Cascade catalysis in membranes with enzyme immobilization for multi-enzymatic conversion of CO2 to methanol

New Biotechnology  Volume 00, Number 00  February 2015

RESEARCH PAPER

Research Paper

Cascade catalysis in membranes with enzyme immobilization for multi-enzymatic conversion of CO2 to methanol Jianquan Luo1, Anne S. Meyer1, R.V. Mateiu2 and Manuel Pinelo1 Q1 1 Department of Chemical and Biochemical Engineering, Center for BioProcess Engineering, Building 229, Technical University of Denmark, DK-2800 Kgs. Lyngby, Denmark 2 Center for Electron Nanoscopy, Danchip, Technical University of Denmark, DK-2800 Kgs. Lyngby, Denmark

Facile co-immobilization of enzymes is highly desirable for bioconversion methods involving multienzymatic cascade reactions. Here we show for the first time that three enzymes can be immobilized in flat-sheet polymeric membranes simultaneously or separately by simple pressure-driven filtration (i.e. by directing membrane fouling formation), without any addition of organic solvent. Such coimmobilization and sequential immobilization systems were examined for the production of methanol from CO2 with formate dehydrogenase (FDH), formaldehyde dehydrogenase (FaldDH) and alcohol dehydrogenase (ADH). Enzyme activity was fully retained by this non-covalent immobilization strategy. The two immobilization systems had similar catalytic efficiencies because the second reaction (formic acid ! formaldehyde) catalyzed by FaldDH was found to be the cascade bottleneck (a threshold substrate concentration was required). Moreover, the trade-off between the mitigation of product inhibition and low substrate concentration for the adjacent enzymes probably made the coimmobilization meaningless. Thus, sequential immobilization could be used for multi-enzymatic cascade reactions, as it allowed the operational conditions for each single step to be optimized, not only during the enzyme immobilization but also during the reaction process, and the pressure-driven mass transfer (flow-through mode) could overcome the diffusion resistance between enzymes. This study not only offers a green and facile immobilization method for multi-enzymatic cascade systems, but also reveals the reaction bottleneck and provides possible solutions for the bioconversion of CO2 to methanol.

Introduction Q2 Global warming and energy shortages demand the development of carbon dioxide (CO2) capture and reutilization technologies. Efficient conversion of atmospheric CO2 to fuels offers a promising approach, which benefits from greenhouse gas fixation and the production of renewable fuels and chemicals [1]. Partial hydrogenation of CO2 has been accomplished by means of heterogeneous catalysis, electrocatalysis and photocatalysis [1,2]. The use of enzymes is particularly appealing since it provides a facile low Corresponding authors: Luo, J. ([email protected]), Pinelo, M. ([email protected])

temperature route for the generation of low-carbon fuels directly from CO2 [3–6]. Obert and Dave were the first to report an enzymatic combination of formate dehydrogenase (FDH), formaldehyde dehydrogenase (FaldDH) and alcohol dehydrogenase (ADH) for sequential reduction of CO2 to methanol, where reduced nicotinamide adenine dinucleotide (NADH) was used as the terminal electron donor for the enzymatic reaction [5]. This reaction was achieved by reversing the biological metabolic pathways. Although the biocatalytic reduction of CO2 is thermodynamically feasible [7], this multi-enzymatic conversion of CO2 to methanol was found to be quite inefficient, taking several hours and only www.elsevier.com/locate/nbt

http://dx.doi.org/10.1016/j.nbt.2015.02.006 1871-6784/ß 2015 Published by Elsevier B.V.

Please cite this article in press as: Luo, J. et al.1, Cascade catalysis in membranes with enzyme immobilization for Biotechnol. (2015), http://dx.doi.org/10.1016/j.nbt.2015.02.006

1 multi-enzymatic conversion

of CO2 to methanol, New

NBT 761 1–9 RESEARCH PAPER

Research Paper

producing a very low concentration of methanol (less than 5 mM) [5]. Various immobilization methods have been used to stabilize the enzymes and increase the catalysis yield [8–10]. Regeneration of NADH by co-immobilized enzymes has also been investigated in order to enhance methanol production [11,12]. For example, Wang et al. constructed an efficient multi-enzyme cascade system based on ultrathin, hybrid microcapsules by combining the unique functions of catechol and gelatin (enzymes were sequentially immobilized in the microcapsules by entrapment and covalent binding), which maintained a methanol yield of 52.6% even after nine recycling steps [10]. El-Zahab et al. immobilized cofactor and enzymes on different polystyrene particles for in situ cofactor regeneration, which allowed methanol to be continuously produced for ten cycles without supplement of NADH [11]. However, these strategies have only marginally improved the final methanol concentration when compared with the first report by Obert and Dave [5]. In order to improve the catalysis efficiency of multi-enzymatic reactions, a more efficient immobilization strategy is required. Firstly, the immobilization procedure should be simple and facile, and enzyme denaturation should be minimized during immobilization. It was found that encapsulation was the most common strategy used for the immobilization of multi enzymes [5,10,12,13]. Secondly, it is necessary to have a high enzyme loading and avoid enzyme leakage during the subsequent immobilization and reaction. Thirdly, the mass transfer resistance of the enzymes in the scaffold should be very low, so the substrate/ intermediate can easily diffuse to the targeted enzymes. Fourthly, the product should be removed immediately from the enzyme active site in order to decrease product inhibition. In situ regeneration of cofactor in multi-enzymatic conversion of CO2 actually is a strategy of product removal. For example, Cazelles et al. found that the methanol productivity increased by 5 times in 3 hours with the NADH regeneration system [12]. Although numerous multi-enzyme immobilization approaches have been developed, which have shown great potential in enhancing the overall enzymatic

New Biotechnology  Volume 00, Number 00  February 2015

performance [14], it is still worth exploring new immobilization methods with cheaper materials and simpler procedures. Recently, biocatalytic membranes with enzymes immobilized in industrially manufactured membranes have attracted growing attention for both biosensor and bioreactor applications [15,16]. The porous membrane, working as a selective barrier as well as a support for enzyme immobilization enabling enzyme re-use, may also improve enzyme stability, eliminate product inhibition, and allow for continuous processing [17]. Fouling-induced enzyme immobilization in polymeric membranes was proposed based on many parallels between membrane fouling mechanisms and enzyme immobilization strategies [18–21]. This can be accomplished by fostering membrane fouling through a simple pressure-driven filtration of the enzyme solution. The enzymes can be immobilized in the membranes via various non-covalent bonds, for example, hydrogen bonding, entrapment and hydrophobic or electrostatic adsorption [19]. It is a promising technique for the coimmobilization of several enzymes as the immobilization can be achieved under the same conditions in a single step and the noncovalent bonding retains their activity [13]. Moreover, the multienzyme cascade system can also be constructed by carrying out layer-by-layer sequential deposition [22] on one membrane or by using multistage membrane filtration (enzymes can be immobilized separately) [18]. It is well known that enzyme cascade reactions ‘in nature’ usually put the enzymes in close proximity to one another; the question therefore is whether it is better to have the enzymes immobilized separately to allow for individual optimization of the enzyme reactions (this reaction relies on a combination of enzymes from various sources, which may therefore have different optimal conditions). To evaluate this, we propose two immobilization strategies for multi-enzymatic catalysis in membranes by fouling-induced enzyme immobilization, called co- and sequential immobilizations, as illustrated in Fig. 1. The hypothesis is that the constant low substrate concentration could decrease the rate of the cascade reaction when enzymes are co-immobilized on the same membrane, but on the other hand it is uncertain whether

FIGURE 1

Two proposed strategies for production of methanol from CO2: co-immobilization versus sequential immobilization of enzymes in membranes. 2

www.elsevier.com/locate/nbt Please cite this article in press as: Luo, J. et al.1, Cascade catalysis in membranes with enzyme immobilization for Biotechnol. (2015), http://dx.doi.org/10.1016/j.nbt.2015.02.006

multi-enzymatic conversion

of CO2 to methanol, New

NBT 761 1–9

the potential advantage of a shorter mass transfer path for a product from one reaction to its subsequent encounter with a new enzyme for which it is now a substrate (for conversion to the next product in the sequence) will be advantageous. To the best of our knowledge, this is the first attempt to construct in vitro a multienzymatic assembly in polymeric membranes. In addition, by comparing the results from free, co- and sequentially immobilized enzymes, the mechanisms of the multi-enzymatic reaction of CO2 into methanol are discussed.

Materials and methods Chemicals and membranes Formate dehydrogenase (EC 1.2.1.2, homo-dimer, 76 kDa) from Candida boidinii (FDH) [23], formaldehyde dehydrogenase (EC 1.2.1.46, homo-dimer, 150 kDa) from Pseudomonas sp. (FaldDH) [24], alcohol dehydrogenase (EC 1.1.1.1, homo-tetramer, 141 kDa) from Saccharomyces cerevisiae (ADH), and L-glutamic dehydrogenase (EC 1.4.1.3, homo-hexamer, 300 kDa) from Bovine liver (GDH) [25] were purchased from Sigma–Aldrich (St. Louis, MO, USA). These commercial powders or liquids are not pure enzymes and the protein content is determined by Bradford protein assay. bNicotinamide adenine dinucleotide reduced form (NADH, >97 wt%), b-nicotinamide adenine dinucleotide hydrate (NAD+), formic acid (>96%), formaldehyde (37%), methanol (99.9%), and Lglutamic acid were purchased from Sigma Aldrich. All the enzyme and substrate solutions were prepared using 0.1 M Tris–HCl buffer (pH = 7.0) unless otherwise stated. CO2 gas (>99.5%) in a cylinder was purchased from AGA A/S (Denmark). Commercial UF membranes (PLTK and PLGC, Millipore) were used in this work, which has a regenerated cellulose skin layer on a polypropylene support, and their molecular weight cut-offs are 30 and 10 kDa, respectively.

Experimental set-up and procedure Pure CO2 gas from a cylinder was first saturated with H2O before entering the reactor. The flow rate of gas (measured by the speed of bubble emission) was controlled in the same manner in all the experiments by controlling the pressure valve. Wet CO2 was continuously bubbled into solution through a syringe needle (0.6 mm  25 mm). The dead-end filtrations were performed in a stirred cell (Amicon 8050, Millipore, USA). The descriptions of equipment and procedure can be found in our previous work [18,19]. The PLTK membranes (30 kDa) were placed on the membrane holder in ‘sandwich’ mode (with their own support layer facing the feed and an extra polypropylene support beneath the skin layer). The membranes were first dipped in a 5% NaCl solution for 30 min and then filtered with deionized water for another 30 min at 1 bar (procedures according to the manufacturers’ instructions). Afterwards, the water permeability of the membranes was measured at 2 bar with buffer for 30 min. Then, the enzyme solutions (50 mL for co-immobilization and 30 mL for sequential immobilization) were put into the cell with a 30 kDa membrane for the enzyme immobilization operations. All the experiments were repeated at least two times.

Enzyme immobilization The amount of enzyme for immobilization was 100 mL liquid of FDH, 1.0 mg solid of FaldDH, and 1.5 mg solid of ADH, respectively.

RESEARCH PAPER

Enzyme immobilization was carried out at a pressure of 2 bar, and permeate was collected in precision cylinders for analysis. The cylinders were replaced manually for every 4 mL. At the end of filtration, the ‘fouled’ membrane was washed with 10 mL of buffer at a pressure of 2 bar, and then rinsed 3 times with buffer without pressure (5 mL of buffer each time). The final retentate and washing/rinsing residue were combined in order to calculate the amount of immobilized enzyme by mass balance. The final steady buffer flux was recorded as permeability for the ‘fouled’ membrane. For sequential immobilization, three membranes with FDH, FaldDH and ADH were sequentially stacked and Parafilm O-rings were placed between membranes to seal the membrane stack. Then the permeability of the tri-membrane stack was determined by buffer solution. For a dead-end filtration process at constant pressure, the laws of filtration can be stated as [26]:  n d2 t dt ¼ K (1) dV dV 2 where t is filtration time (s), V is permeate volume (m3), K is constant and n can have different values depending upon different types of fouling: n = 2 for the complete blocking model, n = 1.5 for the standard blocking model, n = 1 represents the intermediate blocking model, and n = 0 indicates the cake layer model. By integrating Eq. (1), four fouling models can be obtained when fixing the value of n (shown in Table S1). It is possible to identify the most likely fouling mechanisms for enzyme immobilization by fitting the experimental flux data using these models and comparing their Adj. R-Square or Residual Sum of Squares. The amount of immobilized enzyme (loading) was calculated from the mass balance equation, and the immobilization efficiency was expressed as enzyme loading efficiency (%): mi  100 (2) Loading efficiency ¼ mt where mi and mt are immobilized and total enzyme amounts, respectively.

Enzymatic reaction Enzymatic membrane reactor (EMR) with free enzymes The amount of enzyme for this series was 100 mL liquid of FDH, 1.0 mg solid of FaldDH, and 1.5 mg solid of ADH, respectively. NADH solution was prepared with 100 mM Tris–HCl buffer, which was pre-bubbled with CO2 for 30 min. 3 mL NADH solution saturated with CO2 was added into the stirred cell equipped with a clean 10 kDa regenerated cellulose membrane in normal mode, and then 1 mL of the enzyme mixture with saturated CO2 was added (final NADH concentration = 50 mM). EMR with immobilized enzymes NADH solution was prepared with 100 mM Tris–HCl buffer, which was pre-bubbled with CO2 for 30 min. 4 mL NADH solution with saturated CO2 was added into the stirred cell equipped with 30 kDa regenerated cellulose membranes in ‘sandwich’ mode (after co- or sequential immobilization). For both EMRs, the applied pressure was controlled manually to ensure that 4 mL permeate was obtained in 30 min. For the enzyme reuse experiment, when 4 mL of permeate was obtained, the filtration was suspended and another 4 mL of fresh NADH solution

Please cite this article in press as: Luo, J. et al.1, Cascade catalysis in membranes with enzyme immobilization for Biotechnol. (2015), http://dx.doi.org/10.1016/j.nbt.2015.02.006

multi-enzymatic conversion

www.elsevier.com/locate/nbt 3 of CO2 to methanol, New

Research Paper

New Biotechnology  Volume 00, Number 00  February 2015

NBT 761 1–9 RESEARCH PAPER

(50 mM) with saturated CO2 was added for the next cycle (each cycle lasted about 30 min).

Analytical methods

Research Paper

The concentration of enzymes was measured as protein concentration using the Bradford protein assay (Perkin Elmer lambda20 UV/VIS, Germany). A Hewlett Packard HP6890 gas chromatograph (GC) equipped with a FID (250 8C) and a Restek XTI-5 column (30 m  0.25 mm i.d., film thickness 0.25 mm) was used for the methanol concentration. The carrier gas was N2 with a flow rate of 0.4 mL min1. The injector temperature was 150 8C and the injection volume was 1 mL. Methanol GC chromatograms were calibrated with 0.01–1 mM methanol solution in 0.1 M pH 7.0 Tris– HCl buffer. Scanning electron microscopy (SEM) was performed in an FEI Helios EBS3 dual beam electron microscope. The skin and support samples were prepared by cutting a small square of the membrane, which was then attached to an aluminum stub by means of double sided sticky carbon tape. The edges of the sample were grounded to the aluminum stub by means of copper tape. Cross sections of the skin and the support were cut from the liquid nitrogen plunged freeze membrane with a pair of scissors. The cross sections were mounted into a slotted specimen stub and grounded with copper tape. All samples were coated with Pt for 2 s at 80 mA in a Cressington 208HR Sputter Coater, which gave an approximate thickness of 4 nm. The micrographs were acquired with the Everhart Thornley detector at low magnifications and the Thru-the-Lens detector at high magnifications, in high vacuum, at 5 keV acceleration voltage and 43 pA current.

Results and discussion First, we performed the reaction using free enzymes (the details are described in the supplementary information). As illustrated in Fig. S1, GDH was used to regenerate NADH [11]. 2 mL of reaction mixture was prepared in a 15 mL centrifuge tube covered with Parafilm and gaseous CO2 was bubbled into solution through a syringe needle. The methanol produced was maintained between 0.5 and 0.7 mM from 30 to 90 min and decreased after 120 min, regardless of whether there was GDH and glutamate for NADH regeneration or not (Fig. S2). This indicated that CO2 reduction by the three enzymes slowed after 30 min (similar results were found by Wang et al. [10]) and some methanol was stripped out of the tube by CO2 [11], resulting in a decline in the methanol concentration at 120 min. Moreover, and unexpectedly, NADH regeneration by GDH did not seem to work in this case. The reduction of NAD+ to NADH by GDH was tested separately at different pH, showing that the catalysis efficiency was greatly affected by pH and the reaction almost stopped at pH 5 (Fig. S3). We found that saturating the buffer with CO2 decreased the pH of the buffer from 7.0 to 5.6  0.1, which is probably why NADH regeneration by GDH was negligible in the multi-enzymatic reaction.

Fouling-induced enzyme immobilization Three enzymes (i.e. FDH, FaldDH, and ADH) were immobilized together or separately in 30 kDa ultrafiltration membranes by the fouling formation technique, as illustrated in Fig. 1. The membrane consists of a regenerated cellulose skin layer and polypropylene nonwoven support with an effective area of 13.4 cm2. Based on a previous study [27], the membrane support layer 4

New Biotechnology  Volume 00, Number 00  February 2015

was positioned to face the feed in order to increase enzyme loading and stability, and the skin layer was covered by an extra polypropylene support to alleviate membrane compression and prevent peeling off. The permeate volume over time is presented in Fig. 2. Using the fouling models (Table S1), the predominant fouling mechanism can be determined by Adj. R-Square or Residual Sum of Squares [28,29]. It was found that the cake layer model best described the data obtained, regardless of the enzyme type and concentration. As seen in Fig. 2, during the initial filtration stage only the cake layer model fits the experimental data because only the skin layer is considered as the membrane and most of the enzymes deposit on the membrane; as the filtration time increases the other models come closer and closer to the experimental data as the fouling layer is also regarded as an additional ‘membrane’, and thus the enzyme-deposited enzyme interactions are misestimated as a pore blocking phenomenon. That is why the differences of R-Square for all the models are not as obvious as expected. The micrographs in Fig. 3 show both the support side and cross-sections of the membranes before and after co-immobilization. Some enzyme aggregates adsorbed on the surface of the polypropylene support fibers were observed (Figs. 3c and S4) and the regenerated cellulose sublayer became less porous after immobilization (Fig. 3e and f). According to the results from mathematical modeling and microscopy, we concluded that most enzymes are rejected by the skin layer and entrapped in the sublayer [30], forming a cake layer with a cellulose skeleton, while some enzymes bind on the support fibers by hydrophobic adsorption [31]. Accordingly, we believe that this fouling-induced enzyme immobilization involves at least two mechanisms: entrapment and adsorption. After enzyme immobilization, the membrane permeabilities dropped by more than 90% (339  6 L m2 h1 bar1 for new membranes) and the enzyme loading efficiency was 65–90% (Fig. 4). 2.27  0.23 mg proteins were immobilized in one membrane (co-immobilization) and 2.76  0.06 mg proteins were immobilized in the tri-membranes (sequential immobilization).

Enzymatic reaction in EMRs 4 mL of NADH solution with saturated CO2 was added into the Amicon cell equipped with the membranes ‘fouled’ by enzymes. Pressure was controlled manually to ensure that 4 mL of permeate was obtained in 30 min. As seen in Fig. 5a, the methanol concentration increased with NADH concentration, being consistent with previous reports [5,8,11,12]. However, the results were similar to the free, co- and sequentially immobilized enzyme systems in the present work. Therefore, we believe that enzyme activity is retained by non-covalent immobilization and the flow-through mode can effectively avoid enzyme leakage during the reaction. The expected improvement of enzyme stability due to immobilization did not result in any advantage for methanol production over 30 min. Fig. 5b shows the operational stability of the free and immobilized enzymes in a batch mode (fresh substrate was fed for each cycle). Free enzymes were tested in a membrane reactor equipped with a 10 kDa regenerated cellulose membrane in normal mode [18]. Surprisingly, the methanol produced by free enzymes was maintained at 0.10–0.15 mM for ten cycles, indicating that the enzymes were not fully inactivated for at least 5 h. For immobilized enzymes, methanol production increased from 0.10 to 0.25 mM in the first five cycles, and then gradually decreased.

www.elsevier.com/locate/nbt Please cite this article in press as: Luo, J. et al.1, Cascade catalysis in membranes with enzyme immobilization for Biotechnol. (2015), http://dx.doi.org/10.1016/j.nbt.2015.02.006

multi-enzymatic conversion

of CO2 to methanol, New

NBT 761 1–9 RESEARCH PAPER

Research Paper

New Biotechnology  Volume 00, Number 00  February 2015

FIGURE 2

Evaluation of the predominant membrane fouling mechanisms at a constant pressure of 2 bar during (a) co-immobilization and (b)–(d) sequential immobilization.

The initial methanol increase was presumably caused by the NADH retention or adsorption by the ‘fouling layer’ (NADH accumulation in ‘fouled’ membranes). Then part of the NADH might be replaced by NAD+ in the last five cycles. However, for a clean 10 kDa membrane, NADH and NAD+ (molecular weight <1 kDa) were easy to pass through. In principle, the co-immobilization system could accomplish catalysis in a highly efficient way because the intermediates are consumed in situ by the enzymes (i.e. low product inhibition) [32]. However, compared with sequential immobilization (Fig. 1), the methanol yield was not improved by co-immobilization (Fig. 5). There are three possible reasons for this unexpected result. Firstly, the immediate consumption of intermediates by the subsequent enzymes results in a constant low substrate concentration for the reactions, and the trade-off between the mitigation of product inhibition and low substrate concentration for the adjacent enzymes may make co-immobilization meaningless. Secondly, the pressure-driven mass transfer across membranes overcame the diffusion barrier between the membranes (the distance between two enzyme layers was at least 230 mm, the thickness of one membrane), and the intermediates could rapidly reach the targeted enzymes in the sequential immobilization system. The third possible reason involves the enzymatic reaction kinetics. Rusching et al. reported that formate oxidation was 30 times faster than CO2 reduction catalyzed by FDH, and thus the first step is likely to be a bottleneck for the reduction of CO2 [3]. But if formic acid from the

first reaction could be reduced into formaldehyde by FaldDH rapidly, these sequential reactions would go forward because the final step catalyzed by ADH was proven to be highly efficient with low product inhibition [18,33]. If this hypothesis was true, the overall catalytic efficiencies toward the desired pathway (i.e. Fig. S1) would also be largely promoted by in situ NADH regeneration. However, the methanol concentration was still quite low even with regeneration of NADH [11,12].

Reaction mechanism and kinetics In order to unravel this multi-enzymatic reaction, three enzymes were tested separately with corresponding substrates in a cuvette. Saturated CO2 was used to standardize the reaction pH for all the tests. The absorbance of NADH with substrates was stable before adding enzyme, and when the enzyme was injected into the cuvette there was a rapid drop of absorbance for all the cases due to the dilution effect. After this drop, the absorbance at 340 nm decreased which indicated that the NADH was consumed with time and a reaction happened. As shown in Fig. 6a, the FDH indeed catalyzed the hydrogenation of CO2 (NADH was consumed slowly) while the rate of NADH consumption was the same with the three enzymes, indicating that the cascade reactions did not proceed. This was confirmed by the results in Fig. 6b, where FaldDH could not catalyze the production of formaldehyde until the formic acid concentration increased to 10 mM. In this case, the pH decreased to 4.2 due to the addition of formic acid. In Fig. 6c,

Please cite this article in press as: Luo, J. et al.1, Cascade catalysis in membranes with enzyme immobilization for Biotechnol. (2015), http://dx.doi.org/10.1016/j.nbt.2015.02.006

multi-enzymatic conversion

www.elsevier.com/locate/nbt 5 of CO2 to methanol, New

NBT 761 1–9 RESEARCH PAPER

New Biotechnology  Volume 00, Number 00  February 2015

Research Paper FIGURE 3

(a) Overview of support layer (polypropylene nonwoven fibers); (b) surface of support fiber before immobilization and (c) after immobilization; (d) overview of membrane cross-section; (e) regenerated cellulose skin layer and sublayer before immobilization and (f) after immobilization.

the reduction of formaldehyde by ADH is shown to be highly efficient. It is well known that these enzymes can catalyze the oxidation of methanol to CO2 when the electron acceptor is provided [14] and thus these three reactions are reversible. In order to quantify the reaction efficiency in each step, the enzyme kinetics for three enzymes (both forward and reverse reactions) were measured. The measurements were carried out at pH 7 to provide general

information. The CO2 concentration could not be controlled and monitored due to device limitations and the kinetics data for the first reaction (CO2 ! formic acid) were obtained from Ruschig et al. [3]. For the reaction from formic acid to formaldehyde catalyzed by FaldDH, no obvious reaction happened when the pH was higher than 6.0, and the NADH was not stable when the pH was lower than 4.0 (data not shown). Thus, it was very difficult to measure the reaction velocities at different formic acid concentrations with

FIGURE 4

(a) Permeate flux of membranes after enzyme immobilization at 2 bar and (b) enzyme loading and loading efficiency for different approaches. 6

www.elsevier.com/locate/nbt Please cite this article in press as: Luo, J. et al.1, Cascade catalysis in membranes with enzyme immobilization for Biotechnol. (2015), http://dx.doi.org/10.1016/j.nbt.2015.02.006

multi-enzymatic conversion

of CO2 to methanol, New

NBT 761 1–9 New Biotechnology  Volume 00, Number 00  February 2015

RESEARCH PAPER

0.5

1.0

0.8

0.6

0.4

0.2

0.0

b 0.4

0.3

0.2

0.1

0.0

0

10

20

30

40

50

Free enzymes Co-immobilized enzymes Sequential immobilized enzymes

1

2

NADH concentration (mM)

3

4

5

6

7

8

9

10

Number of reuse cylces

FIGURE 5

(a) Methanol production at different NADH concentrations with free and immobilized enzymes and (b) with recycling and reusing of free and immobilized enzymes (NADH = 10 mM). Reaction time = 30 min. Enzyme and NADH concentrations were same for free and immobilized enzyme systems; in order to reuse free enzymes, an enzymatic membrane reactor equipped with 10 kDa regenerated cellulose membrane (skin layer facing feed) was used.

constant pH. For example, at pH 5.7, the reaction velocities (very slow) were the same at the formate concentrations from 10 to 100 mM (data not shown). It seems that there is a dilemma in this reaction, that the formic acid (pKa 3.77) is an effective substrate

while its cofactor (NADH) is not stable in acidic environments (according to the product information from Sigma). As shown in Table 1, for the first enzyme FDH, the efficiency of the reverse reaction (formic acid ! CO2) was much higher than the forward

FIGURE 6

Effect of substrates and enzymes on the reactions concerning CO2 conversion. Buffer pH after CO2 bubbling = 5.6  0.1; 2.5 mL buffer in cuvette; substrate addition: 10, 50, 200 mL 133 mM formic acid and formaldehyde; enzyme addition: FDH = 50 mL (0.14 U, 2.5 mg liquid); FaldDH = 50 mL (0.15 U, 50 mg solid); ADH = 50 mL (>15 U, 5 mg solid), mixture = 100 mL. Standard deviations are less than 5%.

Please cite this article in press as: Luo, J. et al.1, Cascade catalysis in membranes with enzyme immobilization for Biotechnol. (2015), http://dx.doi.org/10.1016/j.nbt.2015.02.006

multi-enzymatic conversion

www.elsevier.com/locate/nbt 7 of CO2 to methanol, New

Research Paper

Free enzymes Co-immobilized enzymes Sequential immobilized enzymes

Methanol concentration (mM)

Methanol concentration (mM)

a

NBT 761 1–9 RESEARCH PAPER

New Biotechnology  Volume 00, Number 00  February 2015

TABLE 1

Kinetic parameters of enzymes for different reactions. Km (mM)

Vmax (mM min1)

CO2 ! HCOOHa

30–50

0.002

HCOOH ! CO2

3.3

0.02

Enzyme/reaction Formate dehydrogenase

b

Formaldehyde dehydrogenase Alcohol dehydrogenase

HCOOH ! HCHO

NA

NA

HCHO ! HCOOH

0.06

0.01

HCHO ! CH3OH

17.5

0.3

CH3OH ! HCHO

275

0.5  103

+

Research Paper

Tris buffer with pH 7; temperature = 23  1 8C; NADH/NAD = 0.25 mM; 2 mL substrate; enzyme addition: FDH = 50 mL (0.14 U, 2.5 mg liquid); FaldDH = 50 mL (0.3 U, 100 mg solid); ADH = 50 mL (>150 U, 50 mg solid). The kinetic parameters were calculated from the results in Fig. S5. Standard deviations are less than 5%. a Data from Ruschig et al. (1976), CO2 concentration was from 0 to 15 mM, temperature = 20 8C, pH = 6.2.b No obvious reaction at low formic acid concentrations (0.1–2.5 mM), and pH was significantly decreased when formic acid concentration was more than 5.0 mM (NADH was not stable when pH was lower than 4).

one (CO2 2 ! formic acid), while the latter was one of the desirable reaction in our study. For the second enzyme FaldDH, the reverse reaction (formaldehyde2 ! formic acid) was quite efficient (Km was lowest in these reactions), while the forward reaction (formic acid2 ! formaldehyde) was in an unfavorable situation. For the third enzyme ADH, the forward reaction (formaldehyde2 ! methanol) was much more efficient than the reverse one (methanol2 ! formaldehyde). Therefore, during the multi-enzymatic conversion of CO2 to methanol, although the first reaction happens very slowly, the second reaction catalyzed by FaldDH is the real bottleneck. The production of formaldehyde was delayed by a slow accumulation of formic acid from the first reaction (the second reaction required a threshold concentration of formic acid to be activated). That may be why the co-immobilization of the three enzymes cannot improve the methanol yield compared to sequential immobilization (Fig. 5). This also explains why a high NADH concentration is necessary for the reaction. Wang et al. also found that the whole reaction velocity fluctuated with time (first a decrease, then an increase, and then a decrease again), which could partly confirm our hypothesis that the FaldDH was ‘waiting’’ for formic acid to accumulate [10]. Cazelles et al. reported that even after increasing the amount of FaldDH, the production of formaldehyde from CO2 was less than 0.1 mM, while using formate with a high concentration (100 mM) as substrate, the production of methanol by FaldDH and ADH was more than 0.5 mM [12].

Acknowledgement

Conclusion

We thank The Hans Christian Ørsted Postdoc Program (DTU) for financial support (J. Luo).

We have evaluated the multi-enzymatic conversion of CO2 to methanol by co- and sequential immobilizations of enzymes in polymeric membranes, which were achieved by fostering membrane fouling by a simple filtration method. Enzymes were immobilized in the membrane as a cake layer in the cellulose sublayer or

as aggregates on polypropylene fibers of the support layer, which fully retain their activity by binding non-covalently. Co-immobilization did not improve methanol production, compared with sequential immobilization, because of the trade-off between the mitigation of product inhibition and low substrate concentration for the adjacent enzymes, and because the second enzyme (FaldDH) could not effectively consume the intermediate (formic acid) from first reaction catalyzed by FDH. The second reaction catalyzed by FaldDH is in an unfavorable situation because there is a chemistry conflict between the required substrates (formic acid and NADH) and this reaction is sensitive to the substrate/product concentration and pH. Thus sequential immobilization can be used for this case as it allows the reaction conditions for each step to be adjusted separately and the pressure-driven mass transfer can overcome the diffusion resistance between enzymes. Extremely high concentrations of NADH can force the reaction toward the desired pathway, but this is probably not economically feasible. In situ regeneration of NADH is an alternative but it suffers from high product inhibition and the low pH induced by solubilized CO2. Engineering FaldDH by targeted mutation, and finding another enzyme or cofactor favoring the reduction of formic acid to formaldehyde are promising alternatives to improve methanol production by this multi-enzymatic catalysis.

Appendix A. Supplementary data Supplementary data associated with this article can be found, in the online version, at http://dx.doi.org/10.1016/j.nbt.2015.02. 006.

References [1] Qiao J, Liu Y, Hong F, Zhang J. A review of catalysts for the electroreduction of carbon dioxide to produce low-carbon fuels. Chem Soc Rev 2014;43:631–75. [2] Kondratenko EV, Mul G, Baltrusaitis J, Larrazabal GO, Perez-Ramirez J. Status and perspectives of CO2 conversion into fuels and chemicals by catalytic, photocatalytic and electrocatalytic processes. Energy Environ Sci 2013;6:3112–35. ¨ ller U, Willnow P, Ho ¨ pner T. CO2 reduction to formate by NADH [3] Ruschig U, Mu catalysed by formate dehydrogenase from Pseudomonas oxalaticus. Eur J Biochem 1976;70:325–30. [4] Kuwabata S, Tsuda R, Yoneyama H. Electrochemical conversion of carbon dioxide to methanol with the assistance of formate dehydrogenase and methanol dehydrogenase as biocatalysts. J Am Chem Soc 1994;116:5437–43.

8

[5] Obert R, Dave BC. Enzymatic conversion of carbon dioxide to methanol: enhanced methanol production in silica solgel matrices. J Am Chem Soc 1999;121:12192–93. ¨ ller V. Direct and reversible hydrogenation of CO2 to [6] Schuchmann K, Mu formate by a bacterial carbon dioxide reductase. Science 2013;342:1382–5. [7] Baskaya FS, Zhao X, Flickinger M, Wang P. Thermodynamic feasibility of enzymatic reduction of carbon dioxide to methanol. Appl Biochem Biotechnol 2010;162:391–8. [8] Sun Q, Jiang Y, Jiang Z, Zhang L, Sun X, Li J. Green and efficient conversion of CO2 to methanol by biomimetic coimmobilization of three dehydrogenases in protamine-templated titania. Ind Eng Chem Res 2009;48:4210–5.

www.elsevier.com/locate/nbt Please cite this article in press as: Luo, J. et al.1, Cascade catalysis in membranes with enzyme immobilization for Biotechnol. (2015), http://dx.doi.org/10.1016/j.nbt.2015.02.006

multi-enzymatic conversion

of CO2 to methanol, New

NBT 761 1–9

[9] Shi J, Wang X, Jiang Z, Liang Y, Zhu Y, Zhang C. Constructing spatially separated multienzyme system through bioadhesion-assisted bio-inspired mineralization for efficient carbon dioxide conversion. Bioresour Technol 2012;118:359–66. [10] Wang X, Li Z, Shi J, Wu H, Jiang Z, Zhang W, et al. Bioinspired approach to multienzyme cascade system construction for efficient carbon dioxide reduction. ACS Catal 2014;4:962–72. [11] El-Zahab B, Donnelly D, Wang P. Particle-tethered NADH for production of methanol from CO2 catalyzed by coimmobilized enzymes. Biotechnol Bioeng 2008;99:508–14. [12] Cazelles R, Drone J, Fajula F, Ersen O, Moldovan S, Galarneau A. Reduction of CO2 to methanol by a polyenzymatic system encapsulated in phospholipidssilica nanocapsules. New J Chem 2013;37:3721–30. [13] Zhao F, Li H, Jiang Y, Wang X, Mu X. Co-immobilization of multi-enzyme on control-reduced graphene oxide by non-covalent bonds: an artificial biocatalytic system for the one-pot production of gluconic acid from starch. Green Chem 2014;16:2558–65. [14] Liu F, Banta S, Chen W. Functional assembly of a multi-enzyme methanol oxidation cascade on a surface-displayed trifunctional scaffold for enhanced NADH production. Chem Commun 2013;49:3766–8. [15] Sassolas A, Blum LJ, Leca-Bouvier BD. Immobilization strategies to develop enzymatic biosensors. Biotechnol Adv 2012;30:489–511. [16] Jochems P, Satyawali Y, Diels L, Dejonghe W. Enzyme immobilization on/in polymeric membranes: status, challenges and perspectives in biocatalytic membrane reactors (BMRs). Green Chem 2011;13:1609–23. [17] Giorno L, Drioli E. Biocatalytic membrane reactors: applications and perspectives. Trends Biotechnol 2000;18:339–49. [18] Luo J, Meyer AS, Jonsson G, Pinelo M. Fouling-induced enzyme immobilization for membrane reactors. Bioresour Technol 2013;147:260–8. [19] Luo J, Marpani F, Brites R, Frederiksen L, Meyer AS, Jonsson G, et al. Directing filtration to optimize enzyme immobilization in reactive membranes. J Membr Sci 2014;459:1–11. [20] Giorno L, Drioli E, Carvoli G, Cassano A, Donato L. Study of an enzyme membrane reactor with immobilized fumarase for production of L-malic acid. Biotechnol Bioeng 2001;72:77–84. [21] Hadidi M, Zydney AL. Fouling behavior of zwitterionic membranes: impact of electrostatic and hydrophobic interactions. J Membr Sci 2014;452:97–103.

RESEARCH PAPER

[22] Kozlovskaya V, Ok S, Sousa A, Libera M, Sukhishvili SA. Hydrogen-bonded polymer capsules formed by layer-by-layer self-assembly. Macromolecules 2003;36:8590–2. ¨ Tte H, Flossdorf J, Sahm H, Kula M-R. Purification and properties of [23] SchU formaldehyde dehydrogenase and formate dehydrogenase from Candida boidinii. Eur J Biochem 1976;62:151–60. [24] Ando M, Yoshtmoto T, Ogushl S, Rikitake K, Shibata S, Tsuru D. Formaldehyde dehydrogenase from pseudomonas putida: purification and some properties. J Biochem 1979;85:1165–72. [25] Reisler E, Pouyet J, Eisenberg H. Molecular weights, association, and frictional resistance of bovine liver glutamate dehydrogenase at low concentrations. Equilibrium and velocity sedmintation, light-scattering studies, and settling experiments with macroscopic models of the enzyme oligomer. Biochemistry 1970;9:3095–102. [26] Hermia J. Constant pressure blocking filtration laws—application to power-law non-newtonian fluids. Trans Inst Chem Eng 1982;60:183–7. [27] Luo J, Meyer AS, Jonsson G, Pinelo M. Enzyme immobilization by fouling in ultrafiltration membranes: impact of membrane configuration and type on flux behavior and biocatalytic conversion efficacy. Biochem Eng J 2014;83:79–89. [28] Affandy A, Keshavarz-Moore E, Versteeg HK. Application of filtration blocking models to describe fouling and transmission of large plasmids DNA in sterile filtration. J Membr Sci 2013;437:150–9. [29] Palacio L, Ho C-C, Zydney AL. Application of a pore-blockage—cake-filtration model to protein fouling during microfiltration. Biotechnol Bioeng 2002;79:260–70. [30] Mazzuca S, Giorno L, Spadafora A, Mazzei R, Drioli E. Immunolocalization of bglucosidase immobilized within polysulphone capillary membrane and evaluation of its activity in situ. J Membr Sci 2006;285:152–8. [31] Jim KJ, Fane AG, Fell CJD, Joy DC. Fouling mechanisms of membranes during protein ultrafiltration. J Membr Sci 1992;68:79–91. [32] Jia F, Narasimhan B, Mallapragada S. Materials-based strategies for multi-enzyme immobilization and co-localization: a review. Biotechnol Bioeng 2014;111:209–22. [33] Wen N, Liu W, Hou Y, Zhao Z. The kinetics behavior of the reduction of formaldehyde catalyzed by alcohol dehydrogenase (ADH) and partial uncompetitive substrate inhibition by NADH. Appl Biochem Biotechnol 2013;170:370–80.

Please cite this article in press as: Luo, J. et al.1, Cascade catalysis in membranes with enzyme immobilization for Biotechnol. (2015), http://dx.doi.org/10.1016/j.nbt.2015.02.006

multi-enzymatic conversion

www.elsevier.com/locate/nbt 9 of CO2 to methanol, New

Research Paper

New Biotechnology  Volume 00, Number 00  February 2015