Toward Design Rules for Enzyme Immobilization in Hierarchical Mesoporous Metal-Organic Frameworks
Metal-organic frameworks (MOFs) are porous, crystalline materials comprised of metal nodes and organic linkers. Here, a Zr-based MOF named NU-1000 is used to encapsulate and protect an enzyme. The encapsulation and subsequent protection of enzymes in solid supports is important for the potential industrialization of enzymes as chemical catalysts. NU-1000 is shown to be capable of stabilizing the enzyme under harsh conditions, and in addition, the encapsulated enzyme is shown to maintain full functionality.
INTRODUCTION<!>Cutinase Immobilization<!>Cutinase Accessibility in NU-1000<!>Activity and Stability of Immobilized Cutinase<!>Comparing the Accessibility of Cutinase Immobilized in NU-1000 versus Other MOF Supports<!>Conclusions<!>EXPERIMENTAL PROCEDURES Materials<!>Measurements of Gas Adsorption<!>Scanning Electron Microscopy<!>Electrospray Ionization Mass Spectrometry<!>Measurement of Zeta Potential<!>ICP-OES Analysis<!>General Procedure for Kinetic Study of the Hydrolysis Reaction<!>Stability and Recyclability Test<!>Labeling Cutinase with Fluorescent Dye<!>CLSM Analysis<!>Titration of Active Sites for Cutinase and Immobilized Cutinase in MOFs<!>SUPPLEMENTAL INFORMATION<!>AUTHOR CONTRIBUTIONS
<p>Enzyme-mediated catalysis is a practical, sustainable, and environmentally benign strategy for the production of industrially relevant chemicals ranging from biofuels to pharmaceuticals and food additives. 1 Advances in protein engineering have led to enzymes with enhanced catalytic performance, yet their use industrially is often hampered by the lack of long-term stability, recyclability, and efficient recovery. 2 These issues can be overcome by using a solid support to immobilize the enzymes and shield them from deactivating reaction conditions (e.g., organic solvents 3 and denaturants 4 ) in addition to providing potential for enzyme recyclability via recovery of the heterogenized catalyst and support.</p><p>Metal-organic frameworks (MOFs) are promising candidates for enzyme immobilization. [5][6][7][8][9][10][11] MOFs 12 are a class of highly tunable, porous materials that have shown promise in a wide range of potential applications including gas storage and release, [13][14][15] light harvesting and energy conversion, [16][17][18] drug delivery, 19 and catalysis. 20 MOFs are potentially superior to other porous materials commonly used to immobilize proteins and enzymes such as sol gels, 21 zeolites, 22 and mesoporous silica supports, [23][24][25] because MOFs are finely tunable and crystalline, thus exhibiting uniformity and long-range ordering from the atomic to microscale regime. 12 Enzyme immobilization in less uniform solid supports typically leads to low protein-loading efficiency, 22,26 low stability at elevated temperatures, and/or enzyme leaching. 27,28 Initial studies have shown that some of these challenges can be overcome by using</p><p>The Bigger Picture Metal-organic frameworks (MOFs), which are simplistically viewed as molecular sponges, have garnered widespread media attention in recent years because of their potential in many popular applications, including naturalgas-powered vehicles. MOFs are porous yet robust materials that offer an interesting platform for encapsulating and protecting enzymes from degradation. Enzymes are nature's catalysts, and the delicate structure of enzymes can be understood by considering the irreversibility of frying an egg. In order to apply enzymes broadly as catalysts in industry, the need to protect them from denaturation under harsh conditions is important. Here, we show the applicability of a robust MOF that has structural features that make it intriguing for enzyme encapsulation. The results are important because the enzyme is protected and maintains high enzymatic activity. The long-term goal is to encapsulate specific enzymes that can be used for the detoxification of chemical-warfare agents.</p><p>MOFs for enzyme immobilization. [5][6][7][8][9][10][11] These initial studies focused on placing enzymes in cage-like materials without emphasis on the spatial distribution, accessibility, or conformation of the encapsulated enzyme or the diffusion of reactants and products throughout a given support. These characteristics coupled with composite stability, however, are critical design elements of an optimal MOF bioreactor. 21 We reasoned that an ideal enzyme support should present a hierarchical pore structure with large pores for enzyme immobilization and small pores for reactant/ product diffusion. We have recently described the preparation and characterization of the water-stable, mesoporous, zirconium-based MOF NU-1000, 29 which features hexagonal channels with a diameter of 3.1 nm as well as triangular channels with an edge length of 1.5 nm and windows connecting the two channels (Figure 1). We reasoned that an MOF with this unusual hierarchical pore structure would be especially suitable for the encapsulation of Fusarium solani pisi cutinase (PDB: 1CEX), an esterase that has shown promise as a biocatalyst in the preparation of aliphatic esters. [30][31][32][33] The sizes of the larger channels of NU-1000 match that of cutinase, an ellipsoid-shaped protein featuring a small-axis length of $3.0 nm. 34 These features, in addition to the high chemical (at pH 1-11) 35 and thermal stability (at >450 C) 29 of NU-1000, make this MOF a promising candidate material for use as a solid support in biocatalysis.</p><!><p>We treated 5-mm-long activated crystals of the MOF (NU-1000-5 mm) with a Trisbuffered saline (TBS) solution of cutinase (100 mM, pH 7.4) at 25 C (Figure 1). The uptake of cutinase by NU-1000-5 mm was determined using UV-visible spectroscopy, and a maximum loading of 5 mmol/g was reached after $75 hr (Figure S1). The solid sample (hereafter denoted as cutinase@NU-1000-5 mm) was then washed five times with detergent-containing buffer solution to ensure full removal of any loosely bound cutinase. Inductively coupled plasma-optical emission spectroscopy (ICP-OES) of the washed cutinase@NU-1000-5 mm sample also revealed a cutinase uptake of 5 mmol/g. The powder X-ray diffraction (PXRD) pattern of NU-1000-5 mm before and after cutinase immobilization confirmed that bulk crystallinity was retained after cutinase encapsulation (Figure 2A). Cutinase@NU-1000-5 mm exhibited a type IV N 2 adsorption-desorption isotherm, much like NU-1000-5 mm itself, but not surprisingly, the MOF exhibited a lower N 2 uptake capacity when cutinase was present in the pores (Figure 2B). The density functional theory pore-size distribution analysis of activated NU-1000-5 mm and cutinase@NU-1000-5 mm showed the two different pores in the hierarchical structure (Figure 2C). The pore volume corresponding to the triangular channels of NU-1000-5 mm dropped from 0.50 to 0.44 cm 3 /g, whereas the pore volume corresponding to the hexagonal channels dropped from 0.75 to 0.22 cm 3 /g after cutinase encapsulation.</p><p>To confirm that cutinase was internalized within NU-1000 and not simply adsorbed on the external surface or within defects of the crystal, we used in situ confocal laser scanning microscopy (CLSM) to image the cutinase encapsulation process with an AlexaFluor 647-labeled enzyme (Cut647) (Figure S1). To study the effect of crystal length on enzyme diffusion, we synthesized NU-1000 crystals with an average length of 10 and 1.5 mm (Figures 3A and 3B). The crystals were immersed in a 50 mM solution of Cut647, and 2D (xy) fluorescence intensity profiles of the MOF samples were obtained at a fixed z depth corresponding to the center layer of the crystal. These images (Figures 3C and 3D) indicate that the dye-labeled enzyme is in fact directed to the center of NU-1000 and that by decreasing the size of the particle from 10 to 1.5 mm, the time required to reach cutinase saturation is decreased by approximately 10-fold. We also modeled the diffusion process using Fick's second law in one dimension (Figures 3E and 3F). The best-fit value for the diffusion coefficient for Cut647 within NU-1000 was 2 3 10 À13 cm 2 /s, which is consistent with diffusive transport through the crystalline channels; the coefficient was found to be identical for 10 and 1.5 mm NU-1000 crystals. For comparison, the diffusion coefficient of cutinase on a 2D trimyristin support was found to be 8.0 3 10 À10 cm 2 /s. 36 These results suggest a strong interaction between Cut647 and NU-1000. In order to explain the nature of this strong interaction, we examined the zeta potential of NU-1000 with varying pH (Figure S2), which indicated an isoelectric point at approximately pH 4.3. This isoelectric point is consistent with our previous study on the Brønsted acidity of NU-1000. 37 Cutinase is known to have an isoelectric point at approximately pH 7.8. 38 Thus, in pH 7 buffer solution, cutinase and NU-1000 are positively and negatively charged, respectively. The strong interaction between Cut647 and NU-1000 can therefore be attributed to columbic forces.</p><p>We also used molecular mechanics calculations to computationally introduce a reported crystal structure of F. solani pisi cutinase into the hexagonal channels of NU-1000. The results indicated that in order to infiltrate the MOF, the long axis of the enzyme must orient along the mesoporous channel of NU-1000. In addition, the shape of cutinase changes slightly to allow for diffusion into the channels of NU-1000, resulting in an ellipsoid that is more elongated than the original cutinase structure (Figures 4A and S3). These models qualitatively support the small diffusion coefficients observed, as the enzyme must deform slightly to fit within the mesopore. In addition, the models indicate that the catalytically active site in cutinase, composed of amino acid residues Ser126, Asp180, and His194, 39 should remain accessible after encapsulation, with no indication of blocking by any framework components.</p><!><p>One property of MOF-encapsulated enzymes that has yet to be explicitly addressed is the permeability of the crystals to small-molecule reactants after enzyme immobilization. We reasoned that the unique hierarchical pore structure of NU-1000 would allow molecules of appropriate dimensions to diffuse into the interior of the crystal, even after enzyme encapsulation, either longitudinally via the triangular pore or laterally via the crystal windows. Molecules larger than either of these apertures should be excluded from the crystal due to blocking of the larger channels by cutinase. To test this idea, we chose two dye molecules (Figure 4B), one with dimensions small enough to diffuse through the smaller apertures (fluorescein: 1.0 3 0.8 3 0.5 nm, l ex = 488 nm, l em = 512 nm) and one too large to do so (4,4 0 ,4 00 ,4%-(porphine-5,10,15,20-tetrayl)tetrakis(benzoic acid): 1.6 3 1.6 3 0.2 nm, l ex = 405 nm, l em = 640 nm), and compared the ability of each dye to permeate empty versus Cut647-loaded NU-1000-10 mm using CLSM. Both dyes have fluorescence emission maxima that lie well outside those of AlexaFluor 647. For empty NU-1000-10 mm, micrographs of samples immersed in solutions of fluorescein and porphyrin showed that both dyes can freely permeate the crystal (Figure S4). In the case of Cut647@NU-1000-10 mm, merged 3D reconstructions of z stacks from the 633 nm (dye-labeled cutinase) and 488 nm (fluorescein) laser channels showed the presence of Cut647 and that fluorescein retains access to the whole of the matrix (Figures 4C  and S4). In the case of the larger porphyrin dye (UV 405 nm laser channel), we observed a size-exclusion effect whereby the molecule was blocked from entering the interior of the crystal and resided primarily on the surface (Figure 4D). This demonstrates that although the large hexagonal channels of NU-1000 are blocked by encapsulated cutinase, a reactant small enough to fit into the smaller triangular channels can still freely diffuse in and out of the framework.</p><p>Given that our computational model suggests that the active site of cutinase remains accessible after immobilization, we performed a real-time in situ CLSM experiment on cutinase@NU-1000-10 mm after the addition of a small aliquot of resorufin butyrate to determine whether the enzyme is accessible and active. 40 Enzymatic hydrolysis of resorufin butyrate by cutinase produces resorufin (l ex = 500 nm, l em = 593 nm), a highly fluorescent dye that can be used to monitor the catalytic activity of the encapsulated enzyme. When resorufin butyrate was added to a solid sample of cutinase@NU-1000-10 mm, the crystals of the MOF rapidly began to fluoresce at 593 nm throughout the crystal, indicating not only that the reactant reached the interior of the matrix but also that the encapsulated cutinase was accessible and reactive there. As the reaction proceeded, the fluorescence intensity of resorufin in solution increased, suggesting that the product can quickly diffuse out of the channels of NU-1000 (Movie S1). Control experiments using empty NU-1000-10 mm showed no  increase in fluorescence over the course of the experiment, indicating that the support has no catalytic activity of its own (Movie S2). To probe specifically the activity of enzymes sited in the MOF interior, we prepared a sample of NU-1000 containing active enzyme only in the interior of the crystal by treating NU-1000-10 mm with a 100 mM solution of cutinase for 2.5 hr followed by extensive washing. The sample was then placed in a 100 mM solution of catalytically inactive cutinase for 2 days to backfill the mesopores. In situ hydrolysis of resorufin butyrate was used to probe the enzyme accessibility and reactivity. We found that hydrolysis still occurred efficiently, indicating that reactant molecules can access cutinase not only on the MOF exterior but also in its crystalline interior (Figure 5 and Movie S3).</p><!><p>Before testing the activity and stability of immobilized cutinase, we compared the hydrolysis of p-nitrophenyl butyrate (PNPB) using cutinase@NU-1000, NU-1000, and the final solution used to soak cutinase@NU-1000 as catalysts (Figure 6A). The results indicate there was no activity for NU-1000 and no active cutinase detectable in the final solution. To gain further insight into the influence of immobilization on activity, the enzyme kinetic parameters k cat and K M for free cutinase and cutinase@NU-1000-5 mm were determined for the hydrolysis of three representative substrates (esters offering different aliphatic chain lengths: p-nitrophenyl acetate [PNPA], PNPB, p-nitrophenyl octanoate [PNPO]) (Table S1 and Figure S5). The results suggest that immobilized cutinase has similar overall activity to free cutinase in solution. We next compared the stability of cutinase@NU-1000-5 mm relative to that of soluble cutinase under several challenging conditions. As a measure of stability, we monitored the catalytic hydrolysis of PNPB, a common esterase substrate, by the free and encapsulated enzyme (Figures 6B-6E) and compared the activity in terms of turnover number (TON) as summarized in Table S2. The activity was monitored: (1) in detergent-containing buffer, (2) detergent-free buffer, as well as in buffers containing (3) the denaturant urea, or (4) the organic solvent tetrahydrofuran (THF). Esterases that also hydrolyze lipids, such as cutinase, are known to show protein-concentration-dependent aggregation in the absence of detergents or stabilizing additives upon reaction with hydrophobic substrates. 41 In the buffer containing detergent, cutinase@NU-1000-5 mm and free cutinase showed comparable catalytic performance (Figure 6B). In buffer without detergent, however, solution-phase cutinase gradually lost its catalytic activity, whereas cutinase@NU-1000-5 mm maintained turnover (Figure 6C). In 30 min, the TON of cutinase@NU-1000-5 mm was almost twice that of free cutinase. In buffer solution containing urea (560 mM), soluble cutinase was completely deactivated after 5 min; in contrast, the catalytic activity of cutinase@NU-1000-5 mm was unchanged after 60 min (Figure 6D). The TON for cutinase@NU-1000-5 mm was almost five times that of free cutinase in 60 min. Similarly, in a buffer solution containing 2.5% THF, free cutinase was rapidly deactivated (8 min), whereas cutinase@NU-1000-5 mm was little affected (Figure 6E). Under these conditions, the TON for cutinase@NU-1000-5 mm was still five times that of free cutinase in 60 min. Finally, we tested the stability of the complex to repeated reaction by following the hydrolysis of 300 mM PNPB catalyzed by 20 mg cutinase@NU-1000-5 mm over five reaction cycles. After each cycle, the complex was isolated by centrifugation, washed extensively with buffer, and subjected to another round of reaction. After five cycles, around 60% of the catalytic activity of cutinase@NU-1000-5 mm was retained (Figure 6F).</p><!><p>We compared the activity of cutinase immobilized in representative channeltype mesoporous MOFs to investigate the role of the unique hierarchical structure of NU-1000 for enzyme encapsulation applications. CYCU-3 42 and PCN-600 43  See also Figures S3 and S4.</p><p>were chosen as control MOFs to compare with NU-1000 (Figure 7A). All of these channel-type MOFs possess mesopores approximately 3.0 nm in diameter but differ in the identity of the metal nodes, organic linkers, and connectivity, resulting in different channel systems and accessibility within these frameworks. PCN-600 contains only hexagonal mesopores, which can, however, communicate via channel windows. CYCU-3 features a hierarchical micro-mesopore architecture similar to NU-1000; however, the walls of the channels in CYCU-3 are too condensed to allow reactant to diffuse freely between neighboring channels. In contrast, NU-1000 has open windows between the microporous and mesoporous channels, allowing for free communication between the different-sized pores. To observe the diffusion of cutinase into PCN-600 and CYCU-3, micro-sized crystals of each MOF were prepared for comparison with NU-1000-5 mm, and the loading of Cut647 in these MOFs was followed by CLSM over 5 days (Figure S6). The diffusion behavior of cutinase in all three MOFs was similar; cutinase slowly diffused from the two ends of each crystal into the center of the MOF. This demonstrates the generality of immobilization of enzymes into channel-type MOFs. To compare enzyme accessibility in CYCU-3, PCN-600, and NU-1000, nano-sized crystals of these three MOFs were synthesized 44 to ensure that the rate of diffusion was maximized and that the maximum loading of cutinase in each framework was achieved within 24 hr (Figure S6A). The final amounts of cutinase encapsulated by nano-sized NU-1000, CYCU-3, and PCN-600 were 12.9, 9.8, and 24.5 nmol/mg, respectively, as determined by ICP-OES. These findings are broadly consistent with the available mesopore volumes, i.e., $0.95 cm 3 /g for NU-1000 versus 0.90 cm 3 /g for CYCU-3 and 1.80 cm 3 /g for PCN-600. To ensure complete removal of any enzyme immobilized on the external surface of each nano-sized MOF prior to accessibility testing, the proteolytic agent trypsin was used to decompose cutinase on the outer surface (Figure 7B). The dimensions of trypsin are similar to those of cutinase, and thus cutinase sited within the MOF should not be decomposed by this process. The percentages of cutinase removed from the exteriors of NU-1000, CYCU-3, and PCN-600 were 3%, 39%, and 2%, respectively (Figure 7C). The comparatively large percentage of cutinase removed from CYCU-3 can be attributed to its comparatively poor stability in water, and therefore CYCU-3 was not studied further.</p><p>To accurately evaluate the accessibility of cutinase after immobilization in NU-1000 versus PCN-600, an organophosphorus ester with a fluorescent resorufin group was synthesized and used as an active-site titrant (Figures S7 and S8). 45 The titration results show that in PCN-600, only 1.5 nmol/mg (6% of the total amount) of cutinase encapsulated in the channels showed activity. Given that PCN-600 contains only hexagonal mesoporous channels, this result is not surprising because many of the mesopores are expected to be blocked by encapsulated cutinase, leaving little room for reactant diffusion. On the other hand, in NU-1000, 93% of the encapsulated cutinase was found to be accessible and catalytically active (Figure 7C). This demonstrates that, in addition to the importance of hierarchical pore structure for reactant diffusion, the presence of windows between the pores is key to achieving enzyme accessibly and hence enzyme turnover in a solid support.</p><!><p>By immobilizing an enzyme in the water-stable, hierarchical, channel-type MOF NU-1000, the enzyme can be stabilized in the array of larger channels, and reactants and products can diffuse through the array of smaller channels in the framework. It is this integration of large and small channels with the windows that connect them throughout the crystal that allows the MOF to be optimized for both binding and stabilization of the enzyme and diffusion of reactants and products. We have shown that NU-1000 is an excellent support for stabilizing the encapsulated enzyme in varying media such as THF and urea, whereas the free enzyme degrades rapidly under these conditions. Considering this stabilization of the enzyme, active-site accessibility, and open structure for diffusion, the design of channel-type mesoporous MOFs with hierarchical pore structure and pore windows should inspire further investigation in enzyme immobilization applications for biocatalysis where high catalytic efficiency and enhanced stability are required. Relevant to studies with other enzymes, and to optimization of enzyme activity, the degree of channel hydrophobicity in NU-1000 has been found to be highly tunable, as has the (bio)chemical composition of the channel lining, e.g., fluorocarbons, polypeptides, organic acids and bases, luminescent dyes, and even metal-sulfide clusters. 46,47 The tunability of MOFs, along with our increasing understanding of the design rules for stabilizing enzymes in an accessible fashion, hold great promise for the immobilization of an ever-expanding array of useful enzyme catalysts.</p><!><p>Zirconyl chloride octahydrate (ZrOCl 2 $8H 2 O), benzoic acid (C 6 H 5 COOH), aluminum chloride nonahydrate (AlCl 3 $9H 2 O), N,N-dimethylformamide, trifluoroacetic acid, 4,4 0 -stilbenedicarboxylic acid (H 2 SDC), PNPA, PNPB, PNPO, diethyl butylphosphonate, tris(hydroxymethyl)aminomethane (Tris), citric acid, fluorescein, resorufin butyrate, and resorufin sodium salt were purchased from Sigma-Aldrich and used as received. Fe(III) meso-tetra(4-carboxyphenyl)porphine chloride (FeTCPP) and meso-tetra(4-carboxyphenyl)porphine (TCPP) were purchased from Frontier Scientific and used without further purification. Iron(III) nitrate nonahydrate (Fe(NO 3 ) 3 $9H 2 O) and sodium acetate trihydrate (NaOOCCH 3 $3H 2 O) were purchased from Alfa Aesar. AlexaFluor 647 dye was purchased from Life Technologies (Thermo Fisher Scientific). The ligand 1,3,6,8-tetrakis(p-benzoic acid)pyrene (H 4 TBAPy) was synthesized according to the published procedure. 29 NU-1000, CYCU-3, precursor [Fe 3 O(OOCCH 3 ) 6 OH], and PCN-600 were synthesized according to published procedures. [42][43][44] F. solani pisi cutinase (molecular weight, 22.5 kDa) was made and purified according to a protocol described elsewhere. 32 Buffered aqueous solutions of cutinase (10 À5 -10 À7 M) were prepared at pH values of 6 (citric acid buffer) and 7.4 (Tris-HCl buffer).</p><!><p>N 2 sorption isotherm measurements were performed on a Micromeritics Tristar II 3020 (Micromeritics) at 77 K. Between 30 and 100 mg of material was used for each measurement.</p><!><p>Scanning electron microscopy images were taken using a Hitachi SU8030 or a Hitachi S4800-II at the EPIC facility (Atomic and Nanoscale Characterization Experimental Center, Northwestern University), which has received support from the MRSEC program (NSF DMR-1121262) at the Materials Research Center, the Nanoscale Science and Engineering Center (NSF EEC-0647560) at the International Institute for Nanotechnology, and the State of Illinois through the International Institute for Nanotechnology.</p><p>NMR Spectroscopy 1 H and 13 C nuclear magnetic resonance (NMR) spectra were recorded on a Bruker 500 FT-NMR spectrometer (500 MHz for 1 H and 126 MHz for 13 C), and 31 P NMR spectra were recorded on an Agilent 400 FT-NMR spectrometer (400 MHz) at the IMSERC (Integrated Molecular Structure Education and Research Center) of Northwestern University.</p><!><p>Data from electrospray ionization mass spectrometry were recorded on a Bruker AmaZon SL Ion Trap at the IMSERC.</p><!><p>The zeta potential of samples was measured using a Malvern Zetasizer Nano ZS. NU-1000 samples were made up in 18.2 MU deionized water at a concentration of 0.1 mg/ml and sonicated for 15 min. The pH of the solution was manually adjusted from 3 to 8 by the addition of 0.1 M HCl or NaOH to 10-15 ml of the suspension before the zeta potential was measured.</p><!><p>Quantification of zirconium (Zr) and sulfur (S) was accomplished using ICP-OES of acid-digested samples using individual Zr and S elemental standards prepared by diluting a 10,000 ppm certified Zr standard and 10,000 ppm certified S standard (Ricca Chemical Company) to 1.5625, 3.125, 6.25, 12.5, 25, and 50 ppm concentrations with 3% nitric acid (v/v), up to a total sample volume of 10 ml. ICP-OES was performed on a computer-controlled (QTEGRA software v.2.2) Thermo iCap 7600 Duo ICP-OES (Thermo Fisher Scientific) operating in standard mode and equipped with a SPRINT valve and CETAC 520 autosampler (Teladyne CETAC). Each sample was acquired using a 5 s sample loop fill (4 ml sample loop), a 4 s loop rinse, and a 4 s extra loop rinse (the rinse was 2% HNO 3 [v/v] and 2% HCl [v/v]). Samples were analyzed for Zr in radial view (339.198, 343.823, and 327.305 nm wavelengths) and S in axial view (180.731, 182.034, and 182.624 nm wavelengths) with three replicates and an exposure time of 20 s. Instrument performance was verified weekly via a performance report (passing manufacturer specifications). The enzyme loading was determined by comparing the experimental Zr/S ratio to the theoretical ratio given by the stoichiometry of Zr in the MOF to the number of cysteine thiols present in cutinase (Zr/S = 96:1).</p><!><p>Hydrolysis profiles of PNPA, PNPB, and PNPO by using cutinase or immobilized cutinase were recorded on a Beckman Coulter DU 640 spectrophotometer. Stock solutions of the esters were prepared using DMSO as a diluent. NU-1000-immobilized cutinase suspensions and free cutinase solution concentrations were adjusted to give similar initial rate profiles and time course length before treatment with various substrate concentrations. The reactions were conducted in a 1 ml cuvette containing 10 ml of the ester stock solution, 10 ml of enzyme solution/MOF-enzyme complex suspension, and 980 ml of TBS + 0.1% Triton X-100. Absorbance data at 410 nm (p-nitrophenyl absorption) were collected versus time. Michealis parameters for the enzyme substrate reactions were obtained using Lineweaver-Burke analysis of the initial rate data.</p><!><p>The stability of 5 nM cutinase and 5 nM immobilized cutinase in NU-1000 were compared using PNPB as a reactant in different media. For the recycling studies with NU-1000-immobilized cutinase, the reaction was performed with 300 mM PNPB using 20 mg of cutinase@NU-1000-5 mm in Tris buffer at pH 7 and room temperature. After the reaction, the mixture was centrifuged, and the supernatant was separated. The resulting solid was washed three times with Tris buffer to remove any soluble residue. The recovered NU-1000-immobilized cutinase was used for the next reaction by adding the same amount of reactant and buffer. The procedure was repeated five times as described above. The relative activity was calculated as a Chem 1, 154-169, July 7, 2016 165 ratio of the enzyme activity at any given cycle versus the enzyme activity during the first cycle.</p><!><p>AlexaFluor 647-labeled cutinase (Cut647) was prepared by reacting cutinase (100 mM) with 1.2 equivalents of an AlexaFluor 647-(ethyl-p-nitrophenyl)-phosphonate conjugate followed by purification of the labeled protein by size-exclusion chromatography. 48 This rendered the enzyme catalytically inactive. AlexaFluor 647 was chosen because of the relative insensitivity of its fluorescence intensity and quantum yield to environmental conditions and because of the excitation and emission maxima (650 and 665 nm, respectively) that occur far outside that of the pyrene struts (390 and 471/529 nm) used to construct the MOF. In the CLSM experiment using deactivated cutinase (Cut647), the complete loss of hydrolysis activity was confirmed by PNPB activity assay.</p><p>Immobilization of Cutinase in MOFs 2 mg of activated MOF (NU-1000, PCN-600, or CYCU-3) was added to 1 ml of deionized water and sonicated for 5 min until a uniform suspension was formed. The well-dispersed solid was isolated by centrifugation at 15,000 rpm for 1 min, and the supernatant was decanted. The solid was then suspended in a solution of cutinase (500 ml, 100 mM) in TBS (pH 7.4) for a given time (1 day for nano-sized MOF and 5 days for micro-sized MOF) at 25 C. The MOF-cutinase composite was then isolated by centrifugation at 15,000 rpm for 1 min, and the supernatant was removed. The solid was washed five times more with TBS containing 0.1% Triton X-100 and soaked in TBS buffer solution before further experiments.</p><!><p>NU-1000-10 mm crystals were used for all CLSM studies to monitor the uptake of cutinase and the distribution of enzymes throughout the matrix. 49 Fluorescence was examined by applying CLSM on a Leica TCS SP5. The Ar laser was set to 5%. Bit depth was set to 12 to achieve intensity resolution of 4,096 gray levels. Laser line 633 with 3% laser power was used to visualize AlexaFluor 647-dyelabeled cutinase on NU-1000. Quantitative analyses were performed using Leica LAS-AF image analysis, where a region of interest (ROI) was manually selected using the line tool. In a selected ROI, measurements of the relative mean intensity of the fluorescence signals were taken by the Leica LAS-AF image analysis program. The Mark and Find panel was used to locate the position (x, y, and z coordinates) of the same single crystal of NU-1000 during the test at different time points. The loading process of Cut647 into a single crystal of NU-1000 was monitored using in situ CLSM. The submerged NU-1000 crystals were first placed into a solution with Tris buffer. 2D (xy) concentration profiles of Cut647 taken at a fixed z depth (corresponding to the center layer) within the crystal of NU-1000 were acquired on a Leica-SP5 CLSM once Cut647 was added to the solution. During imaging, the laser power was set as low as possible (5%, 1 mW HeNe laser) to avoid fluorescence saturation and to minimize photo bleaching. To obtain time-dependent Cut647 concentration profiles for the MOF and avoid decay of the fluorophore caused by repeated illumination by the laser, images were acquired only at given time intervals: 2, 60, 240, 960, 1,440, 2,880, 4,320, and 7,200 min. Using the line tool in ImageJ, 50 fluorescence intensity profiles along the middle of an NU-1000 crystal were obtained for each sample and plotted as a function of the length of the crystal versus time.</p><!><p>The titration experiments were performed using the Molecular Devices Gemini EM fluorescence/chemiluminescence plate reader. A calibration curve relating cutinase activity to the intensity of liberated resorufin was generated by treating cutinase at several concentrations (8 mM, 6 mM, 4 mM, 2 mM, 1 mM, and 500 nM) in 4-morpholineethanesulfonic acid (MES)-saline buffer (20 mM MES, 150 mM NaCl [pH 6.0]) with a solution of 5 mM resorufin phosphonate (RP) in MES-saline at 37 C. Steady-state fluorescence intensities at the completion of the reaction were plotted versus enzyme concentration and fitted to a straight line to generate the calibration curve. For the titration experiments, 5 mg of nano-sized empty NU-1000, empty PCN-600, cutinase@NU-1000, and cutinase@PCN-600 were treated with RP (5 mM) at 37 C, and the fluorescence intensity was monitored for 5 hr. Background-subtracted steady-state fluorescence values for each of the experimental samples were then compared with the calibration curve to yield the concentration of cutinase active sites present in the MOF-enzyme samples.</p><!><p>Supplemental Information includes Supplemental Experimental Procedures, eight figures, two tables, and three movies and can be found with this article online at http://dx.doi.org/10.1016/j.chempr.2016.05.001.</p><!><p>O.K.F., J.T.H., and M.M. initiated the project; P.L. (with some help from A.J.H.) synthesized the MOF and collected and analyzed the data from gas adsorption, X-ray diffraction, scanning electron microscopy, CLSM, ICP-OES, and NMR under the supervision of O.K.F. and J.T.H.; J.A.M. prepared the enzymes, labeled the enzymes with dyes, and performed the titration experiments under the supervision of M.M.; E.V. and P.Z.M. performed the computations and analyzed the results under the supervision of R.Q.S.; P.L., J.A.M., and A.J.H. wrote the paper, and all authors contributed to revising the paper.</p>