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Förster resonance energy transfer study of the improved biocatalytic conversion of COto formaldehyde by co-immobilization of enzymes in siliceous mesostructured cellular foams 2

Pegah S. Nabavi Zadeh, Milene Zezzi do Valle Gomes, Björn Åkerman, and Anders E. C. Palmqvist ACS Catal., Just Accepted Manuscript • DOI: 10.1021/acscatal.8b01806 • Publication Date (Web): 25 Jun 2018 Downloaded from http://pubs.acs.org on June 25, 2018

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ACS Catalysis

Förster resonance energy transfer study of the improved biocatalytic conversion of CO2 to formaldehyde by coimmobilization of enzymes in siliceous mesostructured cellular foams Pegah S. Nabavi Zadeh#a*, Milene Zezzi do Valle Gomes#b, Björn Åkermana and Anders E.C. Palmqvistb a

Chalmers University of Technology, Department of Chemistry and Chemical Engineering,

Physical Chemistry, SE-41296 Gothenburg, Sweden b

Chalmers University of Technology, Department of Chemistry and Chemical Engineering,

Applied Chemistry, SE-41296 Gothenburg, Sweden #

Contributed equally in this work

*Corresponding author: [email protected]

Abstract By combining the two enzymes, formate dehydrogenase (FateDH) and formaldehyde dehydrogenase (FaldDH), it is possible to drive the thermodynamically unfavourable conversion of CO2 to formaldehyde. For this purpose, the enzymes were co-immobilized in siliceous mesostructured cellular foams (MCFs). A high degree of adsorption of both enzymes was achieved by co-immobilizing the enzymes sequentially, i.e. first FateDH and then FaldDH. The highest conversion rate was obtained with an enzyme mass ratio of 1:15 (FateDH: FaldDH). Using MCF functionalized with mercaptopropyl groups (MCF-MP), the activity increased about 4 times compared to the enzymes free in solution. To probe the distance between the two enzymes they were separately labelled with either Cy3 or Cy5 dyes and studied with Förster resonance energy transfer (FRET). An increased energy transfer was observed when the enzymes were co-immobilized in MCF-MP, suggesting that the two enzymes are in close proximity resulting in higher conversion of CO2 to formaldehyde. Keywords: CO2 reduction, mesoporous silica, biocatalysis, Förster resonance energy transfer, carbocyanine dyes, labeled protein

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1. Introduction The environmental impact caused by the high emissions of CO2 is a global concern. Despite the development of new technologies to limit the generation of this greenhouse gas, the demands for energy and the use of fossil fuels continue to increase.1-3 Therefore, it is considered highly important to develop energy efficient technologies to convert CO2 to useful chemicals or fuels and pave the way towards a sustainable society.1, 4 One interesting possibility is to convert CO2 into methanol, which can be used as transportation fuel, as raw material for chemical synthesis, and as an energy storage medium.1,

5

However, the

conversion of CO2 to methanol is a thermodynamically unfavorable reaction, which consumes energy and requires a catalyst to proceed with a relevant rate.2, 6-7 It has been shown that with an appropriate energy supply, a multi-enzymatic system can be used to catalyze the reduction of CO2 to methanol under mild conditions.7-11 This system involves three steps (See Scheme 1).

Scheme 1. Enzymatic cascade conversion of carbon dioxide (CO2) to methanol (CH3OH).

In the first reaction, formate dehydrogenase (FateDH) converts CO2 to formic acid. The formaldehyde dehydrogenase (FaldDH) subsequently reduces formic acid to formaldehyde and in the final step, formaldehyde is converted to methanol by alcohol dehydrogenase (ADH).8 All three enzymes use the cofactor nicotinamide adenine dinucleotide (NADH) as electron donor.8 For the complete system to be energetically sustainable, NADH needs to be regenerated from NAD+, which can be done using solar light and facilitated through photocatalytic reduction of NAD+.12-14 An energetically challenging step of Scheme 1 is the first reaction, the reduction of CO2 to formate (at the pH 6.5 of the present work). To increase the conversion rate of the multi-enzymatic system this is, therefore, a step that requires attention. The

conversion

of

CO2

to

formate

is

thermodynamically

unfavorable

(∆rG’o = 20.6 kJ·mol-1),15 which means that for this reaction to proceed, it is necessary to use high concentrations of CO2 and NADH and to avoid the back reaction converting formate to 2 ACS Paragon Plus Environment

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CO2.10 FateDH very effectively performs this back reaction, with a rate that is about 30 times higher than for the forward reduction of CO2 (at pH 6.2 and 10 oC).16 One strategy to avoid the back reaction is to remove the formate as soon as it is formed so it is not available for the reverse reaction. This can, for instance, be achieved by adding the FaldDH enzyme to the same reactor system, to catalyze the reduction of formate to formaldehyde. The kinetic parameters of FaldDH for the reduction reaction (HCOOHHCHO) has not been determined yet, mainly due to the difficulty to measure the reaction velocities at different formic acid concentrations keeping the pH constant. For the oxidation reaction (HCHOHCOOH) Vmax is equal to 0.01 mM·min -1 (at pH 7.0 and 23 oC).11 FateDH and FaldDH have similar optimum pH and temperature, and also their activities towards the reduction pathways are increased under 5 bars pressure.10 The fact that both enzymes work most efficiently under similar conditions is essential for the cascade reaction to be effective.17 Those conditions, however, are not favorable for the third enzyme in the system, ADH. It has been shown that under such conditions the activity of ADH reduces significantly.18 Therefore, it is of interest to optimize the initial part of the cascade reaction involving only FateDH and FaldDH, as a way to improve the complete reaction system for converting CO2 to methanol. However, cascade reactions with enzymes free in solution are usually not very efficient since the enzymes are not in sufficiently close proximity.17,

19

Therefore, the co-

immobilization of the enzymes in a host material appears as an interesting approach, since confined enzymes tend to be closer and substrate channeling between two enzymes becomes possible.19-21 In such an arrangement, the product of the reaction performed by the first enzyme is directly converted by the second enzyme without equilibration with the bulk phase.20,

22

Theoretical work in free solution23 supports that short distance and suitable

orientation of the two enzymes in a cascade increases the total reaction rate. DNA-based scaffolds have been used to study confinement effects on the total activity of coimmobilized enzymes.24-27 One important observation is that the high charge density of the host material might alter the local pH, with effects on the enzyme activity that adds to or even masks distance effects. This notion is supported by a theoretical study of colocalization of two enzymes in a DNA-mimicking confinement but where electrostatic effects are absent by design.28 In this confined case similar effects of distance and orientation on the cascade efficiency are observed as in the theoretical study in free solution.23 We have shown previously 29 that the pH inside the pores of MPS is similar to the external solution surrounding the particles, so silica-based hosts is an interesting 3 ACS Paragon Plus Environment

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complement to DNA-based scaffolds to study co-confinement effects in an environment where the local pH is not altered by the host material. In addition, we have demonstrated that even though the rotation of two types of proteins (lipase and BSA) is retarded by confinement in the pores of MPS they are not rotationally immobile.30 This may be important since theoretical work23 indicates that the possibility for the enzymes to rotate may be important in order to optimize the cascade efficiency. In our previous work, we studied how the activity of FaldDH can be improved by the immobilization of this enzyme in tailored siliceous mesostructured cellular foams (MCF).31 MCF is a type of mesoporous silica that contains large pores (20-40 nm) connected by smaller windows (10-20 nm).32 By tuning the pore size of the material to about 33 nm and by functionalizing the MCF surface with mercaptopropyl groups, the activity of FaldDH was increased and a low enzyme leakage was observed during the reaction.31 In the present study, an MCF with similar physical properties as used previously for the immobilization of FaldDH, was synthesized and used to co-immobilize FateDH and FaldDH through physical adsorption. This method of immobilization was chosen since it has several advantages. There is no need of special conditions such as organic solvents and extreme pH values, and secondly compared to covalent attachment, it is less likely that the enzymes undergo conformational changes because only weak intermolecular interactions are involved.33-37 However, the interaction between the enzymes and the support material is not controlled, thus the enzymes may be oriented and distributed inside the pores randomly. 33-37 In the co-immobilization process the two enzymes may also compete for entering the pores or adsorbing on outer and inner surface of mesoporous silica, influencing the total amount of each enzyme that can be immobilized. Therefore, in order to optimize the co-immobilization of the FateDH and FaldDH aiming for higher reaction rates, two parameters used in the immobilization were varied in this study, i.e. enzyme concentrations and the order in which the enzymes were added and mixed with the host material. To evaluate the efficiency of the co-immobilization process and the usefulness of coimmobilization on the CO2 conversion efficiency, the two enzymes were labeled with carbocyanine dyes; the FateDH with Sulfo-cyanine3 NHS ester (Cy3) and FaldDH with Sulfo-cyanine5 NHS ester (Cy5). The labeled enzymes were used i) to monitor the degree of immobilization of the two enzymes independently and ii) to determine the average distance between the two enzymes using Förster resonance energy transfer (FRET). More specifically, we investigate the catalytic activity of co-immobilized FateDH and FaldDH in


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tailored MCF and use FRET to investigate the correlation between the transfer efficiency (E) between the two enzymes and the CO2 conversion rates obtained.

2. Experimental section 2.1 Materials Pluronic™ P123 (EO20PO70EO20, Mw=5800), 1,3,5 trimethylbenzene (TMB, 98%), tetraethylorthosilicate (TEOS, ≥98%), hydrochloric acid (HCl, 37 wt%), ammonium fluoride (NH4F, ≥99,9%), 3-mercaptopropyltrimethoxysilane (MPTS, 95%), toluene (anhydrous, 99.8%), potassium phosphate monobasic (KH2PO4, ≥98%), potassium phosphate dibasic (K2HPO4, ≥98%), ammonium acetate (≥98%), acetyl acetone (≥99%), acetic acid (100%), β-nicotinamide adenine dinucleotide reduced disodium salt hydrate (NADH, 98%), Potassium bicarbonate (KHCO3, ≥ 99,5%), formate dehydrogenase from Candida boidinii (5.0-15.0 units/mg protein; lyophilized powder), formaldehyde dehydrogenase from

Pseudomonas sp. (1.0-6.0 units/mg solid; lyophilized powder) were all purchased from Sigma-Aldrich. Sulfo-cyanine3 NHS ester (Cy3) and Sulfo-cyanine5 NHS ester (Cy5) as fluorescent probes were purchased from Lumiprobe, Life Science Solutions. The spectral properties and the structures of the probes can be seen in Table 1 and Figure 1, respectively, as provided by the supplier. Table 1. Spectral Properties of the Carbocyanine Dyes Fluorescent Exb Emc Φd εe a -1 dye (nm) (nm) (M cm-1) Cy3 550 565 0.1 162000 Cy5 650 665 0.2 271000 a

CF280f

DOLh

0.06 0.13

3.1 5.8

Cy3: Sulfo-cyanine3 NHS ester, Cy5: sulfo-cyanine5 NHS ester. Excitation maximum wavelength; obtained experimentally in the present work. c Emission maximum wavelength; obtained experimentally in the present work. d Fluorescence quantum yield; provided by supplier. e Extinction (molar absorption ) coefficient at absorption maximum; provided by supplier. f Correction factor at 280 nm wavelength for calculating degree of labelling; provided by supplier. h Average of degree of labelling, the number of dyes per enzyme. The uncertainty ±0.2 is calculated from the variation between 5 to 6 independent experiments; used the Eq.S1 in Supporting information. b


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Figure 1. Structure of sulfo-cyanine3 NHS ester (Cy3) (left) and sulfo-cyanine5 NHS ester (Cy5) (right).

2.2 Synthesis and characterization of siliceous mesostructured cellular foams. Siliceous mesostructured cellular foams (MCF) were synthesized following the method developed by Schmidt-Winkel et al.

32

Briefly, 2.9 g pluronic™ P123 was dissolved in

100 ml of 1.6 M HCl at room temperature. After that, 6.3 ml of TMB and 0.031g NH4F were added under vigorous stirring and the temperature was raised to 38 ºC. After 1 h, 6.4 ml of TEOS was added and its hydrolysis and condensation was carried out for 20 h under continuous stirring. The suspension was then transferred to a Teflon-lined stainless steel autoclave and aged during 24 h at 100 ºC. The as-prepared sample was isolated by filtration, washed with distilled water and allowed to dry in air at room temperature for 2 days. The organic template was removed by calcination in air at 550 ºC for 8 h (heating rate of 1 ºC·min-1). The surface (internal and external) of the synthesized MCF was functionalized according to the procedure described by Russo et al.38 in which 0.5 g of MCF was dried in a vacuum oven during 4 h at 120 ºC, followed by the addition of 10 ml of toluene and 0.3 ml of MPTS under vigorous stirring at room temperature. Then, the samples were transferred to a Teflon-lined stainless steel autoclave and heated at 100 ºC for 24 h. The functionalized MCFs were filtrated, washed with toluene and dried at 120 ºC overnight. The final product was labeled as MCF-MP. The MCF and MCF-MP pore/window size distribution, specific surface area and pore volume were determined by nitrogen sorption analysis, using a Tristar 3000 instrument from Micromeritics Instrument Corporation. Prior to the measurements, MCF was outgassed for at least 8 h at 180 ºC, whereas MCF-MP was outgassed at 120 ºC overnight. The pore and window size distributions were calculated according to the simplified BdB (BroekhoffdeBoer) - FHH (Frenkel-Halsey-Hill) method,39 whereas the specific surface area (SA) was


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obtained using the BET (Brunhauer-Emmett-Teller) method. The pore volume was calculated using a single point adsorption value at the relative pressure of 0.990. The MCF-MP was also characterized by Thermogravimetric analysis (TGA) carried out on a TGA/DSC 3+ instrument from Mettler Toledo, with heating rate of 10 ºC·min-1 under a N2 flow of 50 ml·min-1. Using the measured weight loss Wloss (in %) per 100 gram of MCF, the surface loading (Ns) 40 and surface density (D) 41 of the MP-ligand were estimated using Eqs. (1) and (2), respectively, where MW is the molecular weight of the ligand and NA is the Avogadro number.

= /

(1)

= ∙ /

(2)

2.3 Labelling FateDH with Cy3 and FaldDH with Cy5 Both spectroscopic probes are sulfonated and amino reactive which make them hydrophilic and ideal for attaching to proteins. The probes were covalently bound to the enzymes, Cy3 to FateDH (Cy3-FateDH) and Cy5 to FaldDH (Cy5-FaldDH), in separate vials based on the reaction of NHS (N-hydroxysuccinimide) ester groups on the probes (Figure 1) with amine groups on each enzyme. The dye modification was performed by mixing the working stock solution of either enzyme (2±0.2 mg of FateDH and 0.7±0.2 mg of FaldDH in 900µl of 0.1M phosphate buffer, pH 8) with 100 µl of the dye stock solution (300±20 µg of Cy3 and 40±2 µg of Cy5 in 100µl of Millli-Q water), based on the labelling protocol provided by Lumiprobe. After vortexing for 1 min, the mixtures were kept on ice overnight. The labeled enzymes were purified on a size-exclusion column (PD Miditrap G-25, GE Healthcare) with 0.1M phosphate buffer and pH 8 to remove all non-bound dyes. The numbers of attached dyes (Cy3 or Cy5) per enzyme (the degree of labelling DOL in Table 1) were determined based on the absorbance ratio of the dye and the enzyme before and after the binding reaction using the molar absorption coefficient of dyes (Table 1) and the extinction coefficient of the enzymes (E1%= 15.9 for FateDH and E1%= 10.0 for FaldDH, as provided by supplier, SigmaAldrich), see Eq S1 in Supporting information. The results show that on the average about 3.1 Cy3 molecules are bound to each FateDH and 5.8 Cy5 molecules to FaldDH, although, we do not know to which of the lysines in the enzymes42 the dyes are attached. The degree of labelling for FaldDH is higher than for FateDH, and one reason can be that the larger FaldDH contains a higher number of lysines available for reaction with Cy5. To avoid protein-protein aggregation of the larger FaldDH during the size-exclusion filtration of the 7 ACS Paragon Plus Environment

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mixture of Cy5 and enzyme, the mass of FaldDH loaded on the column was kept below 1 mg. 2.4 Co-immobilization of the two enzymes Two methods to co-immobilize FateDH and FaldDH in both unmodified MCF and MCF-MP particles were investigated, using three different enzyme mass ratios FateDH:FaldDH of 1:1, 1:3 and 1:15. It should be noted that we have not made an exhaustive optimization of the immobilization strategy, the focus in this study was on comparing two principally different approaches. Moreover, the enzymes used in this work are commercial enzymes obtained as lyophilized powders that may contain impurities. No purifications were performed for sake of comparison between our results and other studies which used the same enzymes. In the first method called simultaneous co-immobilization, 10 µl of FateDH at 2, 10 or 30 mg·ml-1 and 20 µl of FaldDH at 15 mg·ml-1 were added simultaneously to the appropriate volume of MCF or MCF-MP working stock solutions (20 mg·ml-1) to give a total amount of added enzyme of 150 mgenzymes·gparticles-1. The combined enzyme mixture was incubated with the particles for 4 h in a thermomixer at 37 ºC and 900 rpm. 4h immobilization is chosen to ensure that all formaldehyde dehydrogenase can be immobilized.31 In the second method called sequential co-immobilization, initially the 10 µl of FateDH at 2, 10 or 30 mg·ml-1 was mixed with the same volumes of MCF or MCF-MP working stock solution and agitated for 2 h which should be sufficient for FateDH since this enzyme is a smaller enzyme compared to FaldDH.43 After that the 20 µl of FaldDH at 15 mg·ml-1 was added to the pre-incubated FateDH sample and the immobilization was allowed to proceed for another 4 h using a thermomixer at 37 ºC and 900 rpm. The stock solutions of enzymes, MCF and MCF-MP were prepared using 100 mM phosphate buffer and pH 5.6 which is close to the pI of the enzymes resulting in higher enzyme uptaking.31 After the immobilization, the samples were centrifuged and washed three times with 100 mM phosphate buffer and pH 6.5 which has been shown to be the optimum pH of both enzymes for the reduction reactions.10 The concentrations of the enzyme solutions were measured using a Nanodrop One Instrument from Thermo Scientific. The total amount of immobilized enzyme was determined indirectly through the amount of enzymes that remained in the supernatant after immobilization by measuring the protein UV absorption at 280 nm, using the extinction coefficient E1% = 15.9 for FateDH and E1% = 10.0 for FaldDH as provided by supplier, Sigma-Aldrich. 8 ACS Paragon Plus Environment

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In order to evaluate the two different co-immobilization methods, the degree of immobilization for each enzyme type was determined separately by exploiting that the labeled enzymes (Cy3-FateDH and Cy5-FaldDH) have well separated absorption spectra in the visible region, with absorption maxima at 550 nm for Cy3 and at 650 nm for Cy5 (Table 1). Again, the determination was indirect by measuring the absorption of protein-attached Cy3 and Cy5 in the supernatant after the immobilization. By using the DOL of the proteindye complex (Table 1) the concentration of each enzyme was calculated independently.

2.5 Measurement of catalytic activity The catalytic cascade reactions (step 1 and 2 in Scheme 1) in free solution (no particles) were performed by mixing FateDH (0.1, 0.5 or 1.5 mg·ml-1), FaldDH (1.5 mg·ml-1), NADH (100 mM), and KHCO3 (200 mM). All solutions were prepared using 100 mM phosphate buffer pH 6.5. The reactions were performed at 37 ºC in a closed reactor under CO2 atmosphere at 5 bars pressure for 1 h under stirring. The activity tests with co-immobilized enzymes were done under the same conditions including the final total enzyme concentrations, but using the enzymes immobilized as described above. We are not aware of any studies of CO2-solubility in confined water. Assuming the same Henry’s law constant in pore water as in bulk water (KH = 3.3·10-4 (mol·m-3·Pa-1 at 25oC) the equilibrium [CO2(aq)] is approximately 0.16 M at our CO2-pressure of 5 bar.44 This is similar to the expected molar concentrations of substrate but typically a factor 100-1000 higher than the effective protein concentration in the pores according to Nabavi Zadeh et al.30 All activity tests were performed using enzymes which were not labeled with Cy3 and Cy5. The final product, formaldehyde, was quantified using Nash’s method45 since the gas chromatograph with FID detector has low sensitivity for formaldehyde. Nash’s reagent was prepared by mixing 25 g of ammonium acetate, 2.1 ml of acetic acid and 0.2 ml of acetyl acetone in a total volume of 100 ml in water. After the enzymatic reaction, equal amounts of Nash’s reagent and the reaction product were mixed. The samples were agitated for 1 h at 37 ºC. After that, the UV-vis absorbance of the sample at 414 nm was measured.

2.6 Förster resonance energy transfer (FRET) measurements FRET is a process in which energy is transferred non-radiatively via long-range dipoledipole coupling from an excited donor fluorophore to another dye molecule, the acceptor. In this work, the derivatives Sulfo-cyanine3 NHS ester (Cy3) and Sulfo-cyanine5 NHS ester 9 ACS Paragon Plus Environment

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(Cy5) were used as donor and acceptor, because they can be covalently bound to FateDH and FaldDH, respectively. Cy3 and Cy5 are commonly used as a fluorophore pair in FRET studies,46-49 and are suitable for studies in silica particles since the negative charge minimizes binding to the silica pore walls which carry a negative charge (point of zero charge = 2). It also makes them hydrophilic which reduces the dye adsorption to the surface of the water soluble enzyme, and these fluorescent probes are photostable which avoids potential problems caused by photobleaching.43 Cy3 and Cy5 constitute a good donor-acceptor pair, because the fluorescence emission spectrum of the donor Cy3 has a large spectral overlap with the absorption spectrum of the Cy5 acceptor chromophore. To observe FRET, first the orientation of the transition dipole moments of the two chromophores must be approximately parallel to each other.49 Secondly, the donor and acceptor dye must be in close proximity of each other, as characterized by the Förster distance R0 given 50 by Eq.3 51 R0 = 0.211(ƙ2n-4ΦD J(λ))1/6

(3)

where ΦD is the fluorescence quantum yield of the donor in the absence of the acceptor, J(λ) is the spectral overlap integral between the normalized donor fluorescence and the acceptor excitation spectra, ƙ2 is the dipole orientation factor between donor and acceptor transition moments and n is the effective refractive index. The R0 value for the Cy3/Cy5-pair under our conditions is approximately 6 nm in free solution which is in agreement with previous reports.52-53 However, after immobilization, the R0 value can change due to two reasons: (a) possible self-quenching of Cy3 (due to high concentration of labeled protein in the pores) which could reduce the quantum yield of the donor and (b) enhancement the fluorescence intensity of donor due to high viscosity causing a higher quantum yield.54-55 After immobilization in MCF and MCF-MP, the R0 value for each sample was calculated individually to make sure that the values are close enough for comparison of the transfer efficiency values (see Supporting information, section S3 for details). In the calculation of R0, we have used the value ƙ2 = 2/3 for the dipole orientation factor, which corresponds to random orientation of the transition moments of the Cy3 and Cy5 chromophores. The assumption of free movement of the chromophores is supported by time-resolved anisotropy measurements on Cy3-FateDH in free solution and after immobilization (see Supporting information, section S3), which also strongly support that the donor dye does not adsorb to the silica wall as expected since both are negatively charged. 10 ACS Paragon Plus Environment

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Experimentally, the strength of the FRET is presented as transfer efficiency (E). The E value is a direct measure of the fraction of photons absorbed by the donor that is transferred to an acceptor. It is defined as the ratio between transfer rate kT and the total excited state decay rate of the donor, E = kT / (τD-1 + kT)

(4)

where τD is the lifetime of the donor in the absence of acceptors. It is commonly measured as the relative lifetime of the donor in presence (τDA) and absence (τD) of the acceptor,47

E = 1 - (τDA / τD)

(5)

which has the advantage, in turbid samples such as our particle-based system, that it does not involve measurements of absolute intensity. Eq. 6 shows that E depends on the inverse of the sixth power of the distance (r) between the two fluorophores E = R06 / (R06 + r6)

(6)

which has made FRET a useful and sensitive method to measure distances between fluorophores, for example in protein-protein interaction studies.56-57 It is seen that the Förster distance, R0, represents the distance between the two chromophores where the transfer efficiency is 50% for a given fluorophore pair. The FRET efficiency was obtained from fluorescence lifetime measurements using time correlated single photon counting (TCSPC). The samples of co-immobilized enzyme or free enzymes (no particles) were excited by a PicoQuant pulsed (10 MHz) laser diode as the excitation source. All the measurements were performed at 37 °C and in 0.1 M phosphate buffer to give pH 5.6 (if not otherwise stated). Excitation was at 483 nm, where the donor Cy3 can be excited but the acceptor Cy5 has no absorption (See Figure S6 in Supporting information), and the emission intensity time profiles were recorded at 565 nm, where the donor (Cy3) has the emission maximum. The measurement was stopped when 10000 counts had been collected in the peak channel. The emitted light was detected at the magic angle to the excitation light through a monochromator tuned to the maximum emission wavelength of the sample. The photons were collected by a R3809U-50 microchannel plate


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photomultiplier tube from Hammamatsu and fed into a multichannel analyzer (Edinburgh Analytical Instruments) with 4096 channels. Fitting the intensity decays to mono, bi- or triexponential functions was performed using Fluofit Pro v.4 software (PicoQuant GmbH). The average lifetime was amplitude weighted,

=

∑

∑

(7)

where αi is the amplitude of the component with lifetime τi. The lifetime values of both Cy3 and Cy5 are sensitive to the local environment,54 so the spatial distribution of two enzymes should be kept as similar as possible inside the pores during lifetime measurements in presence and absence of acceptor. To that end, Cy3FateDH was co-immobilized with FaldDH in two ways, always keeping the amount of the enzymes the same. Either FaldDH was labelled with Cy5, in which case the sample is denoted Cy3-FateDH+Cy5-FaldDH (donor in presence of acceptor), or the FaldDH lacked the Cy5-label which is referred as the sample with donor in absence of acceptor and denoted Cy3-FateDH+FaldDH.

3. Results and discussion 3.1 Characterization of siliceous mesostructured cellular foams The physical properties of the MCF and MCF-MP are shown in Table 2. The mean pore size for both materials was 32.8 nm. As previously reported,31 this pore size is large enough for the immobilization of FaldDH (molecular dimensions of 8.6 x 8.6 x 19 nm).58 The large pores in the MCF and MCF-MP are connected by narrow windows of 11.3 nm diameter, which is a desirable property that limits enzyme leakage. The specific surface area and pore volume are close to the values obtained previously32, and also it is seen that functionalization of MCF with mercaptopropyl groups slightly reduced these properties of the material. The nitrogen sorption isotherms and pore size distributions are shown in Figure S1 and Figure S2 in the Supporting information. The incorporation of the mercaptopropyl groups in the MCF was confirmed and quantified by TGA (Table 3 and Figure S3 in the Supporting information). The MCF-MP presented surface loading of 0.8 mmol·g- 1 and surface density of 0.8 ligand·nm-2.


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Table 2. Properties of the MCFs Analyzed by Nitrogen Sorption Sample Mean Mean Surface area Window size pore size (m2·g-1 ) (nm) (nm) MCF 11.3 32.8 612 MCF-MP 11.3 32.9 527

Pore volume (cm3·g-1 ) 2.45 2.27

Table 3. Properties of the Functionalized MCFs Calculated from TGA Analysis Sample Weight loss Surface loading (Ns) Surface density (D) (%) (mmol·g-1) (ligand·nm-2) MCF-MP 5.9 0.8 0.8

3.2 Influence of method of co-immobilization Two methods of co-immobilization in MCF without modification were investigated, simultaneous co-immobilization, when the two enzymes were immobilized together (for 4h), and sequential co-immobilization, where FateDH was immobilized first (for 2h) after which the FaldDH was added. The outcome was compared using FateDH and FaldDH at a mass ratio of 1:15, aiming for a total enzyme loading of 150 mgenzymes·gsupport-1. This ratio was used since it has been reported previously by Cazelles et al.

10

as the optimum ratio for the

cascade reaction. Using UV-absorption at 280 nm, it is possible to measure the total amount of immobilized enzyme, which was found to be roughly 95% of the added amount. However, since the initial amount of FateDH represents only 6.25% of the total amount of enzyme added, it is not possible to determine separately how much of this enzyme becomes immobilized. By exploiting the separate Cy-labels on FateDH and FaldDH, which have essentially non-overlapping absorption spectra (See Figure S6 in Supporting information), it was possible to measure how much of each enzyme remained in the supernatant even when they are mixed, and calculate the amount of immobilized FateDH and FaldDH individually. Table 4 shows the degree of immobilization (% of the initial amount of each enzyme that became immobilized) for the FateDH and FaldDH with the two methods of coimmobilization, and the total amount of immobilized enzyme measured by absorption at 280 nm both with and without the dye-labels.


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Table 4. Degree of Immobilization of each Enzyme after Co-Immobilization in MCF using the Concentration Ratio FateDH: FaldDH = 1:15 Method DOI DOI Total amount of Total amount of Cy3-FateDHa Cy5-FaldDHa immobilized enzyme immobilized enzyme (%) (%) with labelling (%)b without labelling (%)c Simultaneous 15 95 90 94 Sequential 95 95 95 95 a

Degree of immobilization (DOI) calculated from absorption of Cy3- and Cy5-labeled enzymes, respectively, in supernatant, as an average from 3 independent experiments, with the uncertainty of ± 5% calculated as half the maximum variation. b Calculated from absorption of total enzyme in the supernatant at 280 nm using the degree of labeling and the absorption of Cy3 and Cy5, respectively at 550 and 650. c Calculated from absorption of total enzyme in the supernatant at 280 nm using extinction coefficient E1% = 10.0, since FaldDH used with higher concentration than FateDH.

It can be seen that at the end of the immobilization process, the simultaneous coimmobilization results in that only 15% of the added FateDH was immobilized, whereas 95% of the FaldDH was adsorbed into the support material. By contrast, in the sequential method 95% of both enzymes were co-immobilized. The low degree of immobilization of FateDH using simultaneous co-immobilization suggests that there is a higher selectivity of the FaldDH to adsorb on the silica surface. Being immobilized more favorably and in higher concentrations, FaldDH, which is a large enzyme (molecular dimensions of 8.6 x 8.6 x 19 nm)

58

would occupy the pores and partially block the windows of the MCFs hindering

FateDH (molecular dimensions of 5.3 x 6.8 x 10.9 nm)59 to access the interior of the pores. Therefore, by co-immobilizing the enzymes sequentially, FateDH could enter the pores more efficiently, leading to a high degree of immobilization.

3.3 Specific catalytic activity The specific catalytic activity (conversion rate of CO2 to formaldehyde per total mass of enzymes) of the cascade reaction was first evaluated using 0.1 mg·ml-1 FateDH and 1.5 mg·ml-1 FaldDH (FateDH:FaldDH = 1:15), which has been found to be a suitable mixing ratio for this cascade reaction.10 The specific catalytic activity amounted to 0.12 ± 0.02 µmol·genzymes-1·min-1 using the enzyme free in solution. Interestingly, the simultaneous co-immobilization in the unmodified MCF, led to no detectable catalytic activity or formation of formaldehyde, most probably due to the low amount of FateDH that became immobilized using this method (Table 4). However, using the sequential method of co-immobilization the specific catalytic activity of the cascade reaction was high enough to


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ACS Catalysis

be measured. Based on these results, all following experiments were performed using only enzymes immobilized by the sequential co-immobilization method. Figure 2 shows the specific catalytic activity in the three cases studied: free in solution and sequentially co-immobilized in MCF and MCF-MP, respectively. For the mass ratio FateDH:FaldDH = 1:15, Figures 2a and 2b show that co-immobilizing the enzymes in MCF did not increase the efficiency of the cascade reaction significantly compared to the enzymes free in solution. In contrast, upon co-immobilization in MCF-MP the specific catalytic activity for formaldehyde formation increased about 4 times, as shown in Figure 2c. This improvement in the specific catalytic activity of the cascade reaction is very promising considering that these two catalytic reaction steps (reduction of CO2 to formate and subsequent reduction of formate to formaldehyde) are the limiting steps for the overall

conversion of CO2 to methanol.10-11, 18 Figure 2. Specific activity of the cascade reaction using FateDH and FaldDH (a) free in solution, and coimmobilized in (b) MCF and (c) MCF-MP. The enzymes were co-immobilized using the sequential method. In parenthesis are the mass ratio of FateDH: FaldDH used in the reactions. Values are presented as means ± standard deviation (n=3).

These results are in agreement with our previous study that showed an increased activity of FaldDH when immobilized in MCF functionalized with mercaptopropyl groups.31 The reason for the improved catalytic activity of the co-immobilized enzymes in MCF-MP can be due to beneficial orientation of the enzymes inside the pores, in such a way that the active sites are more accessible to the substrates.35, 60 Another possible explanation, is that when enzymes are confined inside the pores, they tend to be in closer proximity to each other allowing substrate channeling between the active sites.21 The immobilization of enzymes can also lead to an improved enzyme stability.60


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Furthermore, we evaluated if higher concentrations of FateDH could be used in order to improve the formaldehyde production. However, by increasing the concentration of FateDH to 0.5 mg·ml-1 and keeping the same concentration of FaldDH (1.5 mg·ml-1), resulting in a mass ratio FateDH:FaldDH = 1:3, we found the catalytic activity of the free and coimmobilized enzymes to become significantly reduced (Figures 2a and 2b). When the concentration of FateDH was further increased to 1.5 mg·ml-1 (FateDH:FaldDH = 1:1), the free enzymes in solution did not show a quantifiable formation of formaldehyde, whereas the co-immobilized enzymes in MCF showed a catalytic activity comparable to the 1:3 ratio. When the enzymes were co-immobilized in MCF-MP, shown in Figure 2c, a similar trend in decreasing activity was observed but the activity remained slightly higher than in MCF and free solution also for these enzyme mixing ratios. The decreased activity with more equal enzyme concentrations are in agreement with the results obtained by Cazelles et al..10 The authors reported that it was necessary to use an excess of FaldDH over FateDH to drive the reaction in the direction of the reduction. In this cascade reaction system (Scheme 1), the formic acid is deprotonated at the used pH to formate which participates in two competing reactions: the oxidation of formate to CO2 (catalyzed by FateDH), which is a spontaneous reaction and the reduction of formate to formaldehyde (catalyzed by FaldDH), which is not thermodynamically favored. This can be seen by comparing the free energy (∆rG’o) of the reactions shown in Eqs 8 and 9. CO2 + NADH ↔ HCOO- + NAD+

(8)

HCOO- + NADH +2H+ ↔ HCHO + NAD+ + H2O

(9)

The reduction of CO2 to formate and of formate to formaldehyde are both thermodynamically unfavorable reactions under standard conditions (∆rG’o = 20.6 kJ·mol-1 and 78.2 kJ·mol-1, respectively, at pH=7.3 and an ionic strength of 0.25).15 Therefore, using high concentrations of substrate and cofactor tends to shift the equilibrium towards the production of formaldehyde, and using higher amounts of FaldDH in relation to FateDH also helps to shift the equilibrium towards the final product by removing the intermediary formate more efficiently. A kinetic analysis of the system was not possible, since there is no information available about the Km and Vmax for the conversion of formate to formaldehyde by FaldDH. In fact, it has been proven difficult to measure the kinetic reaction parameters by increasing the concentration of formic acid and keeping the pH constant.11


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ACS Catalysis

3.4 Förster resonance energy transfer Förster resonance energy transfer (FRET) is a spectroscopic process by which an excited fluorophore (the donor) transfers its energy to a neighboring molecule (the acceptor) by non-radiative dipole-dipole interaction.47,

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FRET has become widely used as a

spectroscopic ruler to measure distances on the macromolecular scale because the efficiency of the process depends strongly on the distance between the donor and the acceptor.46 Here, FRET measurements were performed with the aim to measure the distance between the two enzymes when they are co-immobilized in the confining silica materials, in order to investigate to what extent the enzyme-enzyme distance can affect the catalytic activity of the cascade. The same procedure of co-immobilization was used as in the activity measurement, except that the enzymes were labelled with dyes in the FRET measurements (Cy3-FateDH and Cy5-FaldDH). Notably, Table 4 shows that the dye labels had no detectable effect on the degree of immobilization, at least in terms of the total protein uptake, so the FRET results were most likely obtained with the same enzyme concentrations in the pores as the activity measurements. Using Eq.5 the efficiency of the energy transfer (E) between donor and acceptor was calculated from measurements of lifetime values of donor in presence and absence of acceptor. Table 5 shows the measured average lifetime values and calculated energy transfer efficiency of the free and co-immobilized enzymes in unmodified MCF and in MCF-MP at the three different mixing ratios used in the activity measurements. Table 5. Lifetimes and Energy Transfer Efficiency of Enzymes Co-Immobilized in MCF and MCF-MP at Different Enzyme Concentration Ratios Free enzymes MCFa MCF-MPa b Ratio all three ratios 1:15 1:3 1:1 1:15 1:3 1:1 τDA (ns)c 0.64±0.05 0.65±0.05 0.73±0.05 0.88±0.05 0.52±0.05 0.65±0.05 0.85±0.05 τD (ns)d 0.64±0.05 1.45±0.05 1.30±0.05 1.10±0.05 1.45±0.05 1.45±0.05 1.33±0.05 Ee 0 0.56±0.02 0.44±0.02 0.20±0.02 0.64±0.02 0.55±0.02 0.36±0.02 a

Total protein loading 150 mgenzymes·gsupport-1. Mixing ratio Cy3-FateDH: FaldDH-Cy5. c Average lifetime of donor (Cy3) in presence of acceptor (Cy5). d Average lifetime of donor (Cy3) in absence of acceptor (Cy5). e The transfer efficiency calculated from the average lifetime values using Eq 5. All values are average values± maximum variation from 8 independent experiments for each sample. b

As can be seen from Table 5, there is no detectable FRET when the two enzymes are free in solution. This observation is consistent with an average distance of about 40 nm or more between them in the dilute solutions since this value is large compared to the Förster radius R0 in equation (3) of about 6nm. It is also seen that immobilization increases the 17 ACS Paragon Plus Environment

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transfer efficiency significantly, to above 50% in several cases. Importantly, the E value is higher in MCF-MP than in MCF at a given mixing ratio. Furthermore, the FRET efficiency is the highest at the concentration ratio 1:15 for both MCF and MCF-MP, and decreases as the concentration of the two enzymes become more equal. Ideally, when different protein concentrations are compared the fluorescence lifetimes should be constant, but Table 5 shows a slight decrease in

τD when the concentration of FateDH is increased, especially in

the MCF particles. This is most likely due to self-quenching of FateDH-bound Cy3, and as described in Supporting information, section S3, it means that the Förster radius R0 becomes protein-concentration dependent. However, the calculated values of R0 at different enzyme concentrations only vary between 6.6 and 6.8 nm (See Table S3 and Table S4 in Supporting information). Since the maximum variation of 0.2 nm is much smaller than the pore diameter (33 nm), we take R0 to be essentially constant when the transfer efficiency is compared at different enzyme concentrations. Our system is complex from a FRET point of view, because each Cy3-FateDH donor protein may be surrounded by several Cy5-FaldDH acceptors and vice versa. Berney et al.61 have used Monte Carlo simulations to study systems with multiple donors and acceptors. Based on the results they suggest suitable values for the number of donors per acceptor (RDA) in FRET experiments aimed at the effect of changes in the mean distance between donors and acceptors. The RDA values in the present study (number ratios 0.04, 0.17 and 0.53 for the mass mixing ratios 1:15, 1:3 and 1:1 respectively) partly covers the RDA range 0.1-10 where the simulations indicated the highest sensitivity to donor-acceptor distance (for RDA >10 the E is close to zero and for RDA

to Formaldehyde b - ACS Publications - American Chemical Society

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