Insights into sustainable glucose oxidation using magnetically

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Insights into sustainable glucose oxidation using magnetically recoverable biocatalysts Bret Lawson, Ekaterina Golikova, Aleksandrina Sulman, Barry D. Stein, David Gene Morgan, Natalya V. Lakina, Alexey Yu. Karpenkov, Esther M. Sulman, Valentina Matveeva, and Lyudmila M. Bronstein ACS Sustainable Chem. Eng., Just Accepted Manuscript • DOI: 10.1021/acssuschemeng.8b01009 • Publication Date (Web): 16 Jul 2018 Downloaded from http://pubs.acs.org on July 17, 2018

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ACS Sustainable Chemistry & Engineering

Insights into sustainable glucose oxidation using magnetically recoverable biocatalysts Bret P. Lawson1, Ekaterina Golikova2, Aleksandrina M. Sulman2, Barry D. Stein3, David G. Morgan1, Natalya V. Lakina2, Alexey Yu. Karpenkov4, Esther M. Sulman2, Valentina G. Matveeva2,4*, Lyudmila M. Bronstein1, 5, 6* 1

Indiana University, Department of Chemistry, 800 E. Kirkwood Av., Bloomington, IN 47405, USA

2

Tver State Technical University, Department of Biotechnology and Chemistry, 22 A. Nikitina St, 170026, Tver, Russia

3

Indiana University, Department of Biology, 1001 E. Third St., Bloomington, IN 47405, USA 4

Tver State University, Regional Technological Center, Zhelyabova Str., 33, 170100, Tver, Russia

5

A.N. Nesmeyanov Institute of Organoelement Compounds, Russian Academy of Sciences, 28 Vavilov St., Moscow, 119991 Russia

6

King Abdulaziz University, Faculty of Science, Department of Physics, Jeddah, 21589 Saudi Arabia

*

[email protected]; [email protected]

1 Environment ACS Paragon Plus

ACS Sustainable Chemistry & Engineering 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

KEYWORDS. D-glucose, glucose oxidase, magnetite nanoparticles, clusters, mobility

ABSTRACT. Here, we developed magnetically recoverable biocatalysts for enzymatic oxidation of D-glucose to D-gluconic acid with high product yields. The catalyst support is based on nanoparticle clusters (NPCs) composed of magnetite particles and coated with the amino terminated silica layer to facilitate further functionalization. It involves the attachment of the glutaraldehyde linker followed by the covalent attachment of glucose oxidase (GOx) via its amino groups. It was established that the NPCs with a diameter of ~430 nm attach 33% more GOx molecules than NPCs with a diameter of ~285 nm, although the surface area of the former is lower than that of the latter. At the same time, the biocatalyst based on the smaller NPCs shows higher relative activity of 94% than that (87%) of the biocatalyst based on the larger NPCs, both at 50 °C and pH 7 (optimal reaction conditions). This surprising result has been explained by a combination of two major factors such as GOx crowding on the support surface which should prevent denaturation (similar to the enzyme behavior in cells) and the enzyme mobility which should be preserved upon immobilization. Apparently, for the biocatalyst based on 285 nm NPCs, the lower GOx crowding is compensated by its higher mobility. The high stability of these GOx based biocatalysts in ten consecutive reactions as well as facile magnetic recovery combined with excellent catalytic activity in “tolerant” pH range make this biocatalyst design promising for other types of enzymatic catalysts.

Introduction The oxidation of monosaccharides, in particular D-glucose, received considerable attention because of the importance of D-gluconic acid and its salts (gluconates) utilized as drugs, food


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supplements and additives, components of cleaning products, etc.

1-2

Due to the multifunctional

character of D-glucose, however, its oxidation often produces a number of side products depicted in Scheme 1. A selective oxidation of the hemiacetal group without oxidation of hydroxyl groups demands selective and efficient catalytic systems.

Scheme 1. Possible pathways of D-glucose oxidation. Chemical or electrochemical oxidation of D-glucose in the presence of different oxidants is used in the industrial D-gluconic acid production, but the selectivity of the process and the target product yield are generally low because of the formation of side products.1-2Also, the use of inert, anti-corrosion materials for the reactor lining and large amounts of oxidants have an adverse environmental impact, making these processes ecologically unsustainable.1, 3 In recent years heterogeneous catalysts have been actively explored in monosaccharide oxidation. The catalytic oxidation of D-glucose using mono- and bimetallic catalysts containing Pt, Pd, Bi, Au, Rh, Tl, Sn, or Co have been reported.

1, 4-11

The use of heterogeneous catalysts is



definitely a step forward compared to chemical or electrochemical oxidation because they can be separated from the reaction medium and allow for using green oxidizers such as oxygen, peroxide and atmospheric air.12 The highest selectivity (99-100%) to D-gluconic acid was reported for gold containing catalysts.13-15 However, in many cases the conversion was significantly below 100%. For example, in ref. 14 the maximum D-glucose conversion did not exceed 91%, while in ref.15 it was 67%. Another frequent shortcoming of D-glucose oxidation with metal containing heterogeneous catalysts is a need for a basic medium which leads to the formation of alkali gluconates instead of D-gluconic acid. In industry this will result in an additional step of the gluconate protonation.2 This and possible leaching of catalytic metals hamper their practical applications. Microbial oxidation of D-glucose has been actively explored because many bacteria possess an enzyme required for its oxidation in mild conditions and with high selectivity. Different microbial strains such as Gluconobacter spp., Aspergillius niger, Tricholoma robustum and Tricholomabakamatsutake, Gluconobacter oxydans, Zymomonas mobilis, Penicilliumvariabile, Acetobacter methanolicus, Acetobacter diazotrophicus and Acetobacter suboxydans as well as microbial-derived enzymes have been utilized.11, 16-20 The use of enzymes in organic synthesis has numerous advantages such as (i) mild reaction conditions in terms of temperature, pressure, and pH; (ii) low energy demand; (iii) high enantio-, regio- and chemoselectivity of the process. However, the key shortcomings are high costs of enzymes and their single use because their separation is rather unfeasible. These shortcomings were greatly overcome when immobilized enzymatic catalytic systems (biocatalysts) have been developed. 21-22 The advantages of immobilized enzymes in comparison with their native form are the enzyme stability increase, a reuse or a long-term use, reaction rate control, the separation from reaction


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products, the possibility to stop the reaction at any time, enzyme catalytic property modulation, and the prevention of the reaction product contamination.21-23Thus, the enzyme immobilization can directly affect the decrease in the process cost and the increase of the quality of target products.24-25 The immobilization can also lead to a change of enzyme properties.26-28 By choosing the immobilization method, for example, the temperature range of the enzymatic activity can be improved, allowing, in some cases, for the reaction to be conducted at higher temperatures, thereby increasing the reaction rate and product yield.21-28 Currently, the studies of biocatalysts are mainly focused on evaluating the influence of (i) the nature of support, modifier and crosslinking agent; (ii) the introduction of a second enzyme; (iii) conditions of the enzyme immobilization and oxidation on the catalytic process. 21-22, 24-28 The enzyme immobilization on solid supports allows for separation of the catalysts via filtration or sedimentation. However, neither process is energetically favorable. They are time and energy consuming and result in catalyst losses. Magnetically recoverable catalysts allow for easy magnetic separation, leading to energy conservation, cheaper and more pure target products.29-37In a number of cases, iron oxide nanoparticles (NPs) giving magnetic properties to the catalyst, may enhance the catalytic activity via an electron transfer38-39 or due to changes in the reaction mechanism, favoring target molecules.40-41 The immobilization of enzymes on magnetic NPs or magnetic supports is an upcoming trend as well.42-46 Native glucose oxidase (GOx, enzyme of oxidoreductase group EC 1.1.3.4) is employed in the D-glucose selective oxidation.18-20, 47 Carbon nanotubes48-49 and silica50 have been utilized for a glucose oxidase immobilization. First examples of the GOx immobilization on a magnetic support in fluidized bed enzyme reactors have been reported in 70’s and early 80’s.51-52Pieters et al. describedthe GOx immobilization on a magnetic support via modification of the latter with



polyethyleneimine (PEI) followed by the reaction with glutaraldehyde for connecting with the enzyme.53 However, the use of hazardous sodium periodate as an oxidizer made this process environmentally unsound. An interesting example of recyclable multienzyme nanosystems,where magnetic metal-organic framework was used as scaffold, was reported in ref.54. These nanosystems contained two enzymes(one of which was GOx)with different spatial colocalizations that influenced the cascade reaction kinetics and operational stability of immobilized enzymes. However, the maximum D-glucose activity did not exceed 80.6%. In a recent work, Yang et al. synthesized a magnetic enzymatic hybrid catalyst formed by the coating of Fe3O4@C NPs with GOx attached to PEI, followed by the reaction with H2SiO3 for insitu biomineralization.55 This catalyst allowed 81% activity compared to the native enzyme. Although the protection of the enzyme with silica allowed for a significant stability increase, the silica coating may also impede the access of D-glucose molecules to the enzyme. In the present work we developed a sustainable D-glucose oxidation with magnetically recoverable biocatalysts based on magnetite NP clusters (NPCs) with diameters of 285±71 or 430±60 nm to test the influence of the NPC size and the surface area on the catalysis. The NPC are coated with a layer of silica modified with amino groups and used for the GOx covalent attachment via the reaction with glutaraldehyde. We established that NPC sizes affect both the amount of immobilized GOx and the enzyme catalytic activity. Moreover, we demonstrate that these seemingly disjointed factors converge when several aspects influencing the GOx properties are considered. Exceptional catalytic activity and stability of the catalysts developed make them promising for practical applications.


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Experimental Materials Iron (III) chloride hexahydrate (99%), ammonium hydroxide solution (28%), glycerol (≥ 99.5%),

tetraethyl

orthosilicate (TEOS,

99%),

glucose oxidase

lyophilized

powder

(Aspergillusniger, 174.9 U/mg), glutaraldehyde solution (GA, 25%) and PBS buffer were purchased from Sigma-Aldrich and used as received. APTES ((3-aminopropyl)triethoxysilane, 98%) was purchased from Fluka and used without purification. Ethylene glycol (99.5%), succinic acid (99%), and urea (99%) were all purchased from TCI and used without purification. Ethanol was received from Pharmco-Aaper and used as received.

Synthesis of magnetite NPCs The synthesis of magnetite NPCs was carried out according to the procedure reported elsewhere.56 In a typical experiment, 0.8109 g of FeCl3×12H2O (3 mmol), 0.1181 g of succinic acid (1 mmol), and 1.8018 g of urea (30 mmol) were completely dissolved in 30 mL (NPC-1) or 20 mL (NPC-2) of ethylene glycol via vigorous stirring. The resultant solution was placed into a Teflon-lined stainless steel autoclave and heated to 200˚C for 48 hours. The reaction was allowed to cool to room temperature and washed three times with water and three times with ethanol. The sample was dried in a vacuum oven at 60°C overnight.

Coating of NPCs with silica (NPC-S) Coating of iron oxide NPCs with a silica shell was carried out according to the published procedure.57 In a typical experiment, 0.19 g of dry NPCs were dispersed in 3 mL of ethanol via sonication for 4 hours at room temperature. To the dispersed NPCs, 0.222 mL of TEOS and 0.633 mL of ammonia hydroxide were added. The reaction was carried out at 40°C for 12 hours. NPCs coated with silica were separated from the reaction solution and washed three times with



water and three times with ethanol, using magnetic separation each time. The material was dried in vacuum at 60°C overnight.

Functionalization with APTES (NPC-SA) NPC-S (0.3 g) was suspended in 8 mL of the APTES aqueous solution. For the preparation of this solution, glacial acetic acid was added to 10 mL of water until pH 4 was reached. Then 8 mL of the acidified water was mixed with 0.8 mL of APTES and used to suspend the support. After that, 4 mL of glycerol were added. The reaction was carried out under stirring at 90 °C for 5 h. Then, the support was washed with water (three times) and methanol (five times) and dried in a vacuum oven overnight.

Attachment of GA and GOx (NPC-SA-GOx) The dry NPC-SA sample (0.10 g) was added to 20 mL of glutaraldehyde (GA) solution in the PBS buffer and stirred for 1 h. To prepare the GA solution, 0.08 mL of 25% GA in water were mixed with 20 mL of the PBS buffer at pH 7.0. The support functionalized with GA was magnetically separated and washed with water five times. At the same time, 10 mg of GOx were incubated and stirred in 20 mL of the PBS buffer for 1 h. The GOx solution was added to the GA functionalized support and stirred for 1 h. Then the biocatalyst was magnetically separated. The amount of GOx attached was evaluated determining immobilization coefficient (IC):58-59 IC = (C0-C1)/C0 × 100% (1) where C0 is the initial GOx amount (µg/mL) and C1 is the GOx amount (µg/mL) in the filtrate after immobilization and the biocatalyst magnetic separation. The amount of GOx in the solution before and after immobilization was determined via the activity in the D-glucose oxidation, considering that in all experiments the initial GOx activity was 174.9 U/mg.


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Characterization Electron-transparent specimens for transmission electron microscopy (TEM) were prepared by placing a drop of a sample suspension onto a carbon-coated Cu grid. Images were acquired at an accelerating voltage of 80 kV on a JEOL JEM1010 transmission electron microscope. The images were analyzed with an image-processing package ImageJ (the National Institute of Health) to estimate nanoparticle diameters. Scanning TEM (STEM) energy dispersive X-ray spectra (EDS) were acquired at an accelerating voltage of 300 kV on a JEOL 3200FS transmission electron microscope equipped with an Oxford Instruments INCA EDS system. The same TEM grids were used for both analyses. For scanning electron microscopy (SEM), samples were drop cast onto the aluminum SEM stubs. The samples were sputter coated using a Polaron Equipment Ltd,SEM Coating Unit E5100, with a gold/palladium target (Au 60%, Pd 40%) for 2 minutes at 20 mA for a coating of approximately 30 nm. They were imaged on a FEI Quanta 600F with the Everhart Thornley detector at an accelerating potential of 10 kV. X-ray powder diffraction (XRD) patterns were collected on an Empyrean from PANalytical. X-rays were generated from a copper target with a scattering wavelength of 1.54 °A. The step size of the experiment was 0.02. Magnetic measurements were performed on a Quantum Design PPMS-14 magnetometer using the system with DC measurement capabilities. The sample (50 mg) was placed in a standard gelatin capsule. Nitrogen adsorption measurements were carried out at liquid nitrogen temperature on an ASAP 2020 analyzer from Micromeritics. Samples were degassed at 100 °C in vacuum. The total surface area was estimated by the Brunauer–Emmett–Teller (BET) method, while the pore size distribution was determined by the Barrett–Joyner–Halenda (BJH) method using desorption.



D-glucose oxidation and product analysis D-glucose oxidation was carried out at atmospheric pressure in the temperature range of 30 60 °С in a double-jacketed three-neck round-bottom flask equipped with a gas inlet, overhead stirrer, and a reflux condenser, serving as gas outlet. The reaction temperature was maintained by circulating a heating medium in the flask jacket. After reaching the required temperature, 15 mL of the 0.1 M phosphate buffer (pH 7), 10 mg of D-glucose and 0.11 g of the biocatalyst were loaded in the flask and kept stirring for 60 min. Oxygen with a feeding rate of 440 – 450 mL/min was used as oxidant. After the reaction, the catalyst was separated with a rare-earth magnet and the reaction mixture was analyzed using HPLC, UltiMate 3000 (ChromaTech, Russia) equipped with a refractometer detector and a ReproGel H Column (500x10 mm, NTP 160000). The H2SO4 solution (9 mM) was used as eluent at the 0.5 mL/min rate for 30 min at the eluent pressure of 6.5 kPa and the column temperature of 250°С. Only pure D-gluconic acid was used for the product identification because selectivity of the process is 100%. The catalytic activity (A) was quantified using Eq. (2):60 A = Glr /([GOx] × V × τ) (2) where Glr is the amount D-glucose (mmol) reacted, [GOx] is the amount (mg) of native or immobilized GOx, V is the volume of the reaction solution (mL), and τ is the reaction time (60 min in all experiments). The relative activity was calculated as AIE/A0 × 100%, where AIE is the activity of immobilized GOx and A0 is the activity of native GOx.

Results and discussion Catalyst synthesis and characterization The synthesis of biocatalysts has been carried out in several simple steps depicted in Scheme 2. First, magnetic iron oxide NPCs have been synthesized in a polyol process. The NPCs were


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coated with a silica layer (NPC-S), functionalized with amino groups via the APTES attachment (NPC-SA) followed by the binding of the GA linker to create aldehyde groups on the support for the GOx attachment (NPC-SA-GOx).

Scheme 2. Schematic representation of the biocatalyst synthesis via the GOx immobilization on the magnetite NPCs (a). The first step is coating of NPCs with silica using TEOS (b, NPC-S), then functionalization with amino groups via the APTES attachment (c, NPC-SA), followed by the reaction with GA to create terminal aldehyde groups (d), which are reacted with amino groups of GOx (e), allowing for its immobilization (NPC-SA-GOx). For the synthesis of NPCs by the polyol process, we used ethylene glycol as both solvent and reducing agent. This procedure is known to generate iron oxide NPs which tend to assemble into NPCs.56 Varying the iron precursor concentration, we were able to vary the NPC size. The TEM images of NPCs of 430±60 nm (NPC-1) and 285±71 nm (NPC-2) in diameter are displayed in Figure 1. The SEM image of NPC-2 (Fig. 1c) clearly shows that the NPCs are composed of individual NPs of ~ 42-45 nm in diameter.



Figure 1. TEM images of NPC-1 (a) and NPC-2 (b), SEM image (c) and XRD pattern (d) of NPC-2. The scale bar in the SEM image is 1 µm. A representative XRD pattern of iron oxide NPCs (Fig. 1d) shows a set of reflections which are characteristic of a spinel structure, most probably magnetite, Fe3O4 (due to the presence of ethylene glycol which is a reducing agent), according to the standard card (JCPDS 19-629).61 The average crystallite size calculated from the Scherrer equation for the (311) reflection is 43 nm, indicating that the NPCs consist of multiple nanocrystals. This is in a good agreement with the SEM data. The magnetization curves for both NPC samples presented in Figure S1 (the Supporting Information, SI) demonstrate that the saturation magnetization values are 93 and 74 emu/g for NPC-1 and NPC-2, respectively. The former value is close to that of bulk magnetite.62


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The characteristic nitrogen adsorption-desorption isotherms and pore size distribution for NPC-2 are shown in Figure S2 (SI). The BET surface area of this sample is 11.9 m2/g. The isotherms resemble Type IV with the H1 hysteresis loop,63 demonstrating that mesoporosity most probably comes from the spaces between NPCs rather than from the inherent porosity of the NPCs themselves. At the same time, a close look at the TEM images of the NPCs (Fig. 1) shows some spaces with a lower electron density within individual NPCs. These spaces are most likely internal pores formed during NP clustering and Ostwald ripening within NPCs.56 Because they do not have interconnected pore structure which opens to the NPC surface, they do not contribute to the porosity. The BET surface area of NPC-1 is 3.8 m2/g. Assuming that NPCs are not aggregated and have the same density of ~5 g/cm3 (the Fe3O4 density is 5.15 g/cm3, while for SiO2, it is 2.65 g/cm3), the calculated surface area of 430 nm NPCs is estimated to be 2.8 m2/g. For the 285 nm NPCs, the calculated surface area is 4.2 m2/g. Thus, the larger surface areas observed for NPC-1 (3.8 m2/g) and NPC-2 (11.9 m2/g) samples could be associated with a patterned (not smooth) surface of the NPCs formed by the assembly of 42-45 nm NPs (see Fig. 1c). The SEM and TEM images of the NPCs coated with silica and then functionalized with APTES are presented in Figure 2.



Figure 2. SEM images (a, b) and TEM images (c, d) of NPC-S-2 (a, c) and NPC-SA-2 (b, d). The red arrows in (c) and (d) show the silica layer which has a lower electron density than that of the NPCs. The scale bars in the SEM images are 1 µm. The SEM image of NPC-S-2 (Fig. 2a) shows that upon silica coating the surface becomes much smoother. The silica layer is clearly visible in the TEM image of this sample (Fig. 2c) and is approximately 27 nm thick. Functionalization with amino groups (NPC-SA-2) does not change the sample morphology (Fig. 2b), but the silica layer thickness increases to ~32 nm (Fig. 2d), which is most likely due to the addition of the condensed APTES. The STEM dark-field image and EDS maps of NPC-S-2 are presented in Figure 3. The EDS map of Fe shows solid NPCs combined in an aggregate (similar to the dark-field image). The EDS map of Si displays the same shape of the NPC aggregate with hollow particles which is in


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agreement with a silica shell. The superposition of the Fe and Si maps clearly shows core-shell particles with the Fe-containing core and the Si-containing shell. The BET surface areas for NPC-S-1 and NPC-S-2 decrease dramatically to 1.2 m2/g and 2.7 m2/g, respectively, compared to the values obtained for the NPCs without silica coating. This is consistent with both NPC aggregation and smoothing of the NPC surface with silica.

Figure 3. STEM dark-field image (a) and EDS maps of NPC-S-2 for Fe (b), Si (c), and their mix (d). White rectangle in (a) shows the area of EDS mapping. Scale bar in (a) is 1 µm. Scale bars in the EDS maps are 500 nm. After functionalization with GA and the attachment of GOx, the amount of the enzyme attached to the catalyst (immobilization coefficient) has been determined. A standard procedure to assess the amount of immobilized GOx is to evaluate the catalytic activity of the supernatant



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after the enzyme attachment and the catalyst separation (see the Experimental Part). According to these data, NPC-SA-GOx-1 contains 66% of GOx out of the total amount used for functionalization, while NPC-SA-GOx-2 contains only 41%. At the same time, the surface area of the former is less than 50% of the latter, indicating the higher packing density of GOx molecules on the NPC-SA-GOx-1 surface than that on the NPC-SA-GOx-2 surface.

It is

noteworthy that the increase of the GOx loading during immobilization up to 15 mg (normally, 10 mg of GOx was used) did not change the amount of the immobilized enzyme, revealing that a finite number of the GOx molecules can be accommodated on the magnetic support surface and saturation of this capacity was achieved for both NPC-SA-GOx-1 and NPC-SA-GOx-2. Considering that NPC-SA-GOx-1 with a lower surface area accommodates more GOx molecules creating a higher GOx surface packing density, the difference between the two samples can be tentatively attributed to a different NPC curvature, i.e., when the curvature is lower (for NPCSA-GOx-1), more enzyme molecules are assembled on the support surface.

Catalytic performance in the D-glucose oxidation pH influence Since the enzyme activity is dependent on the ionization state of amino acids in the active site, pH plays an important role in maintaining the proper conformation of the enzyme.47 The enzyme immobilization is known to lead to the ionization changes, thus, shifting optimal pH to the more alkaline region or creating a broader рН range, the character of which depends on the enzyme, support, and immobilization types.24 The study of the pH effect on the D-glucose oxidation in the presence of the native enzyme and the biocatalysts developed in this work was performed at 40 °С. Figures 4 and S3, SI, display relative and absolute catalytic activities, respectively, over the pH range. Considering that both graphs show similar trends and relative activities allow for better comparison with published results, here and further we will discuss relative activities. In


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the case of native GOx, the optimal pH is 5. For both immobilized catalysts, however, a comparatively high activity is observed in a broad pH range of 4.8-7.2 with the highest relative activity achieved at pH 7. These data are consistent with the literature data for both native and immobilized enzymes. For native GOx from Aspergillus niger, for example, the highest activity was observed at рН 5-6.11, 47, 64-65 In the case of encapsulated/immobilized GOx, the optimum pH range was determined to be 4-664 or 6-7.66 In our preceding paper for GOx immobilized on alumina and silica, the optimal pH was found to be 7,67 which is consistent with the current findings. These data indicate that the immobilized catalysts developed here can be successfully used in a neutral medium with the relative activity being 17-24% higher than that of native GOx.

b

GOx NPC-SA-GOx-1 NPC-SA-GOx-2

a 80

60

40

20

GOx NPC-SA-GOx-1 NPC-SA-GOx-2

100

Relative activity, %

100


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


90 80 70 60 50 40

4

5

6

7

8

25

30

35

40

45

50

55

60

65

70

Temperature, oC

pH

Figure 4. pH (a) and temperature (b) influence on the D-glucose oxidation. Temperature influence To study the temperature influence on the D-glucose oxidation, the experiments were performed in the temperature range of 30-65 °С at pH 5 for better comparison with native GOx. The dependence of the biocatalyst relative activity on the temperature is presented in Figure 4b. The native enzyme shows the maximum activity at 40 °C, followed by a sharp decrease in the product yield, indicating a loss of the enzyme activity most likely due to denaturation at higher



temperatures.65 In the case of immobilized GOx (for both NPC-SA-GOx-1 and NPC-SA-GOx2), we observed the lower D-gluconic acid yield at 30-40 °C as compared to that of native GOx. This is consistent with prior findings and due to some loss of mobility and reactivity of GOx after immobilization.53 On the other hand, the further temperature increase to 50 °C resulted in much higher product yields (by 14-21%) with the immobilized catalysts than that with the native enzyme. At 65 °C (the highest temperature normally used for enzyme catalyst testing) the native enzyme loses about 60% of the catalytic activity, while the activities of the immobilized GOx catalysts decrease by only 10-15%. This enzyme stabilization is consistent with the data obtained by other authors for GOx immobilized on [email protected] Based on the above studies of the pH and temperature influence, the optimal conditions were found to be pH of 7 and temperature of 50 °C. In these conditions the relative activities of 87 and 94% were achieved for NPC-SAGOx-1 and NPC-SA-GOx-2, respectively. It is noteworthy that in all experiments the relative activity is equal to the conversion, while the selectivity to D-gluconic acid in an enzymatic catalytic process in the presence of GOx is always 100%. NPC-SA-GOx-1 showed consistently lower activity (by about 10%) in the whole temperature range compared to that of NPC-SA-GOx-2.(It is noteworthy that an experimental error in the relative activity assessment did not exceed 2.5%.) As was discussed above, NPC-SA-GOx-1 allows for the attachment of 66% of GOx (from the enzyme loading) on NPCs with the surface area of ~1.2 m2/g, while NPC-SA-GOx-2 attaches only 41% on NPCs with the surface area of ~2.7 m2/g. Because more enzyme was attached to NPC-SA-GOx-1, the higher NPC-SA-GOx-2 activity is counterintuitive. We believe several factors need to be considered when discussing the enzyme activity. On the one hand, the enzyme should be sufficiently mobile after immobilization to allow for normal enzyme function, while the contact with the surface should be minimized to


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prevent denaturation.68 These two conditions can be realized using sufficient length linkers, tethering the enzyme to the support. On the other hand, it is very well established that, in cells, enzyme molecules function in a crowded environment and this environment is beneficial for the enzyme activity.60,

68-69

When enzyme molecules are placed in a diluted buffer solution, they

partially denature which diminishes their activity.70 Thus, ideally the well-designed immobilization on the surfaces or enzyme ordering in confined spaces (pores) may increase the catalytic activity of the enzyme beyond that obtained for a native enzyme in a diluted solution.68 In our case, the native GOx activity is not exceeded, but the best result is close to that. The higher activity of NPC-SA-GOx-2 vs. that of NPC-SA-GOx-1 could be explained by a combination of two key factors: (i) medium crowding of tethered GOx molecules on the NPCSA-GOx-2 support surface preventing denaturing and (ii) a minimal loss of the enzyme mobility, when a lower number of GOx molecules is placed on the higher surface area. A less crowded environment of GOx molecules could be expected due to the higher curvature of the smaller NPCs in NPC-SA-GOx-2 which results in the attachment of less GOx molecules on a larger surface area. A similar trend was observed by us when we studied the functionalization of hydrophobic iron oxide NPs with PEGylated (PEG stands for poly(ethylene glycol)) phospholipids containing terminal carboxyl groups.71We demonstrated, that the amount of PEGylated phospholipids and the charge density (evaluated via ξ-potential measurements) is higher on larger NPs (low curvature) compared to those on smaller NPs (high curvature).For NPC-SA-GOx-1, the crowding is higher (more enzyme molecules on the smaller surface area) but this crowding occurs on the hard surface, which could decrease the mobility of the enzyme. We would like to emphasize that a hypothesis of the NPC curvature influence on the catalytic activity is based on three experimental facts: (i) the higher activity of NPC-SA-GOx-2, (ii) the



lower GOx amount on this catalyst and (iii) its higher surface area. It is noteworthy that the NP curvature and the enzyme activity have been discussed in literature, however, the results are notfully consistent. It was reported that the activity of phosphatidylinositol-specific phospholipase adsorbed on phospholipid vesicles increases with decreasing the diameter of vesicles (increasing curvature).72 The opposite trend was observed for cytosolic phospholipase also adsorbed on vesicles.73 For NP based surfaces, it was demonstrated that the enzyme immobilization on a high curvature surface is beneficial to retain enzymatic activity by preventing denaturing and minimizing changes in the enzyme tertiary structure.74-76Alternatively, in ref. 77 the activity of GOx on the gold flat surface was higher than that on NPs.The lack of consensus regarding the surface curvature influence on the enzyme catalytic activity is probably due to multiple factors influencing the activity, thus making it is difficult to fully decouple the curvature factor. As for catalytic activity of NPC-SA-GOx-2, to the best of our knowledge, the relative activity of 94% is one the highest activities reported to date for immobilized GOx catalysts.78-79 Biocatalyst stability in repeated experiments In order to realize an advantage of magnetically recoverable catalysts and to evaluate the catalyst stability in the repeated use,54-55,

80

ten successive experiments of the D-glucose

oxidation were carried out in the optimal conditions discussed above. After each reaction, the catalyst was magnetically separated with a rare earth magnet and was used again in a consecutive reaction. It is worth noting that the magnetic separation occurs within 30-40 sec. Figure 5 shows that NPC-SA-GOx-1 loses only 7% of its activity after ten catalytic cycles, while NPC-SA-GOx2 loses 10%. The stability of the catalysts discussed in this work exceeds that of GOx immobilized on Fe3O4@C-silica, where 20% of the activity loss occurred after seven catalytic cycles.55 We hypothesize that the higher stability of our catalysts is due to the formation of the


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assembled GOx layer on the support surface which prevents/slows denaturing, while the GOx molecules surrounded by silica, could be isolated and prone to denaturing.55 NPC-SA-GOx-1 NPC-SA-GOx-2

100


1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


80 60 40 20 0 1

2

3

4

5

6

7

8

9

10

Number of cycles

Figure 5. The biocatalyst stability in ten cycles. It is worth noting that the comparatively minor loss of the activity is most likely due to the changes in the GOx structure, rather than the enzyme loss. If the cause would be the enzyme cleavage from the support, some catalytic activity would be observed when the supernatant after the catalyst removal was returned to the catalytic reaction. However, no such activity was detected.

Conclusion We developed novel magnetically recoverable enzymatic catalysts by tethering GOx via a GA linker to the NPCs coated with silica. The NPCs consist of magnetite NPs of about 42-45 nm in diameter, whose close proximity in the NPCs allows for a strong magnetic response and very fast magnetic separation. Depending on the NPC diameter, the magnetic support accommodates different amounts of GOx. For NPC-SA-GOx-1 based on larger NPCs, the amount of



immobilized GOx is approximately 33% higher than that for NPC-SA-GOx-2 based on smaller NPCs. Despite a larger fraction of the enzyme, NPC-SA-GOx-1 is less active allowing for 87% of the relative activity vs. 94% for NPC-SA-GOx-2. This phenomenon is explained by a favorable combination of two major factors: (i) sufficient crowding of tethered GOx molecules on the NPC-SA-GOx-2 support surface preventing denaturing and (ii) a minimal loss of the enzyme mobility, when a lower number of GOx molecules is placed on the higher surface area. For NPC-SA-GOx-1, possible GOx overcrowding could lead to a mobility loss, resulting in the lower activity. The high activity and 100% selectivity as well as exceptional stability of the biocatalyst performance for a very “tolerant” pH range make these immobilized enzymes promising for practical applications.

ASSOCIATED CONTENT Supporting Information. Hysteresis curves, liquid nitrogen adsorption-desorption isotherms, and pore size distributions. This material is available free of charge via the Internet at http://pubs.acs.org. AUTHOR INFORMATION Corresponding Author * [email protected]; [email protected] Author Contributions The manuscript was written through contributions of all authors. ACKNOWLEDGMENT


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The research leading to these results has received funding from the Russian Science Foundation (project 17-19-01408). V.M. and A.S. thank the Russian Foundation for Basic Research (grant 18-08-00468).We also thank the Indiana University Nanoscale Characterization Facility for access to the instrumentation as well as NSF grant #CHE-1048613 which funded the Empyrean from PANalytical.

REFERENCES 1.

Pal, P.; Kumar, R.; Banerjee, S., Manufacture of Gluconic Acid: A Reviewtowards

Process Intensification for Green Production. Chem. Eng. Process. 2016, 104, 160-171. DOI: 10.1016/j.cep.2016.03.009. 2.

Anastassiadis, S. G.; Morgunov, I. G., Gluconic Acid Production. Recent Pat. Biotechnol.

2007, 1 (2), 1-14. DOI: 10.2174/187220807780809472. 3.

Garcia-Morales, N. G.; Garcia-Cerda, L. A.; Puente-Urbina, B. A.; Blanco-Jerez, L. M.;

Antano-Lopez, R.; Castaneda-Zaldivar, F., Electrochemical Glucose Oxidation Using Glassy Carbon Electrodes Modified with Au-Ag Nanoparticles: Influence of Ag Content. J. Nanomater. 2015, 2015, 1-13. DOI: 10.1155/2015/295314. 4.

Da Via, L.; Recchi, C.; Davies, T. E.; Greeves, N.; Lopez-Sanchez, J. A., Back Cover:

Visible-Light-Controlled Oxidation of Glucose Using Titania-Supported Silver Photocatalysts. ChemCatChem 2016, 22, 3475–3483. DOI: 10.1002/cctc.201601429. 5.

Megias-Sayago, C.; Ivanova, S.; Lopez-Cartes, C.; Centeno, M. A.; Odriozola, J. A.,

Gold Catalysts Screening in Base-Free Aerobic Oxidation of Glucose to Gluconic Acid. Catal. Today 2017, 279 (part 2), 148-154. DOI: 10.1016/j.cattod.2016.06.046 6.

Lee, J.; Saha, B.; Vlachos, D. G., Pt Catalysts for Efficient Aerobic Oxidation of Glucose

to Glucaric Acid in Water. . Green Chem. 2016, 18 (13), 3815-3822. DOI: 10.1039/C6GC00460A.



7.

Morawa Eblagon, K.; Pereira, M. F. R.; Figueiredo, J. L., One-Pot Oxidation of

Cellobiose to Gluconic Acid. Unprecedented High Selectivity on Bifunctional Gold Catalysts over Mesoporous Carbon by Integrated Texture and Surface Chemistry Optimization. Appl. Catal. B: Environ. 2016, 184, 381-396. DOI: 10.1016/j.apcatb.2015.10.011. 8.

Xiuhua, L.; Hui, S.; Wei, W.; Shuai, L.; Zaifu, L.; Jiawei, D.; Lisa, X.; Jianshu, L.,

Glucose-Responsive Micelles for Controlled Insulin Release Based on Transformation from Amphiphilic to Double Hydrophilic. J. Nanosci. Nanotechnol. 2016, 16 (6), 5457-5463. DOI: 10.1166/jnn.2016.11655. 9.

Lang, N. J.; Liu, B.; Liu, J., Characterization of Glucose Oxidation by Gold

Nanoparticles Using Nanoceria. J. Colloid Interf Sci. 2014, 428, 78-83. DOI: 10.1016/j.jcis.2014.04.025. 10.

Taketoshi, A.; Takenouchi, S.; Takei, T.; Haruta, M., Reprint of “Synergetic

Combination of an Enzyme and Gold Catalysts for Glucose Oxidation in Neutral Aqueous Solution”. Appl. Catal. B: Environ. 2014, 474, 257-262. DOI: 10.1016/j.apcata.2013.12.012. 11.

Singh, J.; Verma, N., Glucose Oxidase from Aspergillus Niger: Production,

Characterization and Immobilization for Glucose Oxidation. Adv. Appl. Sci. Res. 2013, 4 (3), 250-257. ISSN: 0976-8610. 12.

Wojcieszak, R.; Ferraz, C. P.; Sha, J.; Houda, S.; Rossi, L. M.; Paul, S., Advances in

Base-Free Oxidation of Bio-Based Compounds on Supported Gold Catalysts. Catalysts 2017, 7 (11), 352/1-352/23. DOI: 10.3390/catal7110352. 13.

Biella, S.; Prati, L.; Rossi, M., Selective Oxidation of D-Glucose on Gold Catalyst. J.

Catal. 2002, 206 (2), 242-247. DOI: 10.1006/jcat.2001.3497.


Page 24 of 35

Page 25 of 35 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


14.

Wang, Y.; Van de Vyver, S.; Sharma, K. K.; Roman-Leshkov, Y., Insights into the

Stability of Gold Nanoparticles Supported on Metal Oxides for the Base-Free Oxidation of Glucose to Gluconic Acid. Green Chem. 2014, 16 (2), 719-726. DOI: 10.1039/C3GC41362D. 15.

Miedziak, P. J.; Alshammari, H.; Kondrat, S. A.; Clarke, T. J.; Davies, T. E.; Morad, M.;

Morgan, D. J.; Willock, D. J.; Knight, D. W.; Taylor, S. H.; Hutchings, G. J., Base-Free Glucose Oxidation Using Air with Supported Gold Catalysts. Green Chem. 2014, 16 (6), 3132-3141. DOI: 10.1039/C4GC00087K. 16.

Manuela, D. E.; Gheorghe, M., Temperature Decrease (30–25 °C) Influence on Bi-

Enzymatic Kinetics of D-Glucose Oxidation. J. Mol. Cat. B: Enzym. 2012, 81, 19-24. DOI: 10.1016/j.molcatb.2012.05.001. 17.

Chenhong, W.; Dong, W.; Zhongxi, Z.; Dexin, W.; Jiping, S.; Chul, H., Kim ; Biao, J.;

Zengsheng, H.; Jian, H., Production of Xylonic Acid by Klebsiella Pneumoniae. Appl. Microbiol. Biot. 2016, 100 (23), 10055-10063. DOI: 10.1007/s00253-016-7825-9. 18.

Morthensen, S. T.; Meyer, A. S.; Joergensen, H.; Pinelo, M., Significance of Membrane

Bioreactor Design on the Biocatalytic Performance of Glucose Oxidase and Catalase: Free Vs. Immobilized Enzyme Systems. Biochem. Eng. J. 2017, 117 (Part A), 41-47. DOI: 10.1016/j.bej.2016.09.015. 19.

Huan, Z.; Lulu, S.; Xinlei, M.; Jinping, L.; Dongzhi, W., Enhancement of Cell Growth

and Glycolic Acid Production by Overexpression of Membrane-Bound Alcohol Dehydrogenase in Gluconobacter Oxydans Dsm 2003. J. Biotechnol. 2016, 237, 18-24. DOI: 10.1016/j.jbiotec.2016.09.003.



20.

Caixia, C.; Haibin, C.; Biqiang, C.; Tianwei, T., Genipin Cross-Linked Glucose Oxidase

and Catalase Multi-Enzyme for Gluconic Acid Synthesis. Appl. Biochem. Biotechnol. 2016, 181 (2), 526-535. DOI: 10.1007/s12010-016-2228-z. 21.

Tischer, W.; Wedekind, F., Immobilized Enzymes: Methods and Applications. Top. Curr.

Chem. 1999, 200, 95-126. DOI: 10.1007/3-540-68116-7_4. 22.

Guisan, J. M., Immobilization of Enzymes and Cells. Methods in Biotechnology. .

Humana Press Inc.: Totowa, New Jersey 2006; p 449. DOI: 10.1007/978-1-62703-550-7. 23.

Monier, M.; Ayad, D. M.; Wei, Y.; Sarhan, A. A., Immobilization of Horseradish

Peroxidase on Modified Chitosan Beads. . Int. J. Biol. Macromol. 2010, 46 (3), 324-330. DOI: 10.1016/j.ijbiomac.2009.12.018. 24.

Dwevedi, A., Enzyme Immobilization. Advances in Industry, Agriculture, Medicine, and

the Environment. Springer International Publishing Switzerland: Cham, Switzerland, 2016; p 132. DOI: 10.1007/978-3-319-41418-8. 25.

Brena, B., M.; Batista-Viera, F., Immobilization of Enzymes: A Literature Survey. In

Immobilization of Enzymes and Cells, Guisan, J., M. , Ed. Humana Press Inc.: Totowa, New Jersey, 2006; Chapter 2, pp 15-30. DOI: 10.1007/978-1-62703-550-7_2. 26.

Datta, S.; Christena, L. R.; Rajaram, Y. R. S., Enzyme Immobilization: An Overview on

Techniques and Support Materials. Biotech 2013, 3, 1-9. DOI: 10.1007/s13205-012-0071-7. 27.

Mohamad, N. R.; Marzuki, N. H.; Buang, N. A.; Huyop, F.; Wahab, R. A., Review;

Agriculture and Environmental Biotechnology. An Overview of Technologies for Immobilization of Enzymes and Surface Analysis Techniques for Immobilized Enzymes. Biotechnol. Biotec. Eq. 2015, 29 (2), 205-220. DOI: 10.1080/13102818.2015.1008192.


Page 26 of 35

Page 27 of 35 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


28.

Ahmad, R.; Sardar, M., Enzyme Immobilization: An Overview on Nanoparticles as

Immobilization Matrix. Biochem. Anal. Biochem. 2015, 4 (2), 1-8. DOI: 10.4172/21611009.1000178. 29.

Zhu, Y.; Peng, S. C.; Emi, A.; Su, Z.; Monalisa; Kemp, R. A., Supported Ultra Small

Palladium on Magnetic Nanoparticles Used as Catalysts for Suzuki Cross-Coupling and Heck Reactions. Adv. Synth. Catal. 2007, 349 (11-12), 1917-1922. DOI: 10.1002/adsc.200700021 30.

Polshettiwar, V.; Luque, R.; Fihri, A.; Zhu, H.; Bouhrara, M.; Basset, J.-M., Magnetically

Recoverable Nanocatalysts. Chem. Rev. 2011, 111 (5), 3036-3075. DOI: 10.1021/cr100230z. 31.

Rossi, L. M.; Silva, F. P.; Vono, L. L. R.; Kiyohara, P. K.; Duarte, E. L.; Itri, R.; Landers,

R.; Machado, G., Superparamagnetic Nanoparticle-Supported Palladium: A Highly Stable Magnetically Recoverable and Reusable Catalyst for Hydrogenation Reactions. Green Chem. 2007, 9 (4), 379-385. DOI: 10.1039/b612980c. 32.

Saha, A.; Leazer, J.; Varma, R. S., O-Allylation of Phenols with Allylic Acetates in

Aqueous Media Using a Magnetically Separable Catalytic System. Green Chem. 2012, 14 (1), 67-71. DOI: 10.1039/C1GC16174A. 33.

Vaddula, B. R.; Saha, A.; Leazer, J.; Varma, R. S., A Simple and Facile Heck-Type

Arylation of Alkenes with Diaryliodonium Salts Using Magnetically Recoverable Pd-Catalyst. Green Chem. 2012, 14 (18), 2133-2136. DOI: 10.1039/c2gc35673b. 34.

Wang, D.; Astruc, D., Fast-Growing Field of Magnetically Recyclable Nanocatalysts.

Chem. Rev. 2014, 114 (14), 6949-6985. DOI: 10.1021/cr500134h. 35.

Lu, A.-H.; Salabas, E. L.; Schueth, F., Magnetic Nanoparticles: Synthesis, Protection,

Functionalization, and Application. Angew. Chem. Int. Ed. 2007, 46 (8), 1222-1244 DOI: 10.1002/anie.200602866.



36.

Bazgir, A.; Hosseini, G.; Ghahremanzadeh, R., Copper Ferrite Nanoparticles: An

Efficient and Reusable Nanocatalyst for a Green One-Pot, Three-Component Synthesis of Spirooxindoles in Water. ACS Comb. Sci. 2013, 15 (10), 530-534. DOI: 10.1021/co400057h. 37.

Kundu, D.; Chatterjee, T.; Ranu, B. C., Magnetically Separable Cufe2o4 Nanoparticles

Catalyzed Ligand-Free C-S Coupling in Water: Access to (E)- and (Z)-Styrenyl-, Heteroaryl and Sterically Hindered Aryl Sulfides. Adv. Synth. Catal. 2013, 355 (11-12), 2285-2296. DOI: 10.1002/adsc.201300261. 38.

Kuchkina, N. V.; Morgan, D. G.; Kostopoulou, A.; Lappas, A.; Brintakis, K.; Boris, B.

S.; Yuzik-Klimova, E. Y.; Stein, B. D.; Svergun, D. I.; Spilotros, A.; Sulman, M. G.; Nikoshvili, L. Z.; Sulman, E. M.; Shifrina, Z. B.; Bronstein, L. M., Hydrophobic Periphery Tails of Polyphenylenepyridyl Dendrons Control Nanoparticle Formation and Catalytic Properties. Chem. Mater. 2014, 26 (19), 5654-5663. DOI: 10.1021/cm502336n. 39.

Yuzik-Klimova, E. Y.; Kuchkina, N. V.; Sorokina, S. A.; Boris, B.; Morgan, D. G.;

Nikoshvili, L. Z.; Lyubimova, N. A.; Matveeva, V. G.; Sulman, E. M.; Stein, B. D.; Svergun, D. I.; Spilotros, A.; Kostopoulou, A.; Lappas, A.; Shifrina, Z. B.; Bronstein, L. M., Magnetically Recoverable Catalysts Based on Polyphenylenepyridyl Dendrons and Dendrimers. RSC Adv. 2014, 4 (44), 23271 - 23280. DOI: 10.1039/c4ra00878b. 40.

Mann, J.; Doluda, V. Y.; Leonard, C.; Losovyj, Y. B.; Morgan, D. G.; Bukalov, S. S.;

Shifrina, Z.; Stein, B. D.; Cherkasov, N.; Rebrov, E. V.; Harms, Z. D.; Pink, M.; Sulman, E. M.; Bronstein, L. M., Metal Oxide-Zeolite Composites in Transformation of Methanol to Hydrocarbons: Do Iron Oxide and Nickel Oxide Matter? RSC Adv. 2016, 6 (79), 75166-75177. DOI: 10.1039/C6RA19471K.


Page 28 of 35

Page 29 of 35 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


41.

Manaenkov, O. V.; Mann, J. J.; Kislitza, O. V.; Losovyj, Y.; Stein, B. D.; Morgan, D. G.;

Pink, M.; Lependina, O. L.; Shifrina, Z. B.; Matveeva, V. G.; M.;, S. E.; Bronstein, L. M., RuContaining Magnetically Recoverable Catalysts: A Sustainable Pathway from Cellulose to Ethylene and Propylene Glycols. ACS Appl. Mater. & Interfaces 2016, 8 (33), 21285-21293. DOI: 10.1021/acsami.6b05096. 42.

Akhond, M.; Pashangeh, K.; Karbalaei-Heidari, H. R.; Absalan, G., Efficient

Immobilization of Porcine Pancreatic Α-Amylase on Amino-Functionalized Magnetite Nanoparticles:Characterization and Stability Evaluation of the Immobilized Enzyme. Appl. Biochem. Biotechnol. 2016, 180, 954–968. DOI: 10.1007/s12010-016-2145-1. 43.

Deng, X.; Cao, S.; Li, N.; Wu, H.; Smith, T. J.; Zong, M.; Lou, W., A Magnetic

Biocatalyst Based on Mussel‐Inspired Polydopamine and Its Acylation of Dihydromyricetin. Chinese J. Catal. 2016, 37, 584–595. DOI: 10.1016/S1872-2067(15)61045-2. 44.

Ma, J.; Wu, Y.; Zeng, Y.; Li, Y.; Wu, D., Thermo-Stable Hollow Magnetic

Microspheres: Preparation, Characterization and Recyclable Catalytic Applications. J. Mater. Chem: A 2015, 3, 16762–16773. DOI: 10.1039/c5ta03553h. 45.

Wang, J.; Liu, Z.; Zhou, Z., Improving Pullulanase Catalysis Via Reversible

Immobilization on Modified Fe3o4@Polydopamine Nanoparticles. Appl. Biochem. Biotechnol. 2017, 182 (4), 1467–1477. DOI: 10.1007/s12010-017-2411-x 46.

Zheng, M.; Zhang, S., Immobilization of Glycerol Dehydrogenase on Magnetic Silica

Nanoparticles for Conversion of Glycerol to Value-Added 1,3-Dihydroxyacetone. Biocatal. Biotransfor. 2011, 29 (6), 278-287. DOI: 10.3109/10242422.2011.631212. 47.

Bankar, S., B; Bule, M., V; Singhal, R., S; Ananthanarayan, L., Glucose Oxidase - an

Overview. Biotechnol. Adv. 2009, 27, 489-501. DOI: 10.1016/j.biotechadv.2009.04.003.



48.

Kumar, R.; Leech, D., Coupling of Amine-Containing Osmium Complexes and Glucose

Oxidase with Carboxylic Acid Polymer and Carbon Nanotube Matrix to Provide Enzyme Electrodes for Glucose Oxidation J. Electrochem. Soc. 2014, 161 (13), 3005-3010. DOI: 10.1149/2.0021413jes. 49.

MacAodha, D.; Conghaile, P., Ó.; Egan, B.; Kavanagh, P.; Sygmund, C.; Ludwig, R.;

Leech, D., Comparison of Glucose Oxidation by Crosslinked Redox Polymer Enzyme Electrodes Containing Carbon Nanotubes and a Range of Glucose Oxidising Enzymes. Electroanal. 2013, 25 (1), 94-100. DOI: 10.1002/elan.201200536. 50.

Yang, Y. M.; Wang, J. W.; Tan, R. X., Immobilization of Glucose Oxidase on Chitosan–

Sio2 Gel. Enzyme Microb. Tech. 2004, 34, 126–131. DOI: 10.1016/j.enzmictec.2003.09.007. 51.

Gellf, G.; Boudrant, J., Enzymes Immobilized on a Magnetic Support. Preliminary Study

of a Fluidized Bed Enzyme Reactor. Biochim. Biophys. Acta, Enzymol. 1974, 334 (2), 467-70. DOI: 10.1016/0005-2744(74)90194-6. 52.

Sada, E.; Katoh, S.; Shiozawa, M.; Fukui, T., Performance of Fluidized-Bed Reactors

Utilizing Magnetic Fields. Biotechnol. Bioeng. 1981, 23 (11), 2561-7. DOI: 10.1002/bit.260231113. 53.

Pieters, B., R.; Bardeletti, G., Enzyme Immobilization on a Low-Cost Magnetic Support:

Kinetic Studies on Immobilized and Coimmobilized Glucose Oxidase and Glucoamylase. Enzyme Microb. Technol 1992, 14, 361-370. DOI: 10.1016/0141-0229(92)90004-8. 54.

Chen, S.; Wen, L.; Svec, F.; Tan, T.; Lv, Y., Magnetic Metal-Organic Frameworks as

Scaffolds for Spatial Co-Location and Positional Assembly of Multi-Enzyme Systems Enabling Enhanced Cascade Biocatalysis. RSC Adv. 2017, 7 (34), 21205-21213. DOI: 10.1039/C7RA02291C.


Page 30 of 35

Page 31 of 35 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


55.

Yang, Y.; Zhu, G.; Wang, G.; Li, Y.; Tang, R., Robust Glucose Oxidase with a

Fe3o4@C-Silica Nanohybrid Structure. J. Mater. Chem: B. 2016, 4, 4726-4731. DOI: 10.1039/C6TB01355D. 56.

Cheng, C.; Xu, F.; Gu, H., Facile Synthesis and Morphology Evolution of Magnetic Iron

Oxide Nanoparticles in Different Polyol Processes. New J. Chem. 2011, 35 (5), 1072-1079. DOI: 10.1039/c0nj00986e. 57.

Podolean, I.; Kuncser, V.; Gheorghe, N.; Macovei, D.; Parvulescu, V. I.; Coman, S. M.,

Ru-Based Magnetic Nanoparticles (Mnp) for Succinic Acid Synthesis from Levulinic Acid. Green Chem. 2013, 15 (11), 3077-3082. DOI: 10.1039/c3gc41120f. 58.

Tomotani, E. J.; Vitolo, M., Immobilized Glucose Oxidase as a Catalyst to the

Conversion of Glucose into Gluconic Acid Using a Membrane Reactor. Enzyme Microb. Tech. 2007, 40, 1020-1025. DOI: 10.1016/j.enzmictec.2006.07.039. 59.

Nguyen, L. T.; Yang, K.-L., Combined Cross-Linked Enzyme Aggregates of Horseradish

Peroxidaseand Glucose Oxidase for Catalyzing Cascade Chemical Reactions. Enzyme Microb. Tech. 2017, 100, 52-59. DOI: 10.1016/j.enzmictec.2017.02.007. 60.

Acosta, L. C.; Perez Goncalves, G. M.; Pielak, G. J.; Gorensek-Benitez, A. H., Large

Cosolutes, Small Cosolutes, and Dihydrofolate Reductase Activity. Protein Sci. 2017, 26, 2417—2425. DOI: 10.1002/pro.3316. 61.

Chairam, S.; Somsook, E., Starch Vermicelli Template for Synthesis of Magnetic Iron

Oxide Nanoclusters. J. Magn. Magn. Mater. 2008, 320 (15), 2039-2043. DOI: 10.1016/j.jmmm.2008.02.168.



62.

Goya, G. F.; Berquo, T. S.; Fonseca, F. C.; Morales, M. P., Static and Dynamic Magnetic

Properties of Spherical Magnetite Nanoparticles. J. Appl. Phys. 2003, 94 (5), 3520-3528. DOI: 10.1063/1.1599959. 63.

Sing, K. S. W.; Everett, D. H.; Haul, R. A. W.; Moscou, L.; Pierotti, R. A.; Rouquerol, J.;

Siemieniewska, T., Reporting Physisorption Data for Gas/Solid Systems with Special Reference to the Determination of Surface Area and Porosity (Recommendations 1984). Pure Appl. Chem. 1985, 57 (4), 603-19. DOI: 10.1351/pac198557040603. 64.

Vikartovska, A.; Bucko, M.; Mislovicova, D.; Patoprsty, V.; Lacık, I.; Gemeiner, P.,

Improvement of the Stability of Glucose Oxidase Via Encapsulation in Sodium Alginate– Cellulose Sulfate–Poly(Methylene-Co-Guanidine) Capsules. Enzyme Microb. Tech. 2007, 41, 748–755. DOI: 10.1016/j.enzmictec.2007.06.010. 65.

Wong, C. M.; Wong, K. H.; Chen, X. D., Glucose Oxidase: Natural Occurrence,

Function, Properties and Industrial Applications. Appl. Microbiol. Biotechnol. 2008, 78, 927938. DOI: 10.1007/s00253-008-1407-4. 66.

Wang, F.; Ma, Y.; Liu, Y.-H.; Zhang, X.; Zhang, F.; Linhardt, R. J., Improved Octyl

Glucoside Synthesis Using Immobilized Β-Glucosidase on Pa-M with Reduced Glucose Surplus Inhibition. Biocatal. Biotransform. 2017, 35 (5), 349-362. DOI: 10.1080/10242422.2017.1345886. 67.

Golikova, E. P.; Lakina, N. V.; Grebennikova, O. V.; Matveeva, V. G.; Sulman, E. M., A

Study of Biocatalysts Based on Glucose Oxidase. Faraday Discuss 2017, 202, 303-314. DOI: 10.1039/C7FD00042A.


Page 32 of 35

Page 33 of 35 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


68.

Kao, K.-C.; Lin, T.-S.; Mou, C.-Y., Enhanced Activity and Stability of Lysozyme by

Immobilization in the Matching Nanochannels of Mesoporous Silica Nanoparticles. J. Phys. Chem. C 2014, 118, 6734−6743. DOI: 10.1021/jp4112684. 69.

Lei, C.; Shin, Y.; Magnuson, J. K.; Fryxell, G.; Lasure, L. L.; Elliott, D. C.; Liu, J.;

Ackerman, E. J., Characterization of Functionalized Nanoporous Supports for Protein Confinement. Nanotechnology 2006, 17, 5531–5538. DOI: 10.1088/0957-4484/17/22/001. 70.

Kunkel, J.; Asuri, P., Function, Structure, and Stability of Enzymes Confined in Agarose

Gels. PLoS One 2014, 9 (1), e86785/1-e86785/6, 6 pp. DOI: 10.1371/journal.pone.0086785. 71.

Huang, X.; Bronstein, L. M.; Retrum, J. R.; Dufort, C.; Tsvetkova, I.; Aniagyei, S.; Stein,

B.; Stucky, G.; McKenna, B.; Remmes, N.; Baxter, B.; Kao, C. C.; Dragnea, B., Self-Assembled Virus-Like Particles with Magnetic Cores. Nano Lett. 2007, 7 (8), 2407-2416. DOI: 10.1021/nl071083l. 72.

Ahyayauch, H.; Villar, A. V.; Alonso, A.; Goni, F. M., Modulation of Pi-Specific

Phospholipase C by Membrane Curvature and Molecular Order. Biochemistry 2005, 44 (34), 11592-11600. DOI: 10.1021/bi050715k. 73.

Burke, J. R.; Witmer, M. R.; Tredup, J. A., The Size and Curvature of Anionic Covesicle

Substrate Affects the Catalytic Action of Cytosolic Phospholipase A2. Arch. Biochem. Biophys. 1999, 365 (2), 239-247. DOI: 10.1006/abbi.1999.1151. 74.

Gagner, J. E.; Lopez, M. D.; Dordick, J. S.; Siegel, R. W., Effect of Gold Nanoparticle

Morphology on Adsorbed Protein Structure and Function. Biomaterials 2011, 32 (29), 72417252. DOI: 10.1016/j.biomaterials.2011.05.091.



75.

Chen, L.; Xie, H.; Li, J., Electrochemical Glucose Biosensor Based on Silver

Nanoparticles/Multiwalled Carbon Nanotubes Modified Electrode. J. Solid State Electrochem. 2012, 16 (10), 3323-3329. DOI: 10.1007/s10008-012-1773-9. 76.

Bergman, J.; Wang, Y.; Wigstroem, J.; Cans, A.-S., Counting the Number of Enzymes

Immobilized onto a Nanoparticle-Coated Electrode. Anal. Bioanal. Chem. 2018, 410 (6), 17751783. DOI: 10.1007/s00216-017-0829-1. 77.

Seehuber, A.; Dahint, R., Conformation and Activity of Glucose Oxidase on

Homogeneously Coated and Nanostructured Surfaces. J. Phys. Chem. B 2013, 117 (23), 69806989. DOI: 10.1021/jp401906h. 78.

Yang, Y.; Zhang, R.; Zhou, B.; Song, J.; Su, P.; Yang, Y., High Activity and Convenient

Ratio Control: DNA-Directed Coimmobilization of Multiple Enzymes on Multifunctionalized Magnetic Nanoparticles. ACS Appl. Mater. Interfaces 2017, 9 (42), 37254-37263. DOI: 10.1021/acsami.7b08553. 79.

Li, Z.; Ding, Y.; Li, S.; Jiang, Y.; Liu, Z.; Ge, J., Highly Active, Stable and Self-

Antimicrobial Enzyme Catalysts Prepared by Biomimetic Mineralization of Copper Hydroxysulfate. Nanoscale 2016, 8 (40), 17440-17445. DOI: 10.1039/C6NR06115J. 80.

Zhao, B.; Zhou, L.; Ma, L.; He, Y.; Gao, J.; Li, D.; Jiang, Y., Co-Immobilization of

Glucose Oxidase and Catalase in Silica Inverse Opals for Glucose Removal from Commercial Isomaltooligosaccharide. Int. J. Biol. Macromol. 2018, 107 (Part_B), 2034-2043. DOI: 10.1016/j.ijbiomac.2017.10.0744.


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SYNOPSIS. Sustainable enzymatic D-glucose oxidation with high gluconic acid yields has been developed with magnetically recoverable biocatalysts composed of glucose oxidase immobilized on magnetic support. TABLE OF CONTENTS (TOC) GRAPHIC


Insights into sustainable glucose oxidation using magnetically

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