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Enhancing Enzyme Activity and Immobilization in Nanostructured Inorganic-Enzyme Complexes Xuye Lang, Lingling Zhu, Yingning Gao, and Ian Wheeldon Langmuir, Just Accepted Manuscript • DOI: 10.1021/acs.langmuir.7b02004 • Publication Date (Web): 16 Aug 2017 Downloaded from http://pubs.acs.org on August 16, 2017

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Enhancing Enzyme Activity and Immobilization in Nanostructured Inorganic-Enzyme Complexes

Xuye Lang1, Lingling Zhu2, Yingning Gao1,#, and Ian Wheeldon1,*

1.

Department of Chemical and Environmental Engineering, University of California,

Riverside USA 92521 2.

School of Materials Science and Engineering, Zhengzhou University, Zhengzhou China

450001

*To whom correspondence should be addressed: Phone: (951) 827-2471 Fax: (951) 827-5696 Email: [email protected]

Current address #

Yingning Gao, Nanomedical Diagnostics Inc., 6185 Cornerstone Court East Suite #110, San

Diego, CA 92121, U.S.A.4

ACS Paragon Plus Environment

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ABSTRACT Understanding the chemical and physical interactions at the interface of protein surfaces and inorganic crystals has important implications in the advancement of immobilized enzyme catalysis. Recently, enzyme-inorganic hybrid complexes have been demonstrated as effective materials for enzyme-immobilization. The precipitation of phosphate nanocrystals in the presence of enzymes creates enzyme-Cu3(PO4)2-3H2O particles with high surface-to-volume ratios, enhanced activity, and increased stability. Here, we begin to develop a mechanistic understanding of enzyme loading in such complexes. Using a series of enzymes including horseradish peroxidase (HRP), a thermostable alcohol dehydrogenase (AdhD), diaphorase, catalase, glucose oxidase (GOx), and the protein bovine serum albumin (BSA), we identified a correlation between particle synthesis temperature, overall enzyme charge, and enzyme loading. The model enzyme HRP has a high predicted pI of ~7.5 and maintains an overall positive charge under the synthesis conditions, phosphate buffer pH 7.4. HRP loading in HRP-Cu3(PO4)2 complexes were enhanced by 4.2-fold when synthesis was carried out at 37 °C in comparison with synthesis at 4 °C. HRP loading was further enhanced with synthesis at pH 8.0, correlating with a decrease in overall enzyme charge. Proteins with lower predicted pI values and negative overall charge (AdhD, pI of 5.6; diaphorase, pI of 6.8; GOx, pI of 5.2; catalase, pI of 6.9; and, BSA, pI of 5.7) exhibited higher enzyme loadings with 4 °C synthesis, 2.7-, 2.6-, 2.5-, 1.8-, and 1.7-fold protein loading enhancements, respectively. Using HRP as a model system, we also demonstrate that activity increased concomitantly with enzyme loading, and that particle nanostructure was minimally affected by synthesis temperature. Combined, the results presented here demonstrate the control of enzyme loading in enzyme-inorganic particles opening up new possibilities in enzyme and multienzyme catalysis.


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INTRODUCTION Designing multistep reaction cascades with controllable product distributions, high molecular efficiencies, and high product selectivity is a grand challenge in chemistry.1 Recent research in this area has focused on creating multienzyme systems with tightly controlled spatial organization and tunable ratios of cascade enzymes. For example, two and three-step enzymatic systems have been created using liposomes that immobilize one or more reaction steps on the inside of the structure and localize other reaction steps to the outer surface;2-3 DNA molecular scaffolds have been used to control the nanometer-scale spatial organization of multiple enzymes and their cofactors;4-6 and, inorganic-enzyme particles, the focus of this study, have been used to create enzyme complexes with core-shell structures.7 Metal organic frameworks (MOFs) and other nanoscale materials have also been used to create multicatalytic structures.8-10 Collectively, these systems demonstrate that the architecture of a multienzyme complex can enhance cascade catalysis: nanoscale colocalization of enzymes has been shown to increase presteady state reaction rates and spatial organization can prevent cascade intermediates from diffusing to the bulk solution, thus increasing reaction yields and cascade selectivity.1, 11-13 Kinetic enhancements can also arise from molecular interactions between an enzyme’s substrates and the material used to create a complex.14-17 A common design challenge in enzyme and multienzyme systems is immobilization, that is, enzyme loading in the fabricated particle or multienzyme complex. Low enzyme loadings create systems with poor catalytic capacity, whereas high enzyme loadings can lead to high reaction rates due to an increased number of active sites, help minimize activity losses due to immobilization, and mitigate potential activity losses over time. Low or variable enzyme loadings can also complicate kinetic analysis. An example of this design challenge is the


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immobilization of enzymes on DNA nanostructures. Using a DNA nanoscale tile with enzyme immobilization points at various distances, Fu et al. were able to study the effect of interenzyme distance on cascade catalysis; however, the kinetic analysis was complicated by the fact that immobilization efficiency varied between structures, ranging from ~45% to upwards of ~95%.5 Assembly yields of enzymes in similar DNA nanostructures were subsequently improved, enabling investigations into the effects of DNA scaffolds on enzyme and cascade catalysis.18 Enzyme immobilization in inorganic-protein nano- and microscale structures is useful because enzyme activity can be enhanced, the spatial organization of two and three-step enzyme cascade can be controlled, and the structures have proved to be reusable with limited kinetic losses over time.7, 19-22 This new materials strategy exploits the formation of inorganic crystals on protein surfaces to create enzyme aggregates with demonstrated applications in cascade and tandem catalysis,7, 23 biosensing,21, 24 and proteolytic digestion.25-26 A technical challenge with these materials is that the synthesis procedures result low enzyme loadings. Immobilization of HRP in Cu3(PO4)2 complexes range from 2-17 wt% HRP.12, 21 Synthesis with other enzymes produced similar results, for example glucose oxidase (GOx) was loaded at ~10 wt%,7 trypsin at ~12.5 wt%,21 chymotrypsin at ~10 wt%,26 and amylase at ~20 wt%.27 A reasonably complete understanding of how such enzyme complexes are created has been developed,7, 20, 22-23, 28 but the issue of low enzyme loading has not yet been directly addressed. In this work, we seek to increase enzyme loading in inorganic-enzyme complexes fabricated with protein-Cu3(PO4)2 hybrid materials. Such complexes have utility in cascade catalysis because particle structure and the spatial organization of enzymes within the particle can be controlled, and fabrication is straightforward and facile. These hybrid materials are also generally useful as an enzyme immobilization strategy. By studying the effects of synthesis


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temperature on enzyme loading, activity, and particle structure, we increased the loading of HRP in HRP-Cu3(PO4)2 microscale particles. Translation of our findings to different enzymes with varied chemical properties (specifically, a range of isoelectric points), we begin to develop an understanding the chemical mechanisms responsible for enhanced loading and consequently enzyme particle activity. While this work focuses on increasing the immobilization of a single enzyme within a protein-inorganic particle, the principle is general and can be applied to complexes with two or more enzymes that control the spatial localization of enzyme within the complex.

MATERIALS AND METHODS Chemicals. Horseradish peroxidase (HRP, E.C. 1.11.1.7), glucose oxidase from Aspergillus niger (GOx, E.C. 1.1.3.4), diaphorase from Clostridium kluyven (E.C. 1.6.99.1), catalase from bovine liver (E.C. 1.11.1.6) were purchased from Sigma-Aldrich. Bovine serum albumin (BSA) was purchased from Fisher BioRegents. The alcohol dehydrogenase AdhD from Pyrococcus furiosus was heterologously expressed from E. coli as previously described.29 Sodium phosphate dibasic (Na2HPO4), 3,3’,5,5’-Tetramethylbenzidine (TMB), copper(II) sulfate (CuSO4) and ethylenediaminetetraacetic acid (EDTA) were also purchased form Sigma-Aldrich. Sodium phosphate monobasic (NaH2PO4) and hydrogen peroxidase (H2O2) were purchased from Thermo Fischer Scientific, Fischer Chemical.

Synthesis of Protein-Cu3(PO4)2 Complexes. Protein-Cu3(PO4)2 complexes were prepared using a modified protocol previously described by Ge et al.22 Briefly, complexes were formed by combining 600 µL of 0.5 mg mL-1 protein/enzyme solution in 10 mM PBS buffer (pH 7.4), 300


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µL 20 mM PBS buffer (pH 7.4), and 300 µL of 12 mM CuSO4 in deionized water. Synthesis reactions were carried out in 1.5 mL microcentrifuge tubes. Enzyme solutions were prepared from 1 g L-1 stock solutions. Prior to the addition of CuSO4 to the synthesis mixture, all the solutions were incubated at the desired synthesis temperature (4, 12, 24 or 37 °C). After incubation, 12 mM CuSO4 solution (also pre-heated to the synthesis temperature) prepared in deionized water, was added and complex formation was allowed to proceed for 24 hours. Synthesis temperature was maintained at the desired value by incubating in a temperature controlled shaker-incubator without shaking (Thermo Fisher). To synthesize HRP-Cu3(PO4)2 complexes at different pH values, the method was same as description above with minor modification. 0.5 mg mL-1 HRP solution was prepared in pH 6.8, 7.4 and 8.0 10 mM PBS buffer. Then 600 µL of 0.5 mg mL-1 HRP solution in 10 mM PBS buffer (pH 6.8, 7.4 or 8.0), 300 µL 20 mM PBS buffer (pH 6.8, 7.4 or 8.0), and 300 µL of 12 mM CuSO4 in deionized water were mixed together and incubated at room temperature for 24 h. For the synthesis of core-shell particles, the core protein (CFP or YFP) was first incubated with 12 mM CuSO4 in 10 mM PBS buffer at 4 °C for 24 h. Protein-Cu3(PO4)2 particles were collected by centrifugation (4000 RPM). Secondly, the core particle was incubated with the desired shell protein (YFP or CFP) and 12 mM CuSO4 in 10 mM PBS buffer at 4 °C for another 24 h. The resulting core-shell particles were isolated by centrifugation (4000 RPM).

HRP-Cu3(PO4)2 Particle Density. After 24 hours of complex formation, 10 µL of HRPCu3(PO4)2 synthesis solution was transferred to a standard microscope slide hemocytometer. Particles were observed under bright field using an Olympus BX51 microscope and counted. All particle density measurements were acquired in triplicate.


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Scanning Electron Microscope (SEM) Imaging and Energy Dispersive X-ray Spectroscopy (EDX) Atom Mapping. Immediately after Enzyme-Cu3(PO4)2 complex formation, 2 µL of the synthesis solution was gently transferred to a silicon wafer for imaging. For SEM imaging, samples were coated with Pt/Pd after air drying for 24 h at room temperature. SEM images were acquired using a FEI NNS450 SEM (FEI). For the EDX imaging, samples were air dried for 24 h, but were not coated with Pt/Pd. The EDX atom mapping was obtained by X-Max detector under the Field Free mode of FEI NNS450 SEM.

Thermogravimetric Analysis (TGA). HRP- Cu3(PO4)2 and Cu3(PO4)2 were synthesized as described above and collected by centrifugation at 4000 RPM. In this case, the synthesis of HRPCu3(PO4)2 was carried out in a 20 mL volume of the synthesis reaction described above. For TGA analysis, samples were first held at 100 °C for 2 h to remove water from the complexes. Subsequently, the temperature was increased at a rate of 10 °C min-1 until reaching 500 °C.

Fourier Transform Infrared (FTIR) Spectroscopy. Particles were collected immediately after the synthesis by centrifugation at 4000 RMP using a standard microcentrifuge. The supernatant was discarded and particle aliquots were dried overnight at 37 °C. FTIR spectra of the air-dried particles were collected using a Nicolet 6700 FTIR spectrometer (Thermo) in reflectance mode.

Circular Dichroism (CD) Spectroscopy. Collected enzyme-Cu3(PO4)2 particles were resuspended in 600 µL of 10 mM PBS, pH 7.4. CD spectra were collected in a 1 mm path length quartz cuvette using a Jasco J1500 spectrometer. All spectra were acquired from 180 nm to 350


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nm. The data pitch was 0.2 nm, accumulation number was 3 and the scanning speed was 50 nm min-1.

Enzyme Kinetic Assays. Enzyme-Cu3(PO4)2 particles were collected from the synthesis solution by centrifugation at 4000 RMP for 2 min. Supernatants were removed and saved for kinetic analysis and particles were re-suspended in 120 µL of 10 mM PBS, pH 7.4. Both the resuspended particles and the supernatant were diluted by 200-fold prior to kinetic analysis. Kinetic assays were performed in 96-well flat bottom plates containing 100 µL of 50 mM PBS with 5% v/v diluted particles (or diluted supernatant) and 0.05 mM H2O2 and varying concentrations of 3,3’,5,5’-tetramethylbenzidine (TMB) substrate (0, 2, 4, 10, 20, 40, and 60 µM). A BioTek Synergy 4 Hybrid Microplate Reader was used to detect changes in absorbance at 450 nm every 10 second at room temperature. A molar absorption coefficient of 5.9x104 mol1

cm-1 was used to convert absorption to molar concentrations of oxidized TMB. Reactions were

initiated by adding H2O2 to the reaction mixture. All kinetic assays were repeated in quadruplicate.

Gel Electrophoresis. Collected particles were re-suspended in 120 µL of 60 mM EDTA in deionized water. Incubation in EDTA chelates the copper ions, thus degrading the enzymeCu3(PO4)2 complex. After overnight incubation, SDS-PAGE samples were prepared by combining 3 parts sample with 1 part 4x loading buffer. Samples were incubated at 90 °C for 10 min prior to loading in BioRad anykD precast protein gels.


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Protein Charge and pI. The overall charge and pI of each protein studied here were determined by two independent calculations. Based on the protein amino acid sequence, charge and pI were predicted using the Protein Calculation V3.4 (protcalc.sourceforge.net). These calculated values were in agreement with the charge and pI predicted using the ExPASy online protein calculator (Compute pI/MW; web.expasy.org/compute_pi/).

RESULTS AND DISCUSSION Protein-inorganic complexes were first described by Ge et al. in 2012.22 These hybrid materials were developed as a facile method of creating enzyme aggregates immobilized in high surface-to-volume ratio microscale particles. A schematic of the synthesis method is shown in Figure 1A. First, enzyme molecules form a complex with Cu2+ ions via amide groups of the protein backbone and residue side chains. Second, phosphate crystals form and coprecipitate with Cu2+ complexed enzymes. As synthesis continues (typically 1-3 days at temperatures between 4 and 25 °C), particles grow in size often forming structures that resemble budding flowers. Typical protein-inorganic particles are shown in Figure 1B, where we demonstrate the synthesis of microscale structures fabricated with two different protein components, cyan and yellow fluorescent proteins (CYP and YPF, respectively).


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Figure 1. The synthesis of protein-Cu3(PO4)2 complexes. (A) Schematic diagrams of the synthesis of single and multiprotein systems. (B) SEM images of CFP/YFP- Cu3(PO4)2 complexes, including particles with mixed CFP and YFP, YFP@CFP, and CFP@YFP. The 5 µm scale bar applies to all images.

Particle size and protein position within a particle were controllable. The lefthand images of Figure 1B are ~5 and ~10 µm diameter particles synthesized from mixtures of CFP and YFP. The smaller particles were created from solutions of 0.125 mg mL-1 of protein and 2 mM Cu2+, while the larger particles used 0.5 mg mL-1 of protein with 2 mM Cu2+. The middle images of Figure 1B show core-shell particles with CFP as the core and YFP as the shell (YFP@CFP). Five-µm particles were synthesized from solutions containing 0.25 mg mL-1 of protein and 2.5 mM Cu2+, while 15-20 µm particles were formed from solutions of 0.5 mg mL-1 of protein and 3.0 mM Cu2+. The righthand images show core-shell particles of the opposite configuration, YFP as the core and CFP as the shell (CFP@YFP). Particle size was controlled with the same synthesis condition as YFP@CFP. Core-shell architecture was confirmed by fluorescence


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microscopy showing CYP fluorescence in the interior and YFP fluorescence in the shell (Figure S1). In all cases, the hybrid materials were synthesized at 4 °C with reactions lasting 24 h. The core and shell were synthesized sequentially over 2 days. Reported microstructures of synthesized protein-Cu3(PO4)2 particles vary depending on immobilized protein and synthesis conditions. Those shown in Figure 1 are similar in size and structure to bovine serum albumin (BSA)-Cu3(PO4)2 synthesized at 4 °C by Ge et al.22 After establishing the ability to create hybrid protein-Cu3(PO4)2 microscale particles, we focused on varying synthesis temperature as a parameter for optimizing enzyme loading. As shown in Figure 2, HRP- Cu3(PO4)2 complexes fabricated at 4 °C produced low activity complexes with substantial enzymatic activity remaining in the synthesis solution. Synthesis at 12, 24, and 37 °C resulted in 2.0- to 2.5-fold less HRP activity in solution compared with fabrication at 4 °C (Figure 2A; supernatant). Particles produced at 12 and 24 °C had limited HRP activity, but particles synthesized at 37 °C exhibited 3.5- to 4-fold higher activity (Figure 2A; particles). SDS-PAGE analysis of the HRP-Cu3(PO4)2 hybrid materials suggested that the differences in activity were due to increased HRP loading at 37 °C (Figure 2B). HRP content in the particles was visualized using gel electrophoresis (SDS-PAGE) by first destroying particle structure with the chelating agent EDTA (see Materials and Methods). Based on protein band intensity, 37 °C synthesized materials contained up to 3.5 times as much HRP as complexes synthesized at lower temperatures. These results correspond to the particles activity measurements presented in Figure 2A. In addition, the number density of particles synthesized at each temperature was consistent, with 200-300 HRP-Cu3(PO4)2 particles produced per uL of synthesis volume (Figure S2). Of note, was that all particles exhibited peroxidase activity towards the oxidation of the HRP substrate TMB, and that the observed reaction kinetics


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stemmed from immobilized HRP and not the Cu3(PO4)2 support material, which has been shown to possess weak peroxidase activity (VMAX ~ 3.4x10-9 M s-1, KM ~35 mM).23 Taken together, these results suggest that synthesis at 37 °C creates particles with higher HRP loading then particles synthesized at lower temperatures. To confirm HRP loading, synthesized particles were subjected to thermal gravimetric analysis (TGA; Figure 2C). Synthesis at 37 °C produced 29 wt% HRP in hybrid HRP-Cu3(PO4)2 particles. Consistent with the SDS-PAGE analysis, lower temperatures resulted in lower HRP loadings. Synthesis at 4 and 24 °C yielded complexes with ~16 % HRP, respectively.

Figure 2. Effect of synthesis temperature on HRP activity and immobilization in HRP-Cu3(PO4)2 complexes. (A) The relative activity of HRP-Cu3(PO4)2 particles (right) and particle synthesis


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supernatant (left) at synthesis temperatures of 4, 12, 24, and 37 °C. Supernatant activity was normalized to the highest measure activity, supernatant from the 4 °C synthesis conditions. Particle activity was normalized to the 37 ºC synthesis conditions, the highest measured particle activity. Complex formation was allowed to proceed for 24h prior to collection and kinetic analysis. Activity assays were conducted at room temperature with 60 µM of the HRP substrate TMB and 50 µM H2O2 (50 mM PBS, pH 7.4). The mean and standard deviation of three unique samples are shown. (B) SDS-PAGE analysis of HRP immobilized in HRP-Cu3(PO4)2 complexes synthesized at 4, 24 and 37 °C. (C) HRP loading in particles synthesized at different temperatures as determined by thermal gravimetric analysis (TGA).

In order to study particle microstructure and differences in microstructure due to synthesis temperature, SEM images of formed particles were obtained (Figure 3A). Image analysis revealed that particle size ranged from ~5 to ~10 µm. As previously reported for HRP containing particles, the complexes maintained a microscale structure reminiscent of blooming flowers; this has inspired the common name “nanoflowers”.21-22, 28 More formally, all synthesized particles were observed to be spherical in shape with thin leaflets of Cu3(PO4)2 embedded with HRP enzyme. Only small structural variations were observed over the range of synthesis temperature. For example, at low temperature the leaflets or “petals” of the structure appear to be less well defined and particle shape was somewhat irregular in comparison to the spherical particles produced at higher temperatures. As previously reported, synthesis in the absence of protein does not produce particle structures (Figure 3A, Cu3(PO4)2 control sample).22 With SEM analysis enhanced with energy-dispersive X-ray spectroscopy (EDX) for nitrogen


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elemental mapping, we were able to confirm the presence of HRP protein within the HRPCu3(PO4)2 particles as well as visualize particle structure (Figure 3B). Nitrogen was detected across the imaged area, but was found in higher density in areas that overlapped the SEM imaged HRP-Cu3(PO4)2 particles. Phosphate, copper, and sodium mapping also overlapped with imaged particles (Figure S3).

Figure 3. Scanning electron microscope (SEM) imaging of HRP- Cu3(PO4)2 particles. (A) SEM images of 4, 24 and 37°C synthesized HRP-Cu3(PO4)2 complexes and 24 °C synthesized Cu3(PO4)2 . All samples were air dried and coated with Pt/Pd prior to imaging. (B) SEM imaging of HRP-Cu3(PO4)2 particles and nitrogen mapping by EDX.


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FTIR analysis also supports the conclusion that HRP was immobilized within the Cu3(PO4)2 complexes. As shown in Figure 4A, a peak characteristic of phosphate at 1050 cm-1 was observed in HRP- Cu3(PO4)2 particles and Cu3(PO4)2 control sample (Figure S4). This peak is characteristic of O-P vibration and stretching, thus indicating the presence of phosphate in the sample. CD spectra of these samples also suggested that immobilization did not significantly alter HRP secondary structure (Figure 4B). While it appears that Cu3(PO4)2 complexes increase the signal to noise ratio of the CD spectra, characteristic α-helical secondary structure was maintained in the immobilized state. Specifically, negative peaks at 222 and 208 nm and a positive peak at 190 nm suggest strong α -helical structure typical of HRP.

Figure 4. Spectroscopy analysis of HRP-Cu3(PO4)2 particles. (A) FT-IR spectra of 37 °C synthesis particles. (B) CD spectra of HRP and HRP-Cu3(PO4)2 particles. CD spectral peaks characteristic of α-helical secondary structure (positive CD signal at 190 nm and negative CD signal at 208 and 222 nm) were observed.


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Given the observed trend of enhanced HRP loading with high synthesis temperature, it was critical to determine if enzyme activity followed the same trend. With TMB as an HRP substrate and H2O2 as a cosubstrate, kinetic analysis revealed increased particle activity with 37 °C synthesis (Figure 5). HRP-Cu3(PO4)2 synthesis at 4 and 24 °C produced enzyme complexes with apparent Michaelis constants (KM,app) of 1.9±0.4 and 1.6±0.4 µM and VMAX values of 43±2 and 56±4 µM s-1, respectively. Synthesis at 37 °C increased VMAX by 3- and 4-fold to 177±12 µM s-1, while KM,app increased slightly to 2.9±0.6 µM. The increase in catalytic capacity matches the measured increase in HRP loading, thus providing evidence that the 37 °C particle synthesis does not lead to significant enzyme deactivation and that higher loadings do not adversely affect kinetics.

Figure 5. Kinetic analysis of HRP-Cu3(PO4)2 hybrid complexes synthesized at different temperatures. (A) Michaelis-Menten kinetic analysis with TMB as a substrate and 50 µM H2O2 as a cosubstrate. The relative reaction velocity, ν, normalized to VMAX achieved with particle synthesis at 37 °C is shown. Each assay was conducted with equal amounts of synthesized


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material in 50 mM phosphate buffer, pH 7.4. The means and standard deviation of 3 unique samples is shown. (B) VMAX and KM,app parameters resulting from the kinetic analysis.

Seeking to extend our findings of the temperature-dependent immobilization of HRP in Cu3(PO4)2 complexes, we investigated the synthesis of enzyme-inorganic particles with glucose oxidase (GOx), a thermostable alcohol dehydrogenase (AdhD), diaphorase, catalase, and BSA. The enzymes were selected because they are catalytically useful: GOx and HRP are a wellestablished cascade for glucose detection;30 AdhD oxidizes secondary alcohols and is thermostable to 90 °C;29, 31 diaphorase can recycle NAD(H) cofactor for biotechnology applications;32 and, catalase is widely used in various biotechnologies to decompose and detect H2O2.6, 33 BSA is a common model protein and is widely used as standard for protein assays. The set of proteins was also selected because, together with HRP, they represent a wide range of overall protein charge at pH 7.4 (the standard synthesis condition) and cover a range of isoelectric points. Using densitometric analysis of SDS-PAGE imaging of EDTA-dissolved particles as a proxy for enzyme amount, we quantified enzyme loading as a function of synthesis temperature (Figure 6A). The optimum synthesis temperature for HRP complexes was 37 °C, while GOx, AdhD, Diaphorase, catalase and BSA had highest enzyme loadings at 4 °C. In some cases (see GOx and catalase) there was minimum difference in enzyme loading at 4, 12, and 24 °C, but, with the exception of HRP, loading was significantly reduced with 37 °C synthesis. As presented in Figure 6B, optimum synthesis temperature correlated with overall enzyme charge. Proteins with pI values less than the synthesis pH maintained an overall negative charge, which corresponded to enhanced loading at lower temperature. HRP has a predicted pI


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of 7.5, maintaining a positive charge when synthesis was buffered to pH 7.4, and exhibited an optimum synthesis temperature is 37 °C. We directly tested the hypothesis that overall charge correlates with enzyme loading by repeating the synthesis of HRP-Cu3(PO4)2 particles at pH 6.8, 7.4, and 8.0 (Figure 6C). This range of synthesis conditions resulted in the predicted overall charge of HRP decreasing from +2.2 at pH 6.8 to -2.2 at pH 8.0. Correspondingly, HRP loading increased by ~20% at pH 7.4 and ~40% at pH 8.0.

Figure 6. Protein loading as a function of synthesis temperature. (A) The relative change in protein loading in protein-Cu3(PO4)2 particles as a functional of synthesis temperature for BSA, GOx, AdhD, catalase, diaphorase, and HRP. Protein loading was normalized to the synthesis condition that produced the lowest loading (4 °C for HRP and 37 °C for all other proteins). (B) Summary of optimum synthesis conditions for each of the studied proteins. (C) The relative HRP loading amount in particles that were synthesized at pH 6.8, 7.4, and 8.0.


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The synthesis temperature dependency of enzyme loading in Cu3(PO4)2 particles suggests that the interfacial chemistry between protein, Cu2+ ions, and precipitating phosphate strongly affects enzyme loading. Our data suggests that there are at least two effects at play. First, the overall charge of proteins with pI values lower than the synthesis pH (i.e., BSA, GOx, AdhD, catalase, and diaphorase) was negative, while with a predicted pI of 7.5 the net charge of HRP was positive. Electrostatic repulsion between positively charged HRP and Cu2+ is unfavorable for ion binding. Higher temperatures partially alleviate the electrostatic effect because phosphate buffer pH is a function of temperature with pH increasing with temperature.34 At 37 °C, the solution pH is expected to increase by ~0.2 to 7.6, thus reducing overall charge of HRP and reducing electrostatic repulsion to Cu2+ ions in solution. Our study of HRP loading over a range of pH synthesis conditions supports this idea (Figure 6C). Moreover, previous work suggests that Cu2+ binding to proteins is improved under basic conditions.35 Most enzymes have a pI less than neutral pH, therefore synthesis under basic conditions would preference Cu2+ as the overall protein charge is more negative in comparison with neutral or acidic conditions. Secondly, protein-Cu3(PO4)2 complex formation is spontaneous and Cu2+ ion-protein binding is known to be an entropy driven process.36 Higher temperatures favor entropic processes, thus benefiting the formation of HRP-Cu3(PO4)2 complexes. It is important to note that these two effects (overall charge and an entropy driven process) cannot fully explain the observed variability in loading enhancement across different proteins. For example, BSA has the lowest predicted charge (-18), but loading was only enhanced by 1.65-fold. By comparison, diaphorase loading was enhanced by 2.6-fold with a predicted overall charge of -2.9. We speculate that such differences are likely


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due to the effective number of Cu2+ ion binding sites. However, a clear understanding of the binding interactions between Cu2+, or Cu3(PO4)2 inorganic crystals, and the outer surface of protein structures has not yet by been elucidated. Such interactions are the focus of on-going studies. In the context of our results, it is interesting to compare previously reported enzyme wt%, synthesis temperatures, and pI values for various enzymes. To date, it appears that the highest enzyme loading in Cu3(PO4)2 complexes is ~40 wt% papain, an enzyme of the cysteine protease family. Papain from papaya root is negatively charged under the synthesis conditions (pI of 4.4, synthesis at pH 7.4), and the high loadings where achieved with room temperature synthesis and a high concentration of enzyme in the synthesis solution (2 mg mL-1).19, 37 Similar to our results, 17 wt% HRP was achieved with room temperature synthesis at pH 7.4 over a 3 day period.21 Room temperature synthesis has also been used to immobilize GOx at 10 wt%.7 Lastly, trypsin and chymotrypsin proteases have been immobilized in Cu3(PO4)2 particles at ~13 and ~10 wt%, respectively.25-26 The pI of trypsin isoforms range from 4 to 6, while chymotrypsin from bovine pancreas (commonly available through commercial vendors) has a pI above 8. As we demonstrated with HRP, the high pI of chymotrypsin suggests that higher enzyme loadings could be achieved with higher synthesis temperatures. Conversely, synthesis of GOx-, trypsin-, and papain-Cu3(PO4)2 particles would benefit from incubation at 4 °C. If an enzyme is unstable and sensitive to thermal deactivation, lower synthesis temperatures may also be preferred over higher loading achieved at 37 °C (for example, in the case of chymotrypsin). The studies presented here focus on the immobilization of single enzymes in enzymeCu3(PO4)2 hybrid materials, but importantly the capability to create multienzyme structures for cascade catalysis has been demonstrated (Figure 1 and ref. 7). The synthesis of multienzyme


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complexes, including core-shell particles that spatial localize different enzyme reaction steps within a particle and systems with enzymes mixed throughout the particle, would also benefit from the temperature-dependent immobilization described here. For example, wt% loading of GOx in particle cores could be maximized with synthesis at 4 °C, while HRP in the outer shell could be fabricated at 37 °C. Alternative synthesis temperature-profiles could also be used to controlled the ratio of enzyme activities within a core-shell structured particle, an aspect of cascade catalysis that is critical to high reaction cascade yields.

CONCLUSIONS Enzyme immobilization in nano- and microscale structures can enhance catalytic activity, increase stability, and create multienzyme systems with controlled spatial organization. In this work, we investigate a recently developed hybrid materials technology that immobilizes enzymes and other proteins into high surface-to-volume ratio particles of Cu3(PO4)2. Particle synthesis is spontaneous as Cu2+ ions bound to protein help nucleate inorganic crystal growth, thus creating hybrid enzyme-Cu3(PO4)2 complexes. The materials technology is useful for cascade catalysis as well as for single enzyme application that requires immobilization, because particle structure and the spatial organization of enzymes within a particle can be controlled. Our key finding is that enzyme loading was found to be dependent on both synthesis temperature and protein charge. With an overall positive charge under the studied synthesis conditions, HRP loading was enhanced by 4.2-fold when synthesis was accomplished at 37 °C. HRP activity increased concomitantly with the increased loading. Conversely, proteins with overall negative charges (BSA, GOx, AdhD, catalase, and diaphorase) exhibited lower enzyme loadings at 37 °C and enhanced loading with 4 °C synthesis. These insights into controlling enzyme immobilization in


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protein-Cu3(PO4)2 provide new information on the mechanisms for formation and promise to significantly advance this hybrid materials technology for a broad range of enzyme and multienzyme systems.

ASSOCIATED CONTENT The supporting Information is available free of charge on the ACS Publication website. Fluorescence images of YFP@CFP protein-Cu3(PO4)2 particles; quantification of number particle density described in the text (PDF).

AUTHOR INFROMATION Corresponding Author *E-mail: [email protected] Current Address ##

Yingning Gao, Nanomedical Diagnostics Inc., 6185 Cornerstone Court East Suite #110, San

Diego, CA 92121, U.S.A. Author Contributions YG and IW conceived the study. XL, LZ, YG, and IW designed and analyzed the experiments. XL and LZ conducted the experiments. All authors wrote and edited the manuscript. Notes: The authors declare no competing financial interests.

ACKNOWLEDGMENTS This work was supported by the Defense Threat Reduction Agency (DTRA; HDTRA1-12-10045).


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Figure 1. The synthesis of protein-Cu3(PO4)2 complexes. (A) Schematic diagrams of the synthesis of single and multiprotein systems. (B) SEM images of CFP/YFP- Cu3(PO4)2 complexes, including particles with mixed CFP and YFP, YFP@CFP, and CFP@YFP. The 5 µm scale bar applies to all images. 177x74mm (300 x 300 DPI)


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Figure 2. Effect of synthesis temperature on HRP activity and immobilization in HRP-Cu3(PO4)2 complexes. (A) The relative activity of HRP-Cu3(PO4)2 particles (right) and particle synthesis supernatant (left) at synthesis temperatures of 4, 12, 24, and 37 C. Complex formation was allowed to proceed for 24h prior to collection and kinetic analysis. Activity assays were conducted at room temperature with 60 µM of the HRP substrate TMB and 50 µM H2O2 (50 mM PBS, pH 7.4). The mean and standard deviation of three unique samples are shown. (B) SDS-PAGE analysis of HRP immobilized in HRP-Cu3(PO4)2 complexes synthesized at 4, 24 and 37 C. (C) HRP loading in particles synthesized at different temperatures as determined by thermal gravimetric analysis (TGA). 89x119mm (300 x 300 DPI)


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Figure 3. Scanning electron microscope (SEM) imaging of HRP- Cu3(PO4)2 particles. (A) SEM images of 4C, 24C, 37C synthesized HRP-Cu3(PO4)2 complexes and 24 C synthesized Cu3(PO4)2 particles. All particles were air dried and coated with Pt/Pd prior to imaging. (B) SEM imaging of HRP-Cu3(PO4)2 particles and nitrogen mapping by EDS. 159x128mm (300 x 300 DPI)


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Figure 4. Spectroscopy analysis of HRP-Cu3(PO4)2 particles. (A) FT-IR spectra of 37 C synthesis particles. (B) CD spectra of HRP and HRP-Cu3(PO4)2 particles. CD spectral peaks characteristic of α-helical secondary structure (positive CD signal at 190 nm and negative CD signal at 208 and 222 nm) were observed.

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Figure 5. Kinetic analysis of HRP-Cu3(PO4)2 hybrid complexes synthesized at different temperatures. (A) Michaelis-Menten kinetic analysis with TMB as a substrate and 50 µM H2O2 as a cosubstrate. The relative reaction velocity, , normalized to VMAX achieved with particle synthesis at 37 C is shown. Each assay was conducted with equal amounts of synthesized material in 50 mM phosphate buffer, pH 7.4. The means and standard deviation of 3 unique samples is shown. (B) VMAX and KM,app parameters resulting from the kinetic analysis. 72x55mm (300 x 300 DPI)


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Figure 6. Protein loading as a function of synthesis temperature. (A) The relative change in protein loading in protein-Cu3(PO4)2 particles as a functional of synthesis temperature for BSA, GOx, AdhD, catalase, diaphorase, and HRP. Protein loading was normalized to the synthesis condition that produced the lowest loading (4 C for HRP and 37 C for all other proteins). (B) Summary of optimum synthesis conditions for each of the studied proteins. (C) The relative HRP loading amount in particles that were synthesized at pH 6.8, 7.4, and 8.0.

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Enhancing Enzyme Activity and Immobilization in ... - ACS Publications

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