 7.29 (m, 5H), 5.38 (s, 2H), 4.10 (s, 2H). ¹³C{¹H} (100 MHz, 298K, CDCl₃) δ 137.8, 131.5, 131.2, 130.9, 130.8, 129.8, 129.1, 128.4, 128.0, 127.6, 127.5, 127.5, 127.4, 125.9, 125.2, 124.9, 124.7, 124.5, 123.7, 74.8, 56.7. HRMS (ESI⁺/QTOF) Calcd for C₂₄H₂₀NO m/z: [M+H]⁺ 338.1539; Found 338.1551.

Compound 15c.

The product was purified by flash chromatography over silica gel using 10:1 hexane/ethyl acetate as the eluent to afford the final product as a yellow powder (0.22 g, 59%). ¹H NMR (600 MHz, 298K, CDCl₃) δ 8.42 (d, J = 9.2 Hz, 1H), 8.26 – 8.21 (m, 2H), 8.17 (dd, J = 17.4, 8.4 Hz, 2H), 8.08 – 8.03 (m, 2H), 7.27 (t, J = 7.7 Hz, 2H), 6.93 (td, J = 7.4, 1.0 Hz, 1H), 6.85 (d, J = 8.0 Hz, 1H), 5.48 (s, 2H), 4.13 (s, 2H), 3.78 (s, 3H). ¹³C{¹H} (150 MHz, 298K, CDCl₃) δ 157.8, 131.4, 131.3, 131.1, 130.9, 129.7, 128.8, 127.9, 127.6, 127.5, 127.4, 125.9, 125.4, 125.2, 124.9, 124.8, 124.5, 123.8, 120.5, 110.3, 74.3, 55.2, 52.3. HRMS (ESI⁺/QTOF) Calcd for C₂₅H₂₂NO₂ m/z: [M+H]⁺ 368.1645; Found 368.1649.

Compound 15d.

The product was purified by flash chromatography over silica gel using 10:1 hexane/ethyl acetate as the eluent to afford the final product as a yellow powder (0.26 g, 66%). ¹H NMR (400 MHz, 298K, CDCl₃) δ 8.45 (d, J = 9.2 Hz, 1H), 8.24 – 8.10 (m, 4H), 8.10 – 7.97 (m, 4H), 7.16 (t, J = 8.3 Hz, 1H), 6.50 (d, J = 8.3 Hz, 2H), 5.50 (s, 2H), 4.28 (s, 2H), 3.74 (s, 6H). ¹³C{¹H} (100 MHz, 298K, CDCl₃) δ 159.0, 131.4, 131.3, 131.2, 130.9, 129.7, 128.8, 127.8, 127.5, 127.4, 127.3, 125.8, 125.1, 125.1, 124.9, 124.8, 124.5, 123.9, 113.0, 103.7, 73.9, 55.7, 44.5. HRMS (ESI⁺/QTOF) Calcd for C₂₆H₂₄NO₃ m/z: [M+H]⁺ 398.1751; Found 398.1782.

Compound 15e.

The product was purified by flash chromatography over silica gel using 10:1 hexane/ethyl acetate as the eluent to afford the final product as a yellow powder (0.28 g, 72%). ¹H NMR (600 MHz, 298K, CDCl₃) δ 8.22 – 8.12 (m, 4H), 8.10 – 7.98 (m, 4H), 7.87 (d, J = 9.2 Hz, 1H), 7.83 – 7.77 (m, 4H), 7.54 – 7.46 (m, 3H), 6.04 (s, 1H), 5.38 (s, 2H), 4.25 (s, 2H). ¹³C{¹H} (150 MHz, 298K, CDCl₃) δ 135.3, 133.4, 131.5, 131.2, 130.9, 130.8, 129.8, 128.1, 128.0, 127.9, 127.8, 127.6, 127.6, 127.5, 127.4, 127.0, 125.9, 125.8, 125.7, 125.2, 125.1, 124.9, 124.4, 123.7, 74.9, 56.9. HRMS (ESI⁺/QTOF) Calcd for C₂₈H₂₂NO m/z: [M+H]⁺ 388.1696; Found 388.1683.

Compound 15f.

The product was purified by flash chromatography over silica gel using 10:1 hexane/ethyl acetate as the eluent to afford the final product as a yellow powder (0.27 g, 70%). ¹H NMR (600 MHz, 298K, CDCl₃) δ 8.36 (d, J = 9.2 Hz, 1H), 8.23 (t, J = 7.0 Hz, 2H), 8.16 – 8.03 (m, 6H), 7.99 (d, J = 7.7 Hz, 1H), 7.95 (d, J = 8.4 Hz, 1H), 7.82 (d, J = 8.1 Hz, 1H), 7.76 (d, J = 8.2 Hz, 1H), 7.47 (t, J = 6.3 Hz, 1H), 7.43 – 7.36 (m, 2H), 5.41 (s, 2H), 4.52 (s, 2H). ¹³C{¹H} (150 MHz, 298K, CDCl₃) δ 133.8, 132.7, 132.1, 131.5, 131.3, 131.1, 130.8, 129.8, 128.5, 128.4, 128.3, 127.8, 127.6, 127.5, 127.4, 126.0, 125.9, 125.6, 125.3, 125.2, 124.9, 124.8, 124.4, 124.1, 123.8, 74.8, 54.6. HRMS (ESI⁺/QTOF) Calcd for C₂₈H₂₂NO m/z: [M+H]⁺ 388.1696; Found 388.1659.

General procedure for the synthesis of compounds 1a–f.

Compound 15a–f (0.5 mmol, 0.1 M, 1.0 equiv) and glucose (0.108 g, 0.6 mmol, 1.2 equiv) were dissolved in a mixture of 4 mL of THF and 1 mL of acetic acid. After the reaction mixture was stirred at room temperature for 24 h, solvents were by rotary evaporation. The residue was purified by flash chromatography over silica gel to afford the final product.

Compound 1a.

The product was purified by flash chromatography over silica gel using 10:1 dichloromethane/methanol as the eluent to afford the final product as a white powder (0.15 g, 61%). ¹H NMR (400 MHz, 298K, CD₃OD) δ 8.45 (d, J = 9.3 Hz, 1H), 8.25 – 8.15 (m, 4H), 8.11 – 8.05 (m, 3H), 8.02 (t, J = 7.6 Hz, 1H), 5.59 – 5.47 (m, 2H), 4.20 (d, J = 9.0 Hz, 1H), 3.79 (dd, J = 12.2, 2.2 Hz, 1H), 3.71 – 3.60 (m, 2H), 3.43 (t, J = 8.9 Hz, 1H), 3.35 – 3.31 (m, 1H), 3.26 – 3.19 (m, 1H), 3.04 – 2.95 (m, 2H), 1.52 (p, J = 7.5 Hz, 2H), 1.27 (p, J = 7.1 Hz, 2H), 1.21 – 1.07 (m, 4H), 0.77 (t, J = 6.9 Hz, 3H). ¹³C{¹H} (100 MHz, 298K, CD₃OD) δ 131.7, 131.2, 130.8, 130.2, 129.9, 128.6, 127.5, 127.3, 127.0, 125.8, 125.0, 125.0, 124.5, 124.4, 124.3, 123.3, 92.9, 78.3, 78.2, 74.0, 70.5, 69.8, 61.3, 52.7, 31.4, 27.2, 26.7, 22.1, 12.9. HRMS (ESI⁺/QTOF) Calcd for C₂₉H₃₆NO₆ m/z: [M+H]⁺ 494.2537; Found 494.2560.

Compound 1b.

The product was purified by flash chromatography over silica gel using 10:1 dichloromethane/methanol as the eluent to afford the final product as a yellow powder (0.14 g, 57%). ¹H NMR (400 MHz, 298K, CD₃OD) δ 8.19 – 8.13 (m, 2H), 8.07 (d, J = 7.8 Hz, 1H), 8.05 – 7.95 (m, 4H), 7.85 (dd, J = 8.5, 3.5 Hz, 2H), 7.56 – 7.52 (m, 2H), 7.43 – 7.38 (m, 2H), 7.37 – 7.34 (m, 1H), 5.31 (d, J = 10.1 Hz, 1H), 5.23 (d, J = 10.1 Hz, 1H), 4.24 (d, J = 13.0 Hz, 1H), 4.19 – 4.11 (m, 2H), 3.85 – 3.79 (m, 1H), 3.77 (d, J = 8.9 Hz, 1H), 3.65 (dd, J = 12.2, 5.2 Hz, 1H), 3.41 (t, J = 8.9 Hz, 1H). ¹³C{¹H} (100 MHz, 298K, CD₃OD) δ 137.4, 131.6, 131.1, 130.7, 129.9, 129.9, 129.5, 128.7, 128.0, 127.4, 127.3, 127.3, 126.9, 125.7, 124.9, 124.9, 124.3, 124.2, 124.2, 123.4, 92.5, 78.3, 74.3, 70.5, 69.7, 61.3, 56.9, 53.4. HRMS (ESI⁺/QTOF) Calcd for C₃₀H₃₀NO₆ m/z: [M+H]⁺ 500.2068; Found 500.2042.

Compound 1c.

The product was purified by flash chromatography over silica gel using 10:1 dichloromethane/methanol as the eluent to afford the final product as a yellow powder (0.14 g, 52%). ¹H NMR (600 MHz, 298K, CDCl₃) δ 8.23 – 8.17 (m, 3H), 8.14 – 8.01 (m, 5H), 7.80 (d, J = 7.7 Hz, 1H), 7.17 (dd, J = 7.3, 1.7 Hz, 1H), 7.00 (t, J = 7.7 Hz, 1H), 6.72 – 6.63 (m, 2H), 5.22 (d, J = 11.8 Hz, 1H), 5.09 (d, J = 11.8 Hz, 1H), 4.28 – 4.20 (m, 2H), 4.01 – 3.93 (m, 2H), 3.85 (dd, J = 11.9, 5.0 Hz, 1H), 3.82 (s, 3H), 3.57 – 3.50 (m, 1H), 3.50 – 3.42 (m, 2H), 3.40 – 3.33 (m, 1H). ¹³C{¹H} (150 MHz, 298K, CDCl₃) δ 157.6, 131.7, 131.7, 131.2, 130.7, 130.0, 129.9, 129.0, 128.9, 127.8, 127.7, 127.3, 126.0, 125.3, 125.30, 124.8, 124.8, 124.7, 124.2, 123.6, 120.4, 110.2, 92.8, 75.0, 73.3, 70.7, 70.7, 62.6, 55.3, 49.9, 29.7. HRMS (ESI⁺/QTOF) Calcd for C₃₁H₃₂NO₇ m/z: [M+H]⁺ 530.2173; Found 530.2183.

Compound 1d.

The product was purified by flash chromatography over silica gel using 10:1 dichloromethane/methanol as the eluent to afford the final product as a yellow powder (0.14 g, 51%). ¹H NMR (600 MHz, 298K, CDCl₃) δ 8.30 (d, J = 9.2 Hz, 1H), 8.22 (dd, J = 7.6, 2.5 Hz, 2H), 8.13 (dd, J = 8.4, 6.9 Hz, 2H), 8.09 (d, J = 8.9 Hz, 1H), 8.07 – 8.03 (m, 2H), 7.92 (d, J = 7.6 Hz, 1H), 7.14 (t, J = 8.5 Hz, 1H), 6.50 (d, J = 8.5 Hz, 2H), 5.40 (d, J = 11.4 Hz, 1H), 5.31 (d, J = 11.4 Hz, 1H), 4.19 (d, J = 12.0 Hz, 1H), 3.87 – 3.81 (m, 7H), 3.73 – 3.68 (m, 1H), 3.66 – 3.60 (m, 1H), 3.58 – 3.53 (m, 2H), 3.41 – 3.35 (m, 2H), 3.14 – 3.06 (m, 1H), 2.95 (dd, J = 13.7, 3.3 Hz, 1H). ¹³C{¹H} (150 MHz, 298K, CDCl₃) δ 159.1, 131.6, 131.2, 130.7, 130.6, 129.9, 129.2, 128.5, 127.8, 127.6, 127.4, 126.0, 125.3, 125.2, 124.9, 124.7, 124.4, 123.6, 112.6, 103.7, 74.5, 72.7, 71.6, 71.5, 64.0, 60.4, 55.7, 50.9, 50.1. HRMS (ESI⁺/QTOF) Calcd for C₃₂H₃₄NO₈ m/z: [M+H]⁺ 560.2279; Found 560.2246.

Compound 1e.

The product was purified by flash chromatography over silica gel using 10:1 dichloromethane/methanol as the eluent to afford the final product as an off white powder (0.15 g, 55%). ¹H NMR (400 MHz, 298K, CD₃OD) δ 8.09 (d, J = 7.3 Hz, 1H), 8.01 – 7.89 (m, 5H), 7.86 – 7.70 (m, 6H), 7.54 – 7.44 (m, 4H), 5.25 (d, J = 10.2 Hz, 1H), 5.17 (d, J = 10.2 Hz, 1H), 4.36 (d, J = 13.1 Hz, 1H), 4.31 – 4.21 (m, 2H), 3.86 (dd, J = 12.2, 2.2 Hz, 1H), 3.81 (t, J = 8.9 Hz, 1H), 3.69 (dd, J = 12.2, 5.3 Hz, 1H), 3.36 (t, J = 9.2 Hz, 1H), 3.25 (ddd, J = 9.6, 5.3, 2.2 Hz, 1H). ¹³C{¹H} (100 MHz, 298K, CD₃OD) δ 149.0, 135.0, 133.5, 133.1, 131.5, 131.1, 130.5, 129.9, 129.4, 128.7, 128.7, 127.7, 127.5, 127.3, 127.3, 127.1, 126.8, 125.6, 125.6, 125.5, 124.8, 124.7, 124.2, 124.1, 124.0, 123.3, 92.9, 78.4, 78.3, 74.1, 70.6, 69.8, 61.3, 57.0. HRMS (ESI⁺/QTOF) Calcd for C₃₄H₃₂NO₆ m/z: [M+H]⁺ 550.2224; Found 550.2251.

Compound 1f.

The product was purified by flash chromatography over silica gel using 10:1 dichloromethane/methanol as the eluent to afford the final product as an off white powder (0.16 g, 58%). ¹H NMR (600 MHz, 298K, DMSO-d₆) δ 8.33 – 8.30 (m, 2H), 8.28 (d, J = 7.7 Hz, 1H), 8.23 (d, J = 9.2 Hz, 1H), 8.21 – 8.16 (m, 3H), 8.09 (t, J = 7.6 Hz, 1H), 7.99 (t, J = 7.8 Hz, 2H), 7.88 (dd, J = 17.4, 8.2 Hz, 2H), 7.62 (d, J = 6.9 Hz, 1H), 7.48 (t, J = 7.6 Hz, 2H), 7.36 (ddd, J = 8.2, 6.7, 1.3 Hz, 1H), 4.97 – 4.86 (m, 2H), 4.62 (d, J = 12.2 Hz, 1H), 4.46 (d, J = 12.2 Hz, 1H), 3.87 (d, J = 8.9 Hz, 1H), 3.82 – 3.76 (m, 1H), 3.70 – 3.64 (m, 1H), 3.53 – 3.48 (m, 1H), 3.16 (t, J = 8.5 Hz, 1H), 3.09 – 2.98 (m, 2H). ¹³C{¹H} (150 MHz, 298K, CDCl₃) δ 133.6, 132.3, 131.9, 131.6, 131.0, 130.5, 129.8, 129.5, 129.0, 128.6, 128.5, 127.9, 127.6, 127.2, 126.0, 125.8, 125.7, 125.2, 125.2, 124.6, 124.4, 124.3, 123.4, 91.9, 77.7, 74.3, 70.1, 69.8, 62.0, 54.8, 53.4. HRMS (ESI⁺/QTOF) Calcd for C₃₄H₃₂NO₆ m/z: [M+H]⁺ 550.2224; Found 550.2244.

Preparation of MINP.

A typical procedure is as follows. A solution of 6-vinylbenzoxaborole (2) in methanol (10 μL of a 6.4 mg/mL, 0.0004 mmol) was added to template 1a in methanol (10 μL of 19.7 mg/ml, 0.0004 mmol) in a vial with 5 ml methanol. The mixture was stirred for 6 h at room temperature before the methanol was removed in vacuo. A micellar solution of compound 3a (0.03 mmol) and compound 3b (0.02 mmol) in H₂O (2.0 mL) was then added to the template-boronate complex, followed by divinylbenzene (DVB, 2.8 μL, 0.02 mmol) and 2,2-dimethoxy-2-phenylacetophenone (DMPA,10 μL of a 12.8 mg/ml solution in DMSO, 0.0005 mmol). The mixture was subjected to ultrasonication for 10 min until the solution became clear. CuCl₂ in H₂O (10 μL of a 6.7 mg/mL solution, 0.0005 mmol), and sodium ascorbate in H₂O (10 μL of a 99 mg/mL solution, 0.005 mmol) were added and the reaction mixture was stirred slowly at room temperature for 12 h. The reaction mixture was degassed with N₂ for 15 min, sealed with a rubber stopper, and irradiated in a Rayonet reactor for 12 hours. Compound 4 in H₂O (10 μL, 0.024 mmol), CuCl₂ in H₂O (10 μL of a 6.7 mg/mL solution, 0.0005 mmol), and sodium ascorbate in H₂O (10 μL of a 99 mg/mL solution, 0.005 mmol) were added to the previous solution. After being stirred for additional 8 h, the reaction mixture was then poured and precipitated into acetone (8 mL). The precipitate was collected by centrifugation, washed with a mixture of acetone/water (5 mL/1 mL) and methanol/acetic acid (5 mL/0.1 mL) for three times to remove the template. The MINP precipitate was rinsed with methanol (5 mL) and finally with acetone (5 mL) to neutral before being dried in air to afford the final MINP catalyst as an off-white solid. Typical yields were >80%.

Determination of Binding Constants by ITC.

In general, a solution of an appropriate guest in 10 mM Citric buffer (pH 5.0) with 2.5% 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 site 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 auto generated after curve fitting using Microcal Origin 7. All titrations were performed in duplicates and the errors between the runs were <10% using sodium salts of the templates and acid cofactors.

Catalytic hydrolysis of alkyl glycosides.

Stock solutions of the appropriate glycoside (5.0 mM), MINP (200 μM), and 1-pyrenecarboxylic acid (5) (500 μM) in 10 mM citrate buffer (pH 5.0) were prepared. For each hydrolysis, a 50 μL aliquot of the glycoside solution, 125 μL of the MINP solution, and 100 μL of the acid cofactor solution were mixed and diluted with the same buffer to a final volume of 0.50 mL. The reaction mixture was kept at 90 °C for 24 h in a block heater. Hydrolysis was monitored by LC-MS analysis using an Agilent 1200 Series Binary VWD system with an Agilent 6540 UHD Accurate Mass Q-TOF mass detector. Separation of the products was performed on a Thermo Scientific HILIC-LC column (4.6 mm, 150 mm) at 60 °C. For quantitative analysis, injection volumes were adjusted for the signal intensity to stay within the linear range of the calibration curves (Figure S9–12). All samples were centrifuged at 20,000 RPM before analysis to remove the MINP particles (to avoid column blockage over extended usage).

Catalytic hydrolysis of p-nitrophenyl glycosides.

Stock solutions (5 mM) of p-nitrophenyl glycopyranosides (11–13α and 11–13β) in 10 mM MES buffer pH 6.0 were prepared. The stock solutions were stored in a refrigerator and used within 3 days. A stock solution of catalyst containing MINP(1f) (200 μM) and 1-pyrenecarboxylic acid (500 μM) in 10 mM MES buffer pH 6.0 were prepared. For the kinetic experiments, a typical procedure is as follows. An aliquot of 50 μL of the MINP(1f) stock solution and 40 μL of the 1-pyrenecarboxylic acid stock solution were combined with 1870 μL of 10 mM MES buffer pH 6.0 in a cuvette. The cuvette was vortexed gently before it was placed in a UV/Vis spectrometer and equilibrated to 40.0 °C for 10 min. Then an aliquot of 40 μL of the substrate stock solution was added. The hydrolysis was monitored by the absorbance of p-nitrophenol at 320 nm. The concentration of p-nitrophenol or p-nitrophenoxide was calculated based on the calibration curve (Figure S13) obtained from authentic samples.