Molecular Fundamentals of Enzyme Nanogels - American Chemical

J. Phys. Chem. B 2008, 112, 14319–14324

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Molecular Fundamentals of Enzyme Nanogels Jun Ge,† Diannan Lu,† Jun Wang,† Ming Yan,†,‡ Yunfeng Lu,*,‡ and Zheng Liu*,† Department of Chemical Engineering, Tsinghua UniVersity, Beijing 100084, China, and Department of Chemical and Biomolecular Engieering, UniVersity of California, Los Angeles, California ReceiVed: June 19, 2008; ReVised Manuscript ReceiVed: August 13, 2008

The assembly of a monomer around an enzyme as the essential step in the fabrication of enzyme nanogel by in situ polymerization was illustrated by molecular dynamics simulation and evidenced by a fluorescence resonance energy transfer spectrum, using lipase/acrylamide as a model system. The subsequent polymerization generated a hydrophilic gel network which not only strengthened the protein structural integrity via multipoint linkage but also increased the number of intramolecular H-bonds of the encapsulated protein, as suggested by the blue shift of the fluorescence spectrum of the encapsulated lipase. This greatly enhanced the stability of lipase at high temperature, as experimentally demonstrated. The exclusion of polar solvent molecules from the encapsulated enzyme, in contrast to the enrichment of water molecules, due to the presence of a hydrophilic gel network was displayed. This established a hydrophilic microenvironment for the encapsulated protein and thus gave the encapsulated protein an enhanced tolerance to the organic solvent, as experimentally observed in the present study and reported elsewhere. These results have given a molecular insight into the enzyme nanogel as well as its high potential as a robust enzyme model for an expended application spectrum of enzymatic catalysis. 1. Introduction Nanostructured enzyme hybrids such as enzyme nanoparticles,1 enzyme-carbon nanotube composites,2,3 self-assembled enzyme nanowires,4-8 as well as enzyme or enzyme mimics micelles9-12 incorporate the unlimited structural derivability of chemical molecules and thus pave the way to fully display the expected functionalities of biomolecules in area such as nanoand microdevices,13,14 artificial cells,15,16 nanoscale reactors,17,18 and a “nano-scaled factory”.19 Recent years have witnessed the growing efforts in exploring the ways to fabricate enzyme-polymer nanoparticles, which can be classified into two categories, namely, the “grafting onto” and the “growing from” method. The first one involves the immobilization of the enzyme on polymer nanoparticles,20 while the second entails the polymerization from a protein surface. In this case, small functional groups, such as initiators or vinyl groups, are grafted onto the enzyme surface, followed by in situ polymerization that “grows” polymer from the enzyme surface.21-25 In their work of fabricating enzyme nanoparticles, Kim and Grate1 have obtained enhanced enzyme stability at an insignificant increase in the mass-transfer resistance. Recently, Liu and his co-workers26,27 have proposed an aqueous two-step in situ polymerization method for enzyme nanogels, in which the first step is to generate vinyl groups on the protein surface by acryloylation28,29 and the second step is the aqueous in situ polymerization that encapsulates the acryloylated protein. Encapsulation into the porous and flexible polymer network grants the enzyme an almost identical bioactivity to its native counterpart but significantly enhanced stabilities against high temperatures and polar solvents. The depth of the gel layer can be conveniently controlled at around * To whom correspondence should be addressed. Tel: +86-10-62779876. Fax: +86-10-6277-0304. E-mail: [email protected] (Z.L.); E-mail: [email protected]. (Y.L.). † Tsinghua University. ‡ University of California.

10-50 nm by monomer loading, while different monomers can be included to provide various structures and properties in response to the following applications, which are essential for the application of enzyme nanogel as a biocatalytic building block. The present work is in the context of the above efforts but focuses on the molecular interactions involved in the fabrication, which underlies both the design and the synthesis of the enzyme nanogel. Here, Candida rugosa lipase (CRL),30 an enzyme of growing interest in organic synthesis, was chosen as the enzyme model, and acrylamide was chosen as the monomer for the fabrication of the enzyme nanogel with enhanced biocatalytic performance. Molecular dynamics simulation, structural analysis via a fluorescence resonance energy-transfer spectrum, transmission electron microscopy, and activity assay at various conditions were jointly applied to provide a molecular insight into the synthesis and structure of the enzyme nanogel. 2. Experimental Section 2.1. Materials. Lipase from Candida rugosa (Type VII), p-nitrophenyl palmitate (p-NPP), acrylamide (AM), N-acryloxysuccinimide (NAS), N,N,N′,N′-tetramethylethylenediamine (TEMED), ammonium persulfate (APS), pyrene, dimethyl sulfoxide (DMSO), and methanol were purchased from Sigma-Aldrich, U.S.A. Other chemicals were of analytical grade and used without further purification. 2.2. Purification of Lipase. Candida rugosa lipase (Type VII, Sigma) was purified as described previously.31 Ten mg of crude lipase powder was added to 100 mL of 25 mM pH 7.5 phosphate buffer, followed by 2 h of stirring at 4 °C and 15 min of centrifugation at 16000g. (NH4)2SO4 was added to the suspension up to 30% saturation. The supernatant was collected after centrifugation, followed by the addition of (NH4)2SO4 up to 65% saturation. After centrifugation, the precipitate was collected, dissolved in 25 mM pH 7.5 phosphate buffer, and dialyzed against the same buffer. The lipase solution was then

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14320 J. Phys. Chem. B, Vol. 112, No. 45, 2008 loaded on a Hitrap DEAE-Sepharose column pre-equilibrated with 25 mM pH 7.5 phosphate buffer. After washing the unbound components, lipase was eluted with a linear gradient salt concentration (0-1 M NaCl). The elution rate was 1 mL/ min, and each of the fractions (1 mL) was analyzed for lipase activity. The peak with the major lipase activity was collected and applied to a Hitrap desalting column, followed by lyophilization. 2.3. Fluorescence Resonance Energy Transfer (FRET). The emission spectrum was recorded from 300 to 550 nm with an excitation wavelength of 285 nm using RF-5301 PC, SHIMADZU. The emission spectrum of lipase in the absence of pyrene was recorded with a protein concentration of 0.08 mg/mL, while in the presence of pyrene, it was recorded by adding 10 µL of pyrene/DMSO solution (0.1 mg/mL) to 2 mL of the above solution. The energy-transfer efficiency (ET) was calculated via ET ) 1 - FDA/FD (the relative fluorescence intensity of lipase in the presence (FDA) and absence (FD) of pyrene). Then, by varying the concentrations of acrylamide (0, 0.5, 1, 1.5, 2% (w/v)) in the lipase solution, the value of ET was determined by the same experimental procedure. 2.4. Molecular Dynamics Simulation. The structure of native Candida rugosa lipase was obtained from the Brookhaven Protein Data Bank (PDB code: 1TRH). To be consistent with the experimental conditions, none of the aspartates was protonated. The GROMACS 3.3 package was used to perform molecular dynamics (MD) simulation. This package is a collection of programs and libraries for MD simulation and the subsequent analysis of trajectory data. Simulations were performed using general triclinic cell geometry. Pressure and temperature coupling was implemented for all types of simulation cells. The Berendsen’s weak coupling algorithm scheme was used for both pressure and temperature. During the simulation, lipase was put into the center of a rectangular box using periodic boundary conditions with either water or a water/acrylamide solution. The size of the simulation box was 16.48 × 16.445 × 14.773 nm. The system containing a lipase molecule and a certain number of water molecules or a water/acrylamide mixture was submitted to 500 steps of steepest descent minimization converging to a value of 2000 kJ · mol-1 · nm-1, applying the Particle-Mesh Ewald method at 303 K (25 °C). Then, a 10 ps position-restrained MD simulation was performed by keeping the protein coordinate fixed and allowing the water and acrylamide molecules to equilibrate themselves. Then, a 1000 ps MD was performed at 303 K, and a leapfrog algorithm was used for integrating the Newtonian equations of motion for 500000 simulation steps, with a time step of 0.002 ps. In the case of simulating the effects of methanol on lipase naogel, the coordinate of the protein-acrylamide system was fixed, while water and methanol molecules were mobile. 2.5. Synthesis of Lipase Nanogel. To determine the optimal pH, lipase was dissolved, respectively, in acetic buffer (50 mM, pH 4.0), phosphate buffer (50 mM, pH 6.0), borate buffer (50 mM, pH 8.0), and borate buffer (50 mM, pH 9.4), followed by a 24 h dialysis against the same buffer. The protein concentration in solution was about 4 mg/mL. Then, 12 mg of N-acryloxysuccinimide (NAS) was dissolved in 600 µL of DMSO and slowly added to 5 mL of the enzyme solution, with the molar ratio of NAS to lipase being 200:1. After reaction at 30 °C for 6 h, the solution was collected and dialyzed against phosphate buffer (50 mM, pH 7.0) at 4 °C for 24 h to remove unreacted

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Figure 1. Molecular simulation of the lipase-acrylamide assembly; (1) lipase; (2) water; (3) acrylamide.

reagents. The hydrolytic activity of modified lipase was determined using p-nitrophenyl palmitate (p-NPP) as the substrate. Quantities of 250 mg of acrylamide (AM), 15 mg of ammonium persulfate (APS), and 12 µL of N,N,N′,N′-tetramethylethylenediamine (TEMED) were directly added to 5 mL of acryloylated lipase solution (4.0 mg/mL, 50 mM pH 7.0 phosphate buffer), and polymerization proceeded at 30 °C for 12 h. The product was dialyzed against phosphate buffer (50 mM, pH 7.0) at 4 °C for 24 h to remove unreacted reagents, followed by lyophilization. 2.6. Transmission Electron Microscopy (TEM). TEM of the sample was determined using a Hitachi JEOL200CX highresolution transmission electron microscope. Carbon-coated grids were prepared by adding a drop of protein solution, removing the excess, and applying 1% pH 7.0 sodium phosphotungstate. The sample was then subjected to TEM measurement. 2.7. Assays. The lipase concentration was determined by bicinchoninic acid colorimetric protein assay (BCA) calibrated with bovine serum albumin (BSA).32 Primary amine groups were determined by a fluorometric assay.33 The hydrolytic activity of lipase was determined using p-nitrophenyl palmitate (p-NPP) as the substrate. During a run, p-NPP was first dissolved in acetone and then diluted with phosphate buffer (50 mM, pH 7.0) containing 1.25% (w/v) Triton X-100,34 giving the final concentration of 0.5 mM. The reaction was started by adding 50 µL of enzyme solution to 950 µL of substrate solution and detected at 348 nm. Release of 1 µmol of p-nitrophenol per minute in the assay conditions was defined as one enzyme unit activity (U · mg-1). 3. Results and Discussion 3.1. Molecular Simulations of the Lipase-Acrylamide Assembly. Using protocols mentioned in the experimental methods section, the lipase-acrylamide assembly at a concentration of acrylamide of 50 mg/mL was obtained, as exemplified by Figure 1, in which lipase was encapsulated by the acrylamide aggregates that could form a gel network by a subsequent polymerization. An overview of the assembly of monomers around lipase can be interpreted from the radial distribution function (rdf) of acrylamide molecules to lipase. As shown in Figure 2a, two rdf peaks are presented for acrylamide, in which the first one at 0.370 nm is obtained by a H-bond formed between amino acid

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Figure 2. (a) The rdf of water and acrylamide around the lipase (H2O: rdf of water; H2O/AM: rdf of water in the presence of AM; AM: rdf of AM). (b) Number of H-bonds between lipase and acrylamide/water; (1) H-bond between lipase and water in pure water solution, (2) H-bond between lipase and water in acrylamide solution, (3) H-bond between lipase and acrylamide in acrylamide solution, (4) H-bond between lipase with water and acrylamide in acrylamide solution.

Figure 3. Number of intramolecular H-bonds of lipase in water and in AM solution; (1) intramolecular H-bonds of lipase in water; (2) intramolecular H-bonds of lipase in AM solution.

residues of lipase and the amide groups of acrylamide, while the second one at 4-5 nm is obtained by the intermolecular forces among acrylamide. Recall the diameter of CRL of 7 nm (PDB code: 1TRH), the minimum diameter of the lipase nanogel is expected to be around 15-17 nm after the polymerization. The assembly of acrylamide around lipase can also be interpreted from the H-bond formed between lipase with water in the presence and absence of acrylamide. As shown in Figure 2b, the presence of acrylamide reduces the total number of H-bonds formed between lipase and water, indicating the replacement of water at the enzyme surface by acrylamide. In this case, the total number of H-bonds of the system was also reduced due to the spatial hindrance of acrylamide located at the lipase surface. 3.3. Enhanced Stability of Encapsulated Lipase at High Concentration: Molecular Simulation. Dissociation of the hydrophobic core is one essential reason leading to the deactivation of enzyme at high temperature. For the above-mentioned system with an acrylamide concentration of 50 mg/mL, the intramolecular H-bond between amino acids of lipase was simulated as a function of the molecular assembly. As shown in Figure 3, the formation of the lipase-acrylamide assembly gives a significantly increased number of intramolecular H-bonds within lipase. This strengthened intramolecular interaction allows the encapsulated lipase to withstand a higher temperature, particularly after polymerization that gives multipoint linkage with the porous acrylamide network. An enhanced tolerance to organic solvent will expand to the application of the spectrum of enzymatic catalysis, in which the stripping of essential water from the enzyme surface essentially accounts for the deactivation of the enzyme.35,36 This mechanism can be adequately addressed by the distribution of water and organic solvent. Here, a fixed lipase-acrylamide assembly was used to represent the lipase nanogel, while

methanol was chosen as the model organic solvent. Due to the fact that the water content in the lipase nanogel after lyophilization was 10% (w/w), the solution composition was set as 90% (w/w) methanol and 10% (w/w) water. The simulation was then performed at different contents of acrylamide and plotted in Figure 4. Here, ∆ is applied to account for the changes of the distribution of methanol and water around the “lipase nanogel”, using the rdf obtained when the “lipase nanogel” was put into methanol solution at t ) 0 as the reference. Figure 4a shows the ∆rdf’s of methanol around the acrylamide layer with depths of 1, 3, and 5 nm. Here, positive peaks of ∆rdf are obtained at 1.5, 3.8, and 6.1 nm from the mass center of lipase, indicating that methanol molecules are expelled from the lipase encapsulated into the acrylamide assembly. The increase in the peak value in response to gel depth indicates a simple and straightforward way to enhance the exclusion of methanol. In contrast, as shown in Figure 4b, the negative ∆rdf of water is achieved, which means the loss of water molecules around lipase when immerged into methanol. Here, more water molecules are preserved within the “lipase nanogel” with the increased depth of gel. It should be noted here that (1) this enhanced hydrophilic microenvironment is established in the presence of methanol and (2) the loss of water is declined when the gel layer becomes thick. Thus, the presence of “polyacrylamide gel” leads to contrast changes of rdf for methanol and water, which essentially enables the inhibition of the stripping of water from the lipase surface. An enhanced tolerance to methanol as a representing polar solvent can be obtained for the encapsulated lipase. 3.4. Detection of the Lipase-Acrylamide Assembly. Detection of the lipase-acrylamide assembly in solution was performed by fluorescence resonance energy transfer (FRET),37 in which lipase served as a donor and pyrene as an acceptor. The emission spectrum of lipase in the absence of pyrene was recorded with a lipase concentration of 0.08 mg/mL, while in the presence of pyrene, it was recorded by adding 10 µL of pyrene/DMSO solution (0.1 mg/mL) to 2 mL of the above solution. Figure 5a gives the emission spectrum of lipase and the absorption spectrum of pyrene. Once pyrene was introduced into the lipase-acrylamide solution, FRET occurred from lipase to pyrene, resulting in an increase in the fluorescence intensity of pyrene at 370-500 nm and a simultaneous reduction for lipase at 340 nm (Figure 5b). The energy-transfer efficiency (ET) is determined as38 ET ) 1 - FDA/FD, representing the relative fluorescence intensities of lipase in the presence, FDA, and absence, FD, of pyrene. As shown by Figure 6, ET of the lipase-pyrene pair increased from 0.08 to 0.18 once acrylamide was loaded at concentrations up to 2% (w/v). This indicates

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Figure 4. The changes of distribution of a) methanol and b) water around lipase in the presence of polyacrylamide gel network.

Figure 5. (a) The emission spectrum of lipase and the absorption spectrum of pyrene. (b) FRET from lipase to pyrene (1) in absence of AM and (2) in presence of 0.5% AM in solution.

Figure 6. The energy-transfer efficiency from lipase to pyrene and the intensity ratio of peaks 1 to 3 in the emission spectrum of pyrene at various concentration of AM.

Figure 7. Relative activity and modification degree of lipase nanogel at different pH’s.

that pyrene molecules are pushed to the lipase surface by the acrylamide assembly. In addition, the intensity ratio of peaks 1 to 3 (denoted by I1/I3, I1: 374 nm, I3: 382 nm) in the emission spectrum of pyrene, which serves as an index of the polarity of the microenvironment around pyrene,39 decreased from 1.24 to 1.16 once acrylamide was loaded to the solution at 2% (w/v). This suggests that pyrene molecules are incorporated into a more hydrophobic microenvironment in the presence of acrylamide. In other words, the original water molecules were expelled by the acrylamide molecules that assembled around the lipase. 3.5. Preparation and Characterization of Lipase Nanogel. For the step of lipase modification by acryloylation, it is shown by Figure 7 that while a low pH favors an increased residual activity, the modification degree also decreases. The optimal pH for the acryloylation of lipase with NAS was determined as pH 4.0 with the molar ratio of NAS to lipase of 200:1, at which the residual activity of the modified lipase remained 92% and the modification degree reached 52% after reaction at 30 °C for 5 h. An elevated pH, which increases the nucleophilicity of the amino group that favors the chemical reaction with NAS, leads to an unexpected low residual activity of lipase. This may be due to either the poor stability of lipase at alkaline pH31 or the modification of the residuals essential for maintaining the

active site of lipase. Thus, if not stated otherwise, the surface modification was carried out at pH 4.0, followed by in situ polymerization. Transmission electron microscopy (TEM) was applied to determine the size of the lipase nanogel synthesized using the above-mentioned procedure. As shown by Figure 8a, lipase nanogels have diameters ranging from 10 to 30 nm, recalling that the predicted minimum diameter of the lipase nanogel is 15-17 nm. Figure 8b gives the size exclusion chromatography with florescence detection of native lipase, lipase nanogel, as well as the product obtained from the control synthesis using native lipase instead of acryloylated lipase. The identical elution volume suggests that no enzyme nanogel was obtained from the control synthesis, indicating that the generation of vinyl groups on the enzyme surface is requested for the preparation of the enzyme nanogel using acrylamide for the in situ polymerization. Moreover, as shown by Figure 9, the maximum emission wavelengths of the native lipase, acryloylated lipase, and the lipase nanogel are determined to be 343, 336, and 333 nm, respectively, that is, a blue shift in the maximum emission wavelength of the lipase after acryloylation and encapsulation. The blue shift for lipase nanogel may due to an increased intramolecular H-bond within lipase after encapsulation, as predicted by Figure 3. The residual activity

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Figure 8. (a) TEM of lipase nanogel. (b) SEC of native lipase, lipase nanogel, and the products obtained from the control synthesis using native lipase without acryloylation.

Figure 9. Fluorescence spectra of the native lipase, acryloylated lipase, and lipase nanogel.

Figure 10. Stability of native lipase, acryloylated lipase, and lipase nanogel incubated in 50 mM pH 7.0 phosphate buffter at 50 °C.

of lipase nanogel determined using p-NPP as the substrate remained 85%, indicating that the lipase nanogel may essentially resemble its native counterpart. 3.6. Examination of the Stability of the Lipase Nanogel. Native lipase, acryloylated lipase, and lipase nanogel with the same concentration of 0.1 mg/mL in 50 mM pH 7.0 phosphate buffter were incubated at 50 °C, and samples were taken out at specific times to measure their residual activities by hydrolysis of p-NPP, as mentioned in the Experimental Section. Here, the half-life, the time length at which the enzyme remains as 50% of its initial activity, is applied as the index of enzyme stability. As shown in Figure 10, the half-life of native lipase was 30 min, and acryloylated lipase lost 30% of its initial activity by 500 min, while the lipase nanogel remained almost unchanged over 500 min. The enhanced thermal stability for lipase nanogel, we believe, can be attributed to the multipoint linkage with the gel network and the intensified intramolecular H-bond, as shown in Figure 3. To check the tolerance to methanol, native lipase, acryloylated lipase, and lipase nanogel in powder form were incubated in anhydrous methanol with the protein concentration of about 0.5 mg/mL at 50 °C. Samples were taken out from the solution to

Figure 11. Stability of native lipase, acryloylated lipase, and lipase nanogel incubated in methanol at 50 °C.

determine the residual hydrolysis activity using p-NPP as the substrate. As shown in Figure 11, the native and the acrylated lipase is almost completely deactivated in 30 min, while lipase nanogel preserves most of its activity over 180 min. The enhanced tolerance to polar solvent, as predicted from the molecular simulation shown in Figure 4, is due to the presence of the hydrophilic polyacrylamide network that rejects methanol while enriching the water content at the lipase surface. This enhanced stability of lipase in the presence of polar solvent, as well as the enhanced thermal stability, paves the way to both the extension and intensification of the industrial biocatalysis, for example, production of biodiesel by lipase catalysis in the presence of methanol. 4. Conclusions In conclusion, we have shown by molecular dynamics simulation and FRET the assembly of monomers around an enzyme in aqueous solution via hydrogen bonding, which paves the way to the subsequent polymerization yielding enzyme nanogels. The formation of the hydrophilic polyacrylamide gel network leads to an intensified intramolecular H-bond within lipase and, more importantly, a hydrophilic environment that expels organic solvent while attracting water molecules. All of these, together with the multipoint linkage, result in significantly enhanced thermal stability and tolerance to polar solvent, as experimentally demonstrated, and make the enzyme nanogels robust catalytic building blocks for artificial biological processes and devices at different scales. A molecular insight of the enzyme nanogel established via joint applications of molecular simulation and experimental approaches, as attempted by the present study, facilities the design, synthesis, and application of the enzyme nanogel. In addition to the H-bond, a static electronic force or hydrophobic interactions may also be anticipated to create a monomer-enzyme assembly and fabricate various enzyme nanogels and thus merit further efforts.

14324 J. Phys. Chem. B, Vol. 112, No. 45, 2008 Acknowledgment. This work was supported by the National Natural Science Foundation of China under Grant Number 20776076, the key Project of the National Natural Science Foundation of China under Grant No. 20636040, the Ministry of Science and Technology through 973 Project under Grant No. 2003CB716004, and the Project of the Ministry of Education under Grant Number 20070003091. References and Notes (1) Kim, J.; Grate, J. W. Nano Lett. 2003, 3, 1219–1222. (2) Besteman, K.; Lee, J.; Wiertz, F. G. M.; Heering, H. A.; Dekker, C. Nano Lett. 2003, 3, 727–730. (3) Rege, K.; Raravikar, N. R.; Kim, D.; Schadler, L. S.; Ajayan, P. M.; Dordick, J. S. Nano Lett. 2003, 3, 829–832. (4) Velonia, K.; Rowan, A. E.; Nolte, R. J. M. J. Am. Chem. Soc. 2002, 124, 4224–4225. (5) Jia, H.; Zhu, G.; Vugrinovich, B.; Kataphinan, W.; Reneker, D. H.; Wang, P. Biotechnol. Prog. 2002, 18, 1027–1032. (6) Wang, Q. G.; Yang, Z. M.; Wang, L.; Ma, M. L.; Xu, B. Chem. Commun. 2007, 10, 1032–1034. (7) Wang, Q. G.; Yang, Z. M.; Zhang, X. Q.; Xiao, X. D.; Chang, C. K.; Xu, B. Angew. Chem., Int. Ed. 2007, 46, 4285–4289. (8) Wang, Q. G.; Yang, Z. M.; Ma, M. L.; Chang, C. K.; Xu, B. Chem.sEur. J. 2008, 14, 5073–5078. (9) Boerakker, M. J.; Hannink, J. M.; Bomans, P. H. H.; Frederik, P. M.; Nolte, R. J. M.; Meijer, E. M.; Sommerdijk, N. A. J. M. Angew. Chem., Int. Ed. 2002, 41, 4239–4241. (10) Reynhout, I. C.; Cornelissen, J. J. L. M.; Nolte, R. J. M. J. Am. Chem. Soc. 2007, 129, 2327–2332. (11) Fife, W. K.; Rubinsztajn, S.; Zeldin, M. J. Am. Chem. Soc. 1991, 113, 8535–8537. (12) Bhattacharya, S.; Snehalatha, K. Langmuir 1995, 11, 4653–4660. (13) Xiao, Y.; Patolsky, F.; Katz, E.; Hainfeld, J. F.; Willner, I. Science 2003, 299, 1877–1881. (14) Ebara, M.; Hoffman, J. M.; Hoffman, A. S.; Stayton, P. S. Lab Chip 2006, 6, 843–848. (15) Vriezema, D. M.; Hoogboom, J.; Velonia, K.; Takazawa, K.; Christianen, P. C. M.; Maan, J. C.; Rowan, A. E.; Nolte, R. J. M. Angew. Chem., Int. Ed. 2003, 42, 772–776. (16) Vriezema, D. M.; Garcia, P. M. L.; Oltra, N. S.; Hatzakis, N. S.; Kuiper, S. M.; Nolte, R. J. M.; Rowan, A. E.; van Hest, J. C. M. Angew. Chem., Int. Ed. 2007, 46, 7378–7382.

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