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Article

Two Cascade Reactions with Oleate Hydratases for the Sustainable Biosynthesis of Fatty Acid-Derived Fine Chemicals

by
Sophia A. Prem
,
Martina Haack
,
Felix Melcher
,
Marion Ringel
,
Daniel Garbe
and
Thomas Brück
*
Werner Siemens-Chair of Synthetic Biotechnology, School of Natural Sciences, Technical University of Munich, Lichtenbergstr. 4, 85748 Garching, Germany
*
Author to whom correspondence should be addressed.
Catalysts 2023, 13(9), 1236; https://doi.org/10.3390/catal13091236
Submission received: 11 July 2023 / Revised: 26 July 2023 / Accepted: 8 August 2023 / Published: 25 August 2023
(This article belongs to the Section Biocatalysis)
Browse Figures
Review Reports Versions Notes

Abstract

:
Oleate hydratases (OHs) are of significant industrial interest for the sustainable generation of valuable fine chemicals. When combined with other enzymes in multi-step cascades, the direct formation of fatty acid congeners can be accomplished with minimal processing steps. In this study, two cascade reactions are presented, which can be applied in one-pot approaches. The first cascade was placed “upstream” of an OH derived from Rhodococcus erythropolis (OhyRe), where a lipase from Candida rugosa was applied to hydrolyze triglycerides into free fatty acids, a crucial step for OH conversion. Further, we tested the lipase–OhyRe cascade with various types of renewable triglycerides of plant and microbial origin. In this context, the most efficient conversion was observed for microbial oil from Cutaneotrichosporon oleaginosus leading the way toward its industrial application. In contrast, the second cascade was placed “downstream” of OhyRe, where a novel secondary alcohol dehydrogenase (secADH) was applied to oxidize the hydroxylated fatty acid into a fatty acid ketone. Optimal reaction parameters for the cascade with the secADH were established, which allows this to be applied to high-throughput screens. Moreover, we describe a light-dependent route, thereby extending the catalytic efficiency of the OH enzyme system.

Graphical Abstract

1. Introduction

In nature, metabolic pathways are realized by cascading enzyme-driven reactions within cells, ensuring cell growth and viability. The concept of these cellular cascade reactions can also be transferred into extracellular matrices, which allows targeted reactions to be more easily controlled. Further, fewer side-products are generated [1], and simplified downstream product recovery is achieved compared to complex systems present in a microbial cell factory [2]. The application of enzyme cascade reactions has enabled the formation of the sustainable in vitro biosynthesis of fine chemicals and the advent of efficient screening methods in the fields of biotechnology, enzymology, and other related areas. In this context, one-pot enzyme reaction cascades combined with simplified cofactor regeneration systems, the application of enzyme co-immobilization, scaffolding, or encapsulation leads to the targeted design of effective chemical production systems [3].
The advantages of in vitro enzyme cascades for the industrial biosynthesis of target molecules are reduced downstream processes, leading to lower production costs, and in the case of enzyme recycling, less product and catalyst extraction and purification steps, enabling higher space-time yields given the optimized conversion parameters. Furthermore, less organic solvent and energy consumption are needed, making the process more sustainable. Lastly, reduced end- or side-product inhibition occurs, which can be the case for whole-cell biocatalysts containing enzyme cascades [4,5].
In general, there are several routes to how cascade enzyme reactions can be applied. This might be either by the application of enzymes originating from different organisms in the form of synthetic cascades or the extraction of naturally occurring catalysts contained within an existing metabolic pathway. However, the non-natural enzyme coupling efforts might lead to severe incompatibilities regarding reaction parameters such as side reactions, varying temperature or pH profiles, substrate access, and solubility issues, as well as substrate promiscuity, leading to side reactions [2]. Consequently, in most cases, intensive optimization efforts are required to achieve satisfying reaction outputs, and usually trade-offs are to be expected when applying non-optimal parameters for each biocatalyst component.
So far, enzyme cascade reactions have been applied in industrial processes to generate platform chemicals, such as chiral alcohols [6], and are considered for the production of complex pharmaceutical molecules [7], thereby showcasing the immanent relevance of in vitro catalytic systems.
Additionally, enzyme cascades can be applied to design high-throughput assays as they allow for the rapid and facile screening of biological targets. With regard to the former, a plethora of variations regarding the output signals are available, of which the most common ones are spectrophotometric, including but not limited to absorbance [8,9,10], fluorescence [11,12], and bioluminescence [13,14] signals.
At present, there have been only minor efforts to develop relevant reaction cascades revolving around oleate hydratases (OH), which predominantly convert free oleic acid into 10-(R)-hydroxy stearic acid (10-HSA), a fatty acid-derived fine chemical in the lubricant and cosmetic industry [15]. Naturally, this class of enzymes is considered to be involved in defense measures against free fatty acids, which are reported to be toxic for many microorganisms. Consequently, OHs are not able to convert fatty acids with a protected carboxy group, such as methyl esters or acyl glycerides.
Besides their substrates, OHs exhibit the binding of FAD as a cofactor during catalysis. This cofactor, despite not participating in a redox reaction, has been proposed to have structural significance during the catalytic process [16]. Additionally, it has been observed that an environment with reducing conditions leads to a significant enhancement in activity. This finding suggests that FADH2 is likely the preferred cofactor in the catalytic process. Upon cell lysis, FADH2 is oxidized to FAD, and in many OHs, only low FAD occupancy can be measured after protein purification, thus making the direct proof of FADH2 binding preferences considerably challenging. For instance, OhyRe [17], OH from Pediococcus parvulus (OhyPp) [18], and a linoleate hydratase (LAH) [19] have been observed to fully lose FAD upon purification [18,19].
One of the main goals for the sustainable biosynthesis of 10-HSA is the use of plant and microbial as well as waste oils. The OH substrate oleic acid is the main fatty acid component of many plant and microbial oils, such as high-oleic sunflower oil (HOSO) or yeast triglycerides sourced from the oleaginous yeast Cutaneotrichosporon oleaginosus [20]. The latter provides a new, sustainable substrate for OH-centered reactions due to higher yields and reduced land use compared to conventional plant-based resources. Additionally, the fatty acid profile of C. oleaginosus can be altered by adapting fermentation and media parameters [21,22]. Furthermore, a novel genetic engineering process paves the way for tailored fatty acid profiles, thus prospectively allowing for the efficient production of sustainably sourced 10-HSA [23].
Currently, the analogue 12-HSA is solely produced from castor oil via hydrogenation under extreme pressure and temperature, leading to exposure to fluctuations in quality and demand. To this end, plant and microbial oils with high oleic acid content are more diversely available on the market, and biocatalysts convert under moderate conditions with low energy demand. However, to date, the low stability and performance of the available biocatalysts, resulting in rather low yields, have led to high prices compared to conventional production methods. This is where fewer processing steps are advantageous, resulting in time and material savings and ultimately enhancing the economic viability. 10-HSA, furthermore, is a solid material, and thus, no in-between liquid–liquid separation steps are required. The utilization of customized enzyme cascade reactions has the potential to steer closer to achieving this objective.
Several OHs with varying properties are already described and characterized [17,18,24,25], however, monomeric OHs exhibit the greatest potential for an industrial application, as subunit dissociation can be neglected and immobilization strategies can be adopted easier [26]. Hence, this study focuses on an OH from Rhodococcus erythropolis (OhyRe) as the first identified member of monomeric OHs.
A variety of cascades centered around OHs are reported by Hagedoorn et al. [27], and they would significantly broaden the product and application portfolio. This can, for instance, be achieved by applying different enzymes such as alcohol dehydrogenases, amine transaminase, or lipases, exploiting their esterification capabilities.
For the cascade reaction of 10-HSA into its oxidized product, called 10-keto stearic acid (10-KSA), an alcohol dehydrogenase (ADH) is required, which exhibits substrate preference for longer carbon chain alcohols over short ones. To date, several organisms, such as Rhodococcus erythropolis [28] and Nocardia cholesterolicum NRRL 5767 [29], are reported to convert 10-HSA into 10-KSA, and an isolated ADH from Micrococcus luteus has already been applied in screenings [30].
In this study, we first optimized OhyRe’s conversion efficiency by using broadband white light and subsequently applied two different cascade reactions involving OHs. These reactions are of relevance for the one-pot production of 10-HSA from triglyceride oils and 10-keto stearic acid (10-KSA). The latter reaction might additionally be useful for screening purposes due to its absorbance response.
Since some OHs [24], as well as OhyRe [17], have low turnover numbers, they were considered to be the limiting components in the enzyme cascade reactions. Consequently, prior optimization was required to achieve appropriate outputs, which is why the effect of light on the conversion of OhyRe was investigated.
Next, the first cascade was investigated as depicted in Figure 1a, which is the combination of OhyRe with a lipase from Candida rugosa for the direct conversion of triglyceride oils into 10-HSA without the need for the prior hydrolysis of oils into free fatty acids. Lipases are widely studied enzymes [31], and thus, there are several reports about their application in cascade reactions [32,33,34]. However, to our knowledge, there is no report demonstrating the combination of lipase and OHs in vitro.
The second cascade reaction, as depicted in Figure 1b, comprises the conversion of oleic acid into 10-HSA, followed by the biosynthesis of 10-KSA using a novel secondary alcohol dehydrogenase. Furthermore, the reaction parameters were optimized for this cascade reaction to be able to produce 10-KSA as a fine chemical, and secondly, also to prospectively use it in high-throughput screening assays. Despite an uncharacterized secondary ADH being applied already in a high-throughput screen [30], a detailed characterization of the reaction parameters has been lacking so far, which is required for an adequate application.

2. Results and Discussion

2.1. Optimization of OhyRe Using Light

Before applying the above-mentioned cascade reactions, it was necessary to maximize OH’s activity due to the observed slow turnovers. It was reported that a reducing environment strongly enhances OH’s activity, which before has been achieved in vitro by applying elaborate means, such as employing a cascade involving a flavin oxidoreductase [35], by photobleaching, or with the addition of DTT under anaerobic conditions, as described by Engleder et al. [16].
For this study, a more rapid method for the generation of a reducing environment was needed, and this was achieved by using a light source, where the FAD can be partially reduced without the addition of chemicals and without maintaining an anaerobic environment. Since OhyRe loses most FAD upon purification, an equimolar concentration of FAD was added. The effect of light on the OhyRe conversion was tested, and indeed, it could be observed that the addition of broadband white light during the reaction led to a 40-fold increase in activity compared to when the reaction was conducted in the dark, as depicted in Figure 2a.
Also, specific light colors were tested, and blue light showed a similar positive effect, leading to the conclusion that blue light is indeed responsible for the improved activity of OhyRe. Blue light is known to act as an activator for blue-light-sensing proteins via the excitation of FAD [36,37]. Most likely the reduction of FAD, leading to FADH2, the preferred cofactor of OhyRe, is responsible for the enhanced activity. Next, the optimal light intensity was tested, and a broad optimal range between 243 and 364 µmol/m2·s was observed (Figure 2b). Based on the outcomes of these experiments, the subsequent reactions were conducted using broadband white light at an intensity of 364 µmol/m2·s.
OHs are likely the limiting enzymes in coupled reactions due to their low turnover numbers, making them the primary focus in optimization efforts. Therefore, the improvement of activity through blue and broadband white light illumination is a first step towards the optimization of OhyRe’s catalytic activity within cascade reactions.

2.2. A Cascade Reaction Using a Lipase and OhyRe

Next, by applying the broadband light activation, a cascade reaction using a lipase and OhyRe, the conversion of sustainable microbial and plant oils with varying oleic acid concentrations was analyzed. Commercially available olive, linseed, and rapeseed oils were applied and compared to high-oleic sunflower oil (HOSO) and triglyceride oil from C. oleaginosus. For the first step of the cascade reaction, the commercially available lipase from Candida rugosa was used since it is known as a high-fidelity enzyme for the hydrolysis of triglycerides into free fatty acids and glycerol and is also active over a wide temperature and pH range [38,39]. Additionally, a recent screening experiment showed that Candida rugosa lipase was most efficient for rapeseed oil and HOSO conversion [40].
It has to be considered that oils contain small amounts of free fatty acids originating from the processing and storage conditions [41]. To evaluate whether OhyRe can convert any residual free fatty acids from crude oil, and thus, to determine the background activity, first, the conversion of the oil was tested with OhyRe but without lipase. However, as desired, no production of 10-HSA was observed, thereby pointing towards the fact that the amount of free fatty acids in the tested oils can be neglected.
Instead, when the C. rugosa lipase was added in combination with OhyRe, conversion at different yields was observed depending on the type of oil used. The highest yield was achieved for rapeseed oil, with a conversion of 35% of oleic acid, and the least for linseed oil, with 6% (Figure 3a), which was expected since it also contains the lowest amount of oleic acid amongst all oils tested (Table 1).
To evaluate the effect of the oleic acid content, we calculated the converted 10-HSA per oleic acid content in each respective oil and observed that now the microbial oil from C. oleaginosus showed the highest conversion, while olive and HOSO showed the lowest conversion (Figure 3b). Strikingly, the performance of HOSO was one of the lowest regarding the converted 10-HSA per oleic acid content of oils. In general, there was a decrease in yield with increasing oleic acid content, which might be a sign of substrate inhibition. Also, the types of oils were processed differently, and the remaining ingredients most likely can influence the enzymes’ activities. However, the used HOSO is for for technical applications, and thus, well purified, leading to the depletion of contaminants, yet it still showed low yields. Additionally, the composition of the triglycerides might affect the yield due to the preferences of the lipase [42,43]. Linseed oil contains large amounts of C18:2 and C18:3, which are also substrates for OhyRe [17] and other OHs [18], and thus, could bind and might lead to side reactions, resulting in a lower yield, whereas in C. oleaginosus oil, the most abundant fatty acid after oleic acid is C16:0 (Table 1). Thus, this oil might be beneficial regarding the oleic acid content relative to other C18 species. Recently, our research group successfully utilized CRISPR/Cas to genetically engineer C. oleaginosus, resulting in oil without any polyunsaturated fatty acids [23]. For upcoming experiments, it will be intriguing to investigate if the conversion yield can be further improved using this type of oil.
For the assay applied here, the oil was emulsified with buffer. Even though lipases prefer an aqueous–oil bilayer [44] as opposed to emulsified oil, which is the exact opposite of OHs strictly requiring emulsified oil, there was up to 35% of conversion. Still, further optimizations can be applied to tackle the tradeoffs of the two varying specifications of lipases and OHs regarding the emulsification of oil. This can, for instance, be achieved by testing the effect of surfactants or by applying a two-step conversion with an intermediate emulsification step.
In summary, the conducted experiments show that the lipase from Candida rugosa and OhyRe can indeed be used in a cascade reaction, thus allowing for the efficient conversion of triacyl glycerides-based oils into 10-HSA. However, the conversion efficiency is largely dependent on the used oil substrate and there is room for optimization to further increase the yield.

2.3. Functional Expression of secADH

Subsequently, we opted for the implementation of the second step in the aimed downstream cascade reaction involving an alcohol dehydrogenase to convert 10-HSA into 10-KSA (cf. Figure 1b). Apart from the large-scale production of this potential fine chemical for further downstream reactions, it can be used for high-throughput screening purposes. An alcohol dehydrogenase from M. luteus was already reported to be active on secondary alcohol substrates [45]. However, to expand the toolbox for this reaction, we decided to further search the NCBI database for genes that are related to the one from M. luteus and are also predicted to be L-3-hydroxyacyl-CoA dehydrogenases.
An ADH from Deinococcus radiodurans was identified (NCBI Reference Sequence: WP_027479748.1), which is an extremophile organism, and thus, its proteins are expected to hold superior properties regarding their optimal range of pH and temperature and overall durability and stability [46]. The gene was codon optimized for E. coli and cloned into pet28a(+). SecADH could be expressed in soluble form (Figure S1), which is predicted to be a homo-dimer [47], with one monomer being 32.4 kDa large [48].

2.4. Substrate Spectrum of secADH from Deinococcus radiodurans

To test whether the newly identified secADH was suitable for the cascade together with OHs, the substrate spectrum was investigated (Figure 4).
First, the class of alcohols that can be used as substrate was investigated, testing ethanol (12), 2-propanol (1), and tert-butanol (11), which resulted in the conversion of only (1), indicating that neither primary nor tertiary alcohols are substrates for secADH.
Furthermore, we tested the reaction with diols (Figure S2). SecADH could convert diols when both alcohols were secondary ((7) and (8) in Figure 4), albeit at low conversion rates and only with an excess of each respective substrate. Diols, with one of the hydroxy groups being a primary alcohol, were not converted ((9) and (10) in Figure 4).
The enzyme preferred longer carbon chain substrates compared to short ones and can oxidize hydroxylated fatty acids besides alkanols, which we investigated by testing (2) and (3), and (5) and (6). The enzyme had a similar activity towards (5) and (6), and consequently, there was no discrimination between the two different alcohol positions within these alkyl chains. However, when comparing (2) and (3), there was a clear difference, with (3) being preferred. The used (2) was racemic, indicating that secADH prefers R-configurated alcohols—as 10- and 12-HSA are R-configurated as well—and/or alcohols that are not located at the rim of the alkane. There was no conversion observed for 1,1-dimethoxy-2-propanol (4).
A challenging aspect during the measurement was the general insolubility of longer alkanes and fatty acids in water. All substrate stocks were dissolved in ethanol; however, the enzyme reaction has to be performed in a buffer due to the enzyme’s stability. 10- and 12-HSA are highly insoluble in water. Only at a concentration of 500 µM in 5% (v/v) aqueous ethanol in the final reaction, the two long-chain fatty acids were stable in the buffer.
We could thus demonstrate that the newly identified secADH from Deinococcus radiodurans can be used in the desired cascade reaction.

2.5. ADH-Coupled Assay Reaction Parameter Optimization

One of the challenges in coupled assays is the diverging reaction parameters, which usually results from the different originating organisms of the involved catalysts or their varying place of action within the cell. In addition to OhyRe, other OHs were tested in combination with the secADH from Deinococcus radiodurans to investigate which one of the cascade members dominates the reaction parameters. The different tested OHs originated from Rhodococcus erythropolis, Pediococcus parvulus (OhyPp), and Elizabethkingia meningoseptica (OhyEm). Each of them has specific requirements for their optimal reaction activity.
First, we analyzed the optimal reaction parameters of secADH using 12-HSA as a substrate, and we could observe that it preferred basic environments over acidic ones, with an optimal pH of around 9 (Figure 5a). In addition to the overall optimum being in a basic environment, there was a second smaller optimum at a pH of around 7. Since fatty acids usually show unique behavior in basic environments as opposed to alkane-based alcohols, we performed a pH screening of secADH with 3-pentanol as the substrate (Figure S3). Surprisingly, the first pH optimum shifted towards a more basic environment compared to when 12-HSA was used as the substrate. Since oleic acid has a pka of 5.02 [18], we investigated the effect on the pH and whether this might be the reason for the activity shift. We indeed observed that the pH was shifted when 12-HSA was present in the reaction buffer, but it could not explain the change in the optimum (Table S1), thus indicating that the type of substrate for secADH can influence the reaction optima.
Regarding the temperature optimum of secADH, the highest activity was measured at 18 °C and the lowest at 22 °C, where residual activity of 60% still remained, reflecting a rather broad range (Figure 5b).
The broad-range temperature optimum is beneficial for a one-pot reaction; however, the high pH optimum of secADH is problematic since the optimum of most OHs is rather in the acidic range, presumably due to an evolutionary adaptation to the acidic nature of the substrates. OhyPp [18] and OhyEm [16], for instance, have an optimum of 6. OhyRe, however, is an exception since it prefers rather basic pH environments [17].
We analyzed the pH optima for the combination of secADH with two different OHs, OhyRe and OhyEm, since they hold varying pH optima (7.2 and 6, respectively). As shown in Figure 5c, the pH optimum was dependent on the type of OHs. In the cascade reaction with OhyRe and secADH, the optimum was measured to be around 6.5, which is close to the one from OhyRe alone [17]. On the other hand, in the cascade reaction with OhyEm, two optima were observed at around 7.5 and 9, which is more like the ones from secADH. Surprisingly, the combination of OhyRe and secADH showed a rather low conversion in basic environment despite OhyRe’s robustness at elevated pH values [17] and secADH’s optimum there.
The optimal temperature of the cascade reactions also depended on the type of OH used. Here, OhyPp was used, with its optimal temperature at 18 °C [18] and OhyRe at 28 °C [17]. For ADH and OhyRe, the optimum changed to 26 °C, and for ADH and OhyPp, to 20 °C (Figure 5d).
Next, the melting temperature of secADH was investigated at different pH values using protein thermal shift assays. SecADH is a stable enzyme, with its melting temperature measured to be 57.88 ± 0.22 °C at a pH of 6.5 (Table S2). Even though the highest activity was observed at pH 9, the general stability of secADH decreased with higher surrounding pH values (Figure 5f).
The stability was tested with protein thermal shift assays, where a bound ligand usually leads to a decrease in conformational protein flexibility, and thus, to an increase in its melting temperature [49]. For secADH, however, this was not the case since incubation with 3-pentanol and 2-pentanol did not lead to strong changes in the melting temperatures at pH values of 6.5, 7.5, or 9 (Table S2). On the contrary, when 12-HSA was used as the substrate, there was a significant decrease in the melting temperature at a pH of 9 from 53.43 ± 0.54 to 47.00 ± 0.87 °C (Figure 5e), which shows that the substrate strongly destabilized the protein at that pH. At the pH values of 6.5 and 7.5, secADH with 12-HSA showed only slight reductions in Tm, similar to the other substrates (Table S2).
In the coupled assay, secADH converted fatty acids, which are reported to behave differently depending on the environment, in particular, the pH value and the temperature [50]. Under basic conditions, they form mixtures of acid and soap, creating crystalline solids, which leads to non-accessible molecules and additionally might lead to the destabilization of proteins.
Most likely, the high pH and the soap attributes of the fatty acid, which became stronger at elevated pH values, had destabilizing effects on secADH. The OHs did not show these strong decreases in the melting temperatures during incubation with oleic acid, which suggests that they are evolutionarily better evolved to fatty acids as substrates (Table S3).
In summary, the temperature and pH optima cannot be concluded from the type of used OHs, but were different for every type of enzyme. The use of several enzymes within a cascade consequently led to higher complexity, and hence, the optimal parameters changed unpredictably. The results presented here show that either OH or secADH dominated the reaction conditions, which presumably depended on diverging kinetic behaviors, structural stability, or substrate access. The pH and temperature optima were closer to OhyRe’s than to secADH’s when they were combined in a cascade. In contrast, for OhyEm, the pH optimum followed the one for secADH more, which might have resulted from OhyEm being a faster enzyme compared to OhyRe, and hence, being the lesser rate-limiting factor within the cascade. To conclude, the conditions must be optimized separately for every type of OH.

2.6. ADH-Coupled Assay Reaction Parameter Optimization

Next, the effects of additives on the reaction cascade were tested. Oleic acid is insoluble in water, and OHs only have access to it when it is emulsified with water [15]. Additives like Tween 80 usually have positive effects on emulsification, and thus, on the activity [51,52]. Hence, the effect of the different additives (ethanol, Tween 80, dimethyl sulfoxide, and 1,5-pentanediol) in concentrations of 1, 2, 5, 10, and 20 % (v/v) on the cascade reaction was tested using OhyRe and secADH (Figure 6a). All additives showed positive effects only in low concentrations. At concentrations above 1% (v/v), the reactions started to be inhibited, except for ethanol, which led to the highest activity at 5% (v/v).
The effect was most profound for Tween 80, which, at 10% (v/v), led to a residual activity of only 30%. This is surprising since Tween 80 improved the activity of OhyRe alone strongly and at 10% (v/v), 96% residual activity, compared to the optimum at 5% (v/v), remained (Figure S4). The addition of Tween 80 to the reaction of secADH with 3-pentanol led to a decrease in activity as well, albeit not to such an extent as for the coupled assay. Tween 80 formed micellar structures with 10-HSA, potentially reducing the accessibility for secADH. The strongest increase, although still moderate compared to the control, could be observed for 1,5-pentanediol at 1% (v/v) (Figure 6b).
Since OhyRe’s activity was found to be enhanced by a broadband white light at an optimal range between 243 and 364 µmol/m2·s, the cascade reaction’s performance with varying light intensities was tested (Figure 6c). As expected, a 30-fold increase in the 10-HSA yield was observed for the cascade reaction when comparing dark conditions with a light intensity of 364 µmol/m2·s under broadband white light. The cascade reaction behaved differently on the light intensities compared to OhyRe alone since there was a sharp maximum at 364 µmol/m2·s, whereas for OhyRe alone, there was rather a broad optimal range. Notably, the 10-KSA concentration also increased by a 14-fold factor, proving that the cascade benefitted from the light activation as well.

2.7. Optimization of Enzyme Concentrations

The combination of the two enzymes required harmonized protein concentrations. For this, different enzyme concentrations were tested. Interestingly, the concentration dependencies of secADH and OhyRe behaved differently in the coupled assay. For OhyRe, a typical saturation curve can be observed, where the vmax, in this experiment, was reached at 0.057 ± 0.007 U/mL, and the half-maximal concentration at 0.221 ± 0.065 g/L (Figure 7). For secADH, however, an inhibition course can be observed. Here, vmax was predicted to be at 0.057 ± 0.011 U/mL, and the half-maximal concentration at 0.012 ± 0.011 g/L by non-linear regression, which is 18-fold lower compared to OhyRe and Ki 0.953 ± 0.800 g/L.
Furthermore, the concentration dependency curve of secADH shows the behavior of substrate inhibition. This is surprising for an enzyme saturation curve since it usually behaves like OhyRe (dark grey curve in Figure 7). It might be that the increase in secADH led to an accumulation of 10-KSA, which led to the stronger product inhibition of OhyRe compared to 10-HSA. Furthermore, it might be that product inhibition of 10-KSA on secADH occurred, leading to the observed inhibition curves. The inhibition curve might additionally have resulted from a back reaction, which led to a decrease in the NADH/H+ concentration.

2.8. Conversion of Cascade Reaction with OhyRe and secADH

With all of the optimized parameters, a long-time conversion was performed, investigating the conversion of substrates. In Figure 8a, the conversion over a period of 16 h is shown, which displays that the reaction was rather slow, as expected for OH despite the light-dependent activation. However, without that, the conversion might be even slower, as, in one study, a conversion was conducted over the course of 6 days [53]. The maximal conversion led to a residual substrate concentration of 20% of all measured fatty acids. 10-HSA was the predominant species among the products, and 10-KSA ranged only from 16 to 30% (Figure 8b and Table S4). This is surprising since OHs are often slow enzymes, as observable by their long turnover times [18,54], whereas the alcohol dehydrogenase was observed to act faster [55].
An explanation might be the still-not-optimal conditions for secADH due to the undertaken compromises. Since Wu et al. also noticed that reaction conditions diverted between OH and their secADH from Micrococcus luteus, they decided to go for a consecutive conversion, where first 10-HSA was produced and then secADH and NAD+-regenerating enzymes were added after pH adjustment [55]. Under these conditions, it was possible to reach a conversion of 95% of 10-HSA to 10-KSA. For the industrial production of 10-KSA on a large scale, the one-pot reaction is consequently rather unsuitable due to the low yield. 10-KSA, however, has less economic value compared to 10-HSA due to the unreactive 10-oxo group, and thus, this reaction is rather interesting for high-throughput screenings. However, for screening purposes, performing the reaction in a one-pot approach is strongly preferred since it saves time and consumables, and thus, the here-shown optimization of conditions can largely aid in its application. However, finding a 10-HSA converting secADH with a lower optimal pH might drastically increase the overall yield.
Finally, a combination of lipase, OhyRe, and secADH with oil from C. oleaginosus in an in vitro one-pot reaction was additionally conducted, and it was observed that around 80% of oleic acid was converted over 20 h, with around 60 % being 10-HSA, similar to the conversion shown in Figure 8. However, the complexity strongly increased with three different enzymes, and further optimization of this cascade is required to achieve the maximum output.

3. Materials and Methods

3.1. Chemicals

Chemicals were purchased from Thermo Fisher Scientific (Waltham, MA, USA), Merck (Darmstadt, Germany), and Carl Roth (Karlsruhe, Germany) at the highest available grade. Candida rugosa lipase was purchased from Merck. Commercially available olive, rapeseed, and linseed oils were purchased.

3.2. Cascade Reaction Using OhyRe and Lipase from C. rugosa

First, 1 µL of oil was added to 329 µL of buffer (20 mM Tris-Base, pH 7.2) and emulsified. Then, 50 µL of OhyRe (2 mg/mL with an equimolar concentration of FAD) and 20 µL of lipase (2 mg/mL) were added. Reactions were conducted for 20 h at 28 °C with 100 rpm of horizontal shaking. The samples were methylated and measured using GC-FID with a marine oil mix (Restek, Centre County, PA, USA) as an external standard.

3.3. Protein Expression

For protein expression, pet28a(+) containing secADH, OhyRe, OhyPp, and OhyEm was transformed into BL21DE3. Precultures were grown overnight in LB containing 50 µg/mL kanamycin. For the main culture, 500 mL of TB medium was inoculated to an OD600 of 0.05–0.1 and grown until they reached an OD600 of 0.6–0.8 in an Innova44R shaker (Eppendorf, Hamburg, Germany), after which the temperature was decreased to 16°C, and gene expression was induced with a final concentration of 0.1 mM IPTG. After 16 h, the cells were harvested and resuspended in resuspension buffer (20 mM Tris-Base pH 7.2, 20 mM imidazole, 500 mM NaCl,). The cells were disrupted using a high-pressure homogenizer (EmulsiFlex-B15, AVESTIN, Ottawa, ON, Canada), and debris was removed by centrifugation at 20,000× g for 40 min at 4 °C. Finally, the proteins were purified using Ni2+-NTA beads (Thermo Fisher Scientific), which were mixed and incubated overnight with the cell-free lysate. The beads were washed with resuspension buffer, proteins eluted using elution buffer (20 mM Tris-Base pH 7.2, 250 mM imidazole, 500 mM NaCl), and buffer exchanged with desalting columns (PD MidiTrap G-25; Cytiva, Marlborough, MA, USA). The final storage buffer was dependent on the type of enzyme and experiment and is described separately in the following paragraphs and underneath the figures. SecADH was stored in 20 mM Tris, pH 7.2. The protein concentrations were determined using ROTI®Quant with bovine serum albumin (both from Carl Roth).

3.4. Analytics

Assays were either analyzed using a multimode microplate reader (Enspire2; Perkin Elmer, Waltham, MA, USA) for 30 min at 340 nm or by GC-FID. The reactions were monitored for 30 min at 340 nm. For the calculation of U/mL, the initial linear slope of the absorbance curve of NADH, its extinction coefficient ε = 6.22 mM−1·cm−1, and a plate thickness of 0.59 cm were used. U is defined as (initial slope·total volume)/(plate thickness·enzyme volume· ε). For GC-FID, the reactions were stopped, and the fatty acids were extracted with 1 mL ethyl acetate, which was evaporated afterward. All fatty acids and oils measured on GC-FID were methylated and measured as previously described [18].

3.5. Light Intensity Test

For the light intensity tests, reactions were performed in an incubation shaker equipped with LEDs of adjustable light intensity and color (TB2000, FutureLED, Berlin, Germany). For the light color test, reactions were conducted in broadband white, blue (425 nm), and red (680 nm) light at photosynthetic active radiation of 191 µmol/m2·s for each condition, and in the dark. For the light intensity tests, broadband white light was used at 48, 243, 364, and 486 µmol/m2·s. Then, 1 µL oleic acid was emulsified with 149 µL buffer (20 mM Tris-Base pH 7.2, 200 mM NaCl), and 50 µL of OhyRe (30 µM with 30 µM FAD) was added. Reactions were conducted for 3 h at 28 °C and 100 rpm, followed by extraction with 1 mL of ethyl acetate with subsequent methylation. All following reactions were conducted in the LED shaker.

3.6. Optimal Conditions Test of secADH

For the pH optimum, 5 µL of 10 mM 12-HSA in ethanol was mixed with 10 µL of the secADH (0.2 g/L), 165 µL of each respective buffer, and 20 µL of NAD+ (10 mM). The reactions were measured with a multimode microplate reader.
For the temperature optimum, 25 µL of 12-HSA (10 mg/mL), 100 µL of the secADH (2 mg/mL), 375 µL of 100 mM Tris-Base pH 8, and 20 µL NAD+ (10 mM) were mixed and incubated for 10 min. The temperatures of 16, 18, 20, 22, 24, and 26 °C were tested. The samples were analyzed using GC-FID.

3.7. Optimal Conditions Test of Cascade Reaction with OH and secADH

For testing the pH optimum, oleic acid was emulsified with 20 mM Tris-Base buffer pH 7.2 to a concentration of 10 mM, and 20 µL of it was added to 20 µL of OhyRe or OhyEm (30 µM in 50 mM MES buffer pH 6.5, with an equimolar concentration of FAD for OhyRe), 20 µL NAD+ (10 mM), 3 µL ADH (2 mg/mL), and the respective volume of each respective buffer to a total volume of 200 µL. The reactions were measured with a multimode microplate reader.
For testing the temperature optimum, 1 µL of oleic acid was added to 126 µL of 50 mM MES pH 6.5 and emulsified. Then, 50 µL OhyRe or OhyPp (30 µM in 50 mM MES buffer pH 6.5 for OhyRe and in 20 mM Tris-Base pH 7.2 for OhyPp, with an equimolar concentration of FAD), 20 µL NAD+ (10 mM) and 3 µL of the secADH (2 mg/mL) were added. The experiments were conducted in triplicate for 15 min, and the samples were analyzed using GC-FID.

3.8. Buffer for pH-Tests

For the analysis of the pH optima, the following buffers were used: pH 5: citrate; pH 5.5-6.5: MES; pH 7-8.5: Tris-Base; pH 9-9.5: TAPS; pH 10: CAPS, all with a concentration of 100 mM.

3.9. Substrate Specificity Test for secADH

To initially investigate the substrate specificity, 10 µL of ethanol, tert.-butanol, and 2-propanol were mixed with 10 µL of ADH (2 mg/mL), 20 µL of NAD+ (10 mM), and 160 µL of 20 mM Tris-Base, pH 7.2. For the conversion of diols, 1 M stocks, and for the secondary alcohols, 10 mM stocks, were prepared in ethanol. The reactions were conducted as described above and analyzed in a multimode microplate reader at 340 nm.

3.10. Determination of Melting Temperatures

First, 5 µL of ADH (2 mg/mL) was mixed with 2.5 µL SYPRO™ Orange (Thermo Fisher Scientific) (1:200), 1.3 µL of ethanol or 1.3 µL of 10 mM of each of the secondary alcohols dissolved in ethanol and 16.2 µL buffer in qPCR tubes with optically clear lids. For pH 6.5 MES, for pH 7.5 Tris-Base, and for pH 9, TAPS buffer was used (all with a concentration of 100 mM). After putting the samples on ice, they were transferred to an RT-PCR cycler (CFX Opus 96, Bio-Rad, Feldkirchen, Germany). The ramp went from 10 to 95 °C in 0.5 °C steps, holding for 10 s per cycle. The experiments were conducted in quadruplicate.

3.11. Testing of Different Additives

First, 10 mM oleic acid was mixed with 50 mM MES buffer pH 6.5 to an emulsion, and 20 µL of it was added to 20 µL OhyRe (30 µM with an equimolar concentration of FAD), 3 µL of the secADH (2 mg/mL), 20 µL NAD+ (10 mM) of the respective volume of the additive, and 50 mM MES buffer pH 6.5 to a total volume of 200 µL. The reactions were monitored at 340 nm for 30 min.

3.12. Optimization of Enzyme Concentrations

For the fixed secADH concentrations (dark grey curve in Figure 7), the following conditions were used: oleic acid was mixed with 50 mM MES buffer pH 6.5 to an emulsion of 10 mM, and 20 µL of it was added to different final concentrations of OhyRe (stored with an equimolar concentration of FAD) and 0.01 g/L ADH, 20 µL NAD+ (10 mM), and 50 mM MES buffer pH 6.5 to a total volume of 200 µL. For the fixed OhyRe concentrations (red curve in Figure 7), a final concentration of OhyRe of 1 g/L was used. The reactions were monitored at 340 nm for 30 min.

4. Conclusions

In summary, rapid light-driven activation of OhyRe was presented with broadband white light at an intensity of 364 µmol/m2·s, leading to the highest increase in activity. This most likely can be attributed to a partial reduction of FAD to FADH2 without the utilization of anaerobic conditions.
Furthermore, two cascade reactions were presented, with one being upstream and one being downstream of OhyRe. For the upstream one, a lipase from Candida rugosa and OhyRe was presented, leading to the hydrolysis of triglyceride oils into free fatty acids and their subsequent hydroxylation. The yield was decreased with increasing amounts of oleic acid; however, other factors, such as remaining impurities, may also play a role.
The downstream cascade reaction comprised a novel alcohol dehydrogenase from Deinococcus radiodurans, which was functionally expressed, and its substrate specificity was investigated, proving a preference for long-carbon-chain secondary alcohols. It also converted hydroxylated fatty acids, which makes it an ideal candidate for high-throughput screenings of optimized OHs and large-scale conversions of 10-HSA. To be suitable for screenings, the optimal reaction conditions were investigated. The results show that the pH and temperature optima depended on the type of OHs and might either follow OH’s or secADH’s preferences. Furthermore, hydroxylated fatty acids destabilized secADH at elevated pH values and became more unstable with increasing pH despite showing the highest activity in basic environments. Since OHs prefer oleic acid as substrate in an emulsion, they had increased activity with emulsifiers such as ethanol or Tween 80. However, when using them together with secADH, the activity improvement was only marginal at low concentrations of these additives. At higher concentrations, a drastic decrease in activity occurred. Furthermore, it could be shown that a coupled assay can benefit from improvements in the OH reaction. The cascade reaction showed an increase in activity by a factor of 30 upon incubation under light. Still, the concentration of 10-KSA was rather low compared to 10-HSA. Additional optimizations can facilitate the achievement of further enhanced product concentrations, including the engineering of a secADH to enhance resistance to pH variations and additives.
We also showcased a proof-of-concept, indicating that a coupled assay using lipase, OhyRe, and a secADH to generate 10-KSA out of oil from C. oleaginosus is feasible.
In general, the utilization of cascade reactions in conjunction with OHs could potentially significantly impact catalyst optimization screenings and industrial applications in the future. This is due to their ability to reduce the number of processing steps, leading to savings in both time and costs. The data presented here represent initial strides toward the implementation of these cascade reactions.
In future activities, our group will validate and optimize the herein-described enzyme cascades at technical scales to generate data for a techno-economic analysis that will guide potential subsequent commercialization efforts.
In this context, fermentative enzyme production expenses are a main cost driver that may be an economic barrier in the industrialization process. The alleviation of these economic constraints requires the use of cost-efficient, complex feedstocks in enzyme production [56]. Significant progress has been made in achieving cost-efficient production of lipase, an industrially advanced enzyme system [57]. In follow-up experiments, lipase and OH can be produced from biomass residue streams, such as enzymatically hydrolyzed sunflower husks or wheat spelts, which are current milling by-products with little valorization. Our group has significant experience with these feedstocks and is in the process of developing economically viable processes in this respect.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/catal13091236/s1. Figure S1: SDS-page gel of the novel secADH from Deinococcus radiodurans. Predicted molecular weight is 32.4 kDa. Figure S1: Comparison of the conversion in U/ml of the diols 2,4-pentanediol, 2,5-hexanediol, 1,3-butanediol with the secondary alcohol 2-propanol. 1,2-butanediol didn’t show any conversion. Figure S2: pH-optimum of secADH using 12-HSA compared with 3-pentanol as substrates. 5 µL of 10 mM 12-HSA in ethanol were mixed with 10 µL of secADH (0.2 mg/mL), 20 µL of NAD+ (10 mM) and 165 µL of each respective buffer (100 mM concentrations). Figure S3: Effect of Tween 80 on the conversion of oleic acid using OhyRe. Oleic acid was mixed with 20 mM Tris-Base buffer pH 7.2 to an emulsion of 10 mM and 20 µL of it were added to 20 µL OhyRe (30 µM with an equimolar concentration of FAD), 20 µL NAD+ (10 mM) the respective volume of the additive and 20 mM Tris-Base buffer pH 7.2 to a total volume of 200 µL. Table S1: Anticipated pH and actual (measured) pH using 5 µL of 10 mM 12-HSA in ethanol mixed with 10 µL of secADH (0.2 mg/mL in 20 mM Tris-Base pH 7.2), 20 µL of NAD+ (10 mM in 20 mM Tris-Base pH 7.2) and 165 µL of each respective buffer (100 mM concentrations). Table S2: Melting temperatures (in °C) of secADH incubated with different secondary alcohols. Table S3: Melting temperatures (in °C) of OHs incubated with or without oleic acid in 200-molar excess. Oleic acid was dissolved in ethanol, which was evaporated prior the addition of the other components to assure an appropriate final concentration. The protein thermal shift assays were conducted at a pH of 7.2 for OhyRe and at a pH of 6 for OhyPp and OhyEm. Table S1: Conversion ratio of 10-KSA to total product concentration after 1, 2, 4, 6 and 16 h.

Author Contributions

Conceptualization, S.A.P., M.H. and F.M.; methodology, S.A.P., M.H. and M.R.; validation, S.A.P. and M.H.; formal analysis, S.A.P.; investigation, S.A.P. and M.R.; resources, S.A.P.; data curation, S.A.P.; writing—original draft preparation, S.A.P. and F.M.; writing—review and editing, S.A.P., M.H., M.R., F.M., D.G. and T.B.; supervision, D.G. and T.B.; project administration, D.G. and T.B.; funding acquisition, T.B. All authors have read and agreed to the published version of the manuscript.

Funding

We greatly acknowledge the German Federal Ministry of Education and Research (Bundesministerium für Bildung und Forschung), which funded this research project (grant number 03SF0577A).

Data Availability Statement

The data presented in this study are available upon request from the corresponding author.

Acknowledgments

We want to thank Maria Bandookwala for her support in the data acquisition. Furthermore, we appreciate Mahmoud Masri and Global Sustainable Transformation GmbH for providing microbial oil from C. oleaginosus. We would like to extend our sincere gratitude to FUCHS LUBRICANTS GERMANY GmbH for their generous provision of 12-HSA and HOSO for use in our research.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Claassens, N.J.; Burgener, S.; Vögeli, B.; Erb, T.J.; Bar-Even, A. A Critical Comparison of Cellular and Cell-Free Bioproduction Systems. Curr. Opin. Biotechnol. 2019, 60, 221–229. [Google Scholar] [CrossRef]
  2. Siedentop, R.; Claaßen, C.; Rother, D.; Lütz, S.; Rosenthal, K. Getting the Most out of Enzyme Cascades: Strategies to Optimize in Vitro Multi-enzymatic Reactions. Catalysts 2021, 11, 1183. [Google Scholar] [CrossRef]
  3. Hwang, E.T.; Lee, S. Multienzymatic Cascade Reactions via Enzyme Complex by Immobilization. ACS Catal. 2019, 9, 4402–4425. [Google Scholar] [CrossRef]
  4. Lin, B.; Tao, Y. Whole-Cell Biocatalysts by Design. Microb. Cell Fact. 2017, 16, 1–12. [Google Scholar] [CrossRef] [PubMed]
  5. Guterl, J.K.; Garbe, D.; Carsten, J.; Steffler, F.; Sommer, B.; Reiße, S.; Philipp, A.; Haack, M.; Rühmann, B.; Koltermann, A.; et al. Cell-Free Metabolic Engineering: Production of Chemicals by Minimized Reaction Cascades. ChemSusChem 2012, 5, 2165–2172. [Google Scholar] [CrossRef]
  6. Voss, M.; Küng, R.; Hayashi, T.; Jonczyk, M.; Niklaus, M.; Iding, H.; Wetzl, D.; Buller, R. Multi-Faceted Set-up of a Diverse Ketoreductase Library Enables the Synthesis of Pharmaceutically-Relevant Secondary Alcohols. ChemCatChem 2020, 13, 1538–1545. [Google Scholar] [CrossRef]
  7. Siedentop, R.; Rosenthal, K. Industrially Relevant Enzyme Cascades for Drug Synthesis and Their Ecological Assessment. Int. J. Mol. Sci. 2022, 23, 3605. [Google Scholar] [CrossRef]
  8. Begander, B.; Huber, A.; Döring, M.; Sperl, J.; Sieber, V. Development of an Improved Peroxidase-Based High-Throughput Screening for the Optimization of d-Glycerate Dehydratase Activity. Int. J. Mol. Sci. 2020, 21, 335. [Google Scholar] [CrossRef]
  9. Bub, O.; Jager, S.; Dold, S.M.; Zimmermann, S.; Hamacher, K.; Schmitz, K.; Rudat, J. Statistical Evaluation of Hts Assays for Enzymatic Hydrolysis of β-Keto Esters. PLoS ONE 2016, 11, e0146104. [Google Scholar] [CrossRef]
  10. Kiianitsa, K.; Solinger, J.A.; Heyer, W.D. NADH-Coupled Microplate Photometric Assay for Kinetic Studies of ATP-Hydrolyzing Enzymes with Low and High Specific Activities. Anal. Biochem. 2003, 321, 266–271. [Google Scholar] [CrossRef]
  11. Fang, X.; Zheng, Y.; Duan, Y.; Liu, Y.; Zhong, W. Recent Advances in Design of Fluorescence-Based Assays for High-Throughput Screening. Anal. Chem. 2019, 91, 482–504. [Google Scholar] [CrossRef] [PubMed]
  12. Kozaeva, E.; Mol, V.; Nikel, P.I.; Nielsen, A.T. High-Throughput Colorimetric Assays Optimized for Detection of Ketones and Aldehydes Produced by Microbial Cell Factories. Microb. Biotechnol. 2022, 15, 2426–2438. [Google Scholar] [CrossRef] [PubMed]
  13. Leippe, D.M.; Nguyen, D.; Zhou, M.; Good, T.; Kirkland, T.A.; Scurria, M.; Bernad, L.; Ugo, T.; Vidugiriene, J.; Cali, J.J.; et al. A Bioluminescent Assay for the Sensitive Detection of Proteases. Biotechniques 2011, 51, 105–110. [Google Scholar] [CrossRef] [PubMed]
  14. Zegzouti, H.; Zdanovskaia, M.; Hsiao, K.; Goueli, S.A. ADP-Glo: A Bioluminescent and Homogeneous Adp Monitoring Assay for Kinases. Assay Drug Dev. Technol. 2009, 7, 560–572. [Google Scholar] [CrossRef]
  15. Prem, S.; Helmer, C.P.O.; Dimos, N.; Himpich, S.; Brück, T.; Garbe, D.; Loll, B. Towards an Understanding of Oleate Hydratases and Their Application in Industrial Processes. Microb. Cell Fact. 2022, 21, 1–15. [Google Scholar] [CrossRef]
  16. Engleder, M.; Pavkov-Keller, T.; Emmerstorfer, A.; Hromic, A.; Schrempf, S.; Steinkellner, G.; Wriessnegger, T.; Leitner, E.; Strohmeier, G.A.; Kaluzna, I.; et al. Structure-Based Mechanism of Oleate Hydratase from Elizabethkingia Meningoseptica. ChemBioChem 2015, 16, 1730–1734. [Google Scholar] [CrossRef]
  17. Lorenzen, J.; Driller, R.; Waldow, A.; Qoura, F.; Loll, B.; Brück, T. Rhodococcus Erythropolis Oleate Hydratase: A New Member in the Oleate Hydratase Family Tree—Biochemical and Structural Studies. ChemCatChem 2018, 10, 407–414. [Google Scholar] [CrossRef]
  18. Prem, S.A.; Helmer, C.P.O.; Loll, B.; Garbe, D.; Brück, T. Expanding the Portfolio by a Novel Monomeric Oleate Hydratase from Pediococcus Parvulus. ChemCatChem 2023, 15. [Google Scholar] [CrossRef]
  19. Volkov, A.; Khoshnevis, S.; Neumann, P.; Herrfurth, C.; Wohlwend, D.; Ficner, R.; Feussner, I. Crystal Structure Analysis of a Fatty Acid Double-Bond Hydratase from Lactobacillus Acidophilus. Acta Crystallogr. Sect. D Biol. Crystallogr. 2013, 69, 648–657. [Google Scholar] [CrossRef]
  20. Masri, M.A.; Garbe, D.; Mehlmer, N.; Brück, T.B. A Sustainable, High-Performance Process for the Economic Production of Waste-Free Microbial Oils That Can Replace Plant-Based Equivalents. Energy Environ. Sci. 2019, 12, 2717–2732. [Google Scholar] [CrossRef]
  21. Ochsenreither, K.; Glück, C.; Stressler, T.; Fischer, L.; Syldatk, C. Production Strategies and Applications of Microbial Single Cell Oils. Front. Microbiol. 2016, 7. [Google Scholar] [CrossRef] [PubMed]
  22. Fuchs, T.; Melcher, F.; Rerop, Z.S.; Lorenzen, J.; Shaigani, P.; Awad, D.; Haack, M.; Prem, S.A.; Masri, M.; Mehlmer, N.; et al. Identifying Carbohydrate-Active Enzymes of Cutaneotrichosporon Oleaginosus Using Systems Biology. Microb. Cell Fact. 2021, 20, 1–18. [Google Scholar] [CrossRef]
  23. Shaigani, P.; Fuchs, T.; Graban, P.; Prem, S.; Haack, M.; Masri, M.; Mehlmer, N.; Brueck, T. Mastering Targeted Genome Engineering of GC-Rich Oleaginous Yeast for Tailored Plant Oil Alternatives for the Food and Chemical Sector. Microb. Cell Fact. 2023, 22, 1–14. [Google Scholar] [CrossRef] [PubMed]
  24. Zhang, Y.; Eser, B.E.; Kristensen, P.; Guo, Z. Fatty Acid Hydratase for Value-Added Biotransformation: A Review. Chin. J. Chem. Eng. 2020, 28, 2051–2063. [Google Scholar] [CrossRef]
  25. Schmid, J.; Steiner, L.; Fademrecht, S.; Pleiss, J.; Otte, K.B.; Hauer, B. Biocatalytic Study of Novel Oleate Hydratases. J. Mol. Catal. B Enzym. 2016, 133, S243–S249. [Google Scholar] [CrossRef]
  26. Betancor, L.; Hidalgo, A.; Fernández-Lorente, G.; Mateo, C.; Rodríguez, V.; Fuentes, M.; López-Gallego, F.; Fernández-Lafuente, R.; Guisan, J.M. Use of Physicochemical Tools to Determine the Choice of Optimal Enzyme: Stabilization of D-Amino Acid Oxidase. Biotechnol. Prog. 2003, 19, 784–788. [Google Scholar] [CrossRef]
  27. Hagedoorn, P.L.; Hoolmann, F.; Hanefeld, U. Novel Oleate Hydratases and Potential Biotechnological Applications. Appl. Microbiol. Biotechnol. 2021, 105, 6159–6172. [Google Scholar] [CrossRef]
  28. Boratynski, F.; Szczepanska, E.; De Simeis, D.; Serra, S.; Brenna, E. Bacterial Biotransformation of Oleic Acid: New Findings on the Formation of γ-Dodecalactone and 10-Ketostearic Acid in the Culture of Micrococcus Luteus. Molecules 2020, 25, 3024. [Google Scholar] [CrossRef]
  29. Huang, J.K.; Samassekou, K.; Alhmadi, H.B.; VanDerway, D.R.; Diaz, J.D.; Seiver, J.A.; McClenahan, S.W.; Holt, S.M.; Wen, L. Knockout of Secondary Alcohol Dehydrogenase in Nocardia Cholesterolicum NRRL 5767 by CRISPR/Cas9 Genome Editing Technology. PLoS ONE 2020, 15, e0230915. [Google Scholar] [CrossRef]
  30. Sun, Q.F.; Zheng, Y.C.; Chen, Q.; Xu, J.H.; Pan, J. Engineering of an Oleate Hydratase for Efficient C10-Functionalization of Oleic Acid. Biochem. Biophys. Res. Commun. 2021, 537, 64–70. [Google Scholar] [CrossRef]
  31. Chandra, P.; Enespa; Singh, R.; Arora, P.K. Microbial Lipases and Their Industrial Applications: A Comprehensive Review. Microb. Cell Fact. 2020, 19. [Google Scholar] [CrossRef] [PubMed]
  32. Ranganathan, S.; Gärtner, T.; Wiemann, L.O.; Sieber, V. A One Pot Reaction Cascade of in Situ Hydrogen Peroxide Production and Lipase Mediated in Situ Production of Peracids for the Epoxidation of Monoterpenes. J. Mol. Catal. B Enzym. 2015, 114, 72–76. [Google Scholar] [CrossRef]
  33. Van Dongen, S.F.M.; Nallani, M.; Cornelissen, J.J.L.M.; Nolte, R.J.M.; Hest, J.C.M. Van A Three-Enzyme Cascade Reaction through Positional Assembly of Enzymes in a Polymersome Nanoreactor. Chemistry (Easton) 2009, 15, 1107–1114. [Google Scholar] [CrossRef]
  34. Srinivasamurthy, V.S.T.; Böttcher, D.; Bornscheuer, U.T. A Multi-Enzyme Cascade Reaction for the Production of 6-Hydroxyhexanoic Acid. Zeitschrift fur Naturforsch.-Sect. C J. Biosci. 2019, 74, 71–76. [Google Scholar] [CrossRef]
  35. Serra, S.; De Simeis, D.; Marzorati, S.; Valentino, M. Oleate Hydratase from Lactobacillus Rhamnosus Atcc 53103: A Fadh2-Dependent Enzyme with Remarkable Industrial Potential. Catalysts 2021, 11, 1051. [Google Scholar] [CrossRef]
  36. Lukacs, A.; Haigney, A.; Brust, R.; Zhao, R.K.; Stelling, A.L.; Clark, I.P.; Towrie, M.; Greetham, G.M.; Meech, S.R.; Tonge, P.J. Photoexcitation of the Blue Light Using FAD Photoreceptor AppA Results in Ultrafast Changes to the Protein Matrix. J. Am. Chem. Soc. 2011, 133, 16893–16900. [Google Scholar] [CrossRef] [PubMed]
  37. Masuda, S.; Hasegawa, K.; Ishii, A.; Ono, T.A. Light-Induced Structural Changes in a Putative Blue-Light Receptor with a Novel FAD Binding Fold Sensor of Blue-Light Using FAD (BLUF); Slr1694 of Synechocystis Sp. PCC6803. Biochemistry 2004, 43, 5304–5313. [Google Scholar] [CrossRef]
  38. Benjamin, S.; Pandey, A. Candida Rugosa Lipases: Molecular Biology and Versatility in Biotechnology. Yeast 1998, 14, 1069–1087. [Google Scholar] [CrossRef]
  39. Macrae, A.R.; Hammond, R.C. Present and Future Applications of Lipases. Biotechnol. Genet. Eng. Rev. 1985, 3, 193–217. [Google Scholar] [CrossRef]
  40. Melcher, F.; Ringel, M.; Vogelgsang, F.; Haack, M.; Masri, M.; Brück, T.; Garbe, D.; Roth, A. Lipase-Mediated Plant Oil Hydrolysis—Toward a Quantitative Glycerol Recovery for the Synthesis of Pure Allyl Alcohol and Acrylonitrile. Eur. J. Lipid Sci. Technol. 2023, 2200196, 1–12. [Google Scholar] [CrossRef]
  41. Di Pietro, M.E.; Mannu, A.; Mele, A. NMR Determination of Free Fatty Acids in Vegetable Oils. Processes 2020, 8, 410. [Google Scholar] [CrossRef]
  42. Ostojčić, M.; Budžaki, S.; Flanjak, I.; Bilić Rajs, B.; Barišić, I.; Tran, N.N.; Hessel, V.; Strelec, I. Production of Biodiesel by Burkholderia Cepacia Lipase as a Function of Process Parameters. Biotechnol. Prog. 2021, 37, e3109. [Google Scholar] [CrossRef]
  43. Yiitolu, M.; Temoçin, Z. Immobilization of Candida Rugosa Lipase on Glutaraldehyde-Activated Polyester Fiber and Its Application for Hydrolysis of Some Vegetable Oils. J. Mol. Catal. B Enzym. 2010, 66, 130–135. [Google Scholar] [CrossRef]
  44. Reis, P.; Holmberg, K.; Watzke, H.; Leser, M.E.; Miller, R. Lipases at Interfaces: A Review. Adv. Colloid Interface Sci. 2009, 147–148, 237–250. [Google Scholar] [CrossRef]
  45. Huang, J.-K.; Park, J.K.; Dhungana, B.R.; Youngblut, N.D.; Lin, C.-T.; Wen, L. A Novel Secondary Alcohol Dehydrogenase from Micrococcus Luteus WIUJH20: Purification, Cloning, and Properties. FASEB J. 2010, 24, 835.5. [Google Scholar] [CrossRef]
  46. Li, J.; Webster, T.J.; Tian, B. Functionalized Nanomaterial Assembling and Biosynthesis Using the Extremophile Deinococcus Radiodurans for Multifunctional Applications. Small 2019, 15, 1–15. [Google Scholar] [CrossRef]
  47. Waterhouse, A.; Bertoni, M.; Bienert, S.; Studer, G.; Tauriello, G.; Gumienny, R.; Heer, F.T.; De Beer, T.A.P.; Rempfer, C.; Bordoli, L.; et al. SWISS-MODEL: Homology Modelling of Protein Structures and Complexes. Nucleic Acids Res. 2018, 46, W296–W303. [Google Scholar] [CrossRef] [PubMed]
  48. Gasteiger, E.; Hoogland, C.; Gattiker, A.; Duvaud, S.; Wilkins, M.R.; Appel, R.D.; Bairoch, A. Protein Identification and Analysis Tools on the Expasy Server. In The Proteomics Protocols Handbook; Humana: Totowa, NJ, Canada, 2005; pp. 571–607. [Google Scholar]
  49. Celej, M.S.; Montich, G.G.; Fidelio, G.D. Protein Stability Induced by Ligand Binding Correlates with Changes in Protein Flexibility. Protein Sci. 2003, 12, 1496–1506. [Google Scholar] [CrossRef]
  50. Small, D.M. Physical Properties of Fatty Acids and Their Extracellular and Intracellular Distribution. Polyunsaturated Fat. Acids Hum. Nutr. Nestlé Nutr. Work. Ser. 1992, 28, 25–39. [Google Scholar]
  51. Park, J.Y.; Lee, S.H.; Kim, K.R.; Park, J.B.; Oh, D.K. Production of 13S-Hydroxy-9(Z)-Octadecenoic Acid from Linoleic Acid by Whole Recombinant Cells Expressing Linoleate 13-Hydratase from Lactobacillus Acidophilus. J. Biotechnol. 2015, 208, 1–10. [Google Scholar] [CrossRef]
  52. Kim, B.N.; Yeom, S.J.; Oh, D.K. Conversion of Oleic Acid to 10-Hydroxystearic Acid by Whole Cells of Stenotrophomonas Nitritireducens. Biotechnol. Lett. 2011, 33, 993–997. [Google Scholar] [CrossRef] [PubMed]
  53. Busch, H.; Tonin, F.; Alvarenga, N.; van den Broek, M.; Lu, S.; Daran, J.M.; Hanefeld, U.; Hagedoorn, P.L. Exploring the Abundance of Oleate Hydratases in the Genus Rhodococcus—Discovery of Novel Enzymes with Complementary Substrate Scope. Appl. Microbiol. Biotechnol. 2020, 104, 5801–5812. [Google Scholar] [CrossRef]
  54. Zhang, J.; Bilal, M.; Liu, S.; Zhang, J.; Lu, H.; Luo, H.; Luo, C.; Shi, H.; Iqbal, H.M.N.; Zhao, Y. Sustainable Biotransformation of Oleic Acid to 10-Hydroxystearic Acid by a Recombinant Oleate Hydratase from Lactococcus Garvieae. Processes 2019, 7, 326. [Google Scholar] [CrossRef]
  55. Wu, Y.X.; Pan, J.; Yu, H.L.; Xu, J.H. Enzymatic Synthesis of 10-Oxostearic Acid in High Space-Time Yield via Cascade Reaction of a New Oleate Hydratase and an Alcohol Dehydrogenase. J. Biotechnol. X 2019, 2, 3–7. [Google Scholar] [CrossRef] [PubMed]
  56. Sakhuja, D.; Ghai, H.; Rathour, R.K.; Kumar, P.; Bhatt, A.K.; Bhatia, R.K. Cost-Effective Production of Biocatalysts Using Inexpensive Plant Biomass: A Review. 3 Biotech 2021, 11, 280. [Google Scholar] [CrossRef]
  57. Abdelmoez, W.; Mostafa, N.A.; Mustafa, A. Utilization of Oleochemical Industry Residues as Substrates for Lipase Production for Enzymatic Sunflower Oil Hydrolysis. J. Clean. Prod. 2013, 59, 290–297. [Google Scholar] [CrossRef]
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Figure 1. (a) “Upstream” cascade reaction, with the combination of a lipase and an OH to produce 10-HSA without the need for prior hydrolysis of oils into free fatty acids. (b) “Downstream” cascade reaction, with the combination of an OH and a secondary alcohol dehydrogenase to produce 10-KSA.
Figure 1. (a) “Upstream” cascade reaction, with the combination of a lipase and an OH to produce 10-HSA without the need for prior hydrolysis of oils into free fatty acids. (b) “Downstream” cascade reaction, with the combination of an OH and a secondary alcohol dehydrogenase to produce 10-KSA.
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Figure 2. (a) Effects of different light colors (broadband white, blue (425 nm), red (680 nm), and no light (dark)) on the conversion of oleic acid into 10-HSA by OhyRe. Conversions are presented relative to the highest yield, which was observed under broadband white light. The photosynthetic active radiation was 191 µmol/m2·s or each condition. (b) Conversion rates of the biosynthesis of 10-HSA by applying varying light intensity in µmol/m2·s using broadband white light. Reactions were conducted in triplicates.
Figure 2. (a) Effects of different light colors (broadband white, blue (425 nm), red (680 nm), and no light (dark)) on the conversion of oleic acid into 10-HSA by OhyRe. Conversions are presented relative to the highest yield, which was observed under broadband white light. The photosynthetic active radiation was 191 µmol/m2·s or each condition. (b) Conversion rates of the biosynthesis of 10-HSA by applying varying light intensity in µmol/m2·s using broadband white light. Reactions were conducted in triplicates.
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Catalysts 13 01236 g003
Figure 3. Conversion of different types of oils to free fatty acids and 10-HSA using a lipase from C. rugosa and OhyRe. Experiments were conducted in triplicates. (a) Conversion of oleic acid to 10-HSA in (%) for high-oleic sunflower oil, olive oil, rapeseed oil, microbial oil from C.o., and linseed oil. (b) Conversion of oleic acid to 10-HSA in (%) per oleic acid content of each oil type and the oleic acid content in (%).
Figure 3. Conversion of different types of oils to free fatty acids and 10-HSA using a lipase from C. rugosa and OhyRe. Experiments were conducted in triplicates. (a) Conversion of oleic acid to 10-HSA in (%) for high-oleic sunflower oil, olive oil, rapeseed oil, microbial oil from C.o., and linseed oil. (b) Conversion of oleic acid to 10-HSA in (%) per oleic acid content of each oil type and the oleic acid content in (%).
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Figure 4. Structures of different substrates tested for conversion by secADH. (a) Secondary alcohols, (b) diols, (c) primary/tertiary alcohols. The substrates were 2-propanol (1), 2-pentanol (2), 3-pentanol (3), 1,1-dimethoxy-2-propanol (4), 10-HSA (5), 12-HSA (6), 2,4-pentanediol (7), 2,5-hexanediol (8), 1,3-butanediol (9), 1,2-butanediol (10), tert-butanol (11), ethanol (12). (d) Conversion of the main substrates 12-HSA, 10-HSA, 3-pentanol, 2-pentanol, and 2-propanol in U/mL.
Figure 4. Structures of different substrates tested for conversion by secADH. (a) Secondary alcohols, (b) diols, (c) primary/tertiary alcohols. The substrates were 2-propanol (1), 2-pentanol (2), 3-pentanol (3), 1,1-dimethoxy-2-propanol (4), 10-HSA (5), 12-HSA (6), 2,4-pentanediol (7), 2,5-hexanediol (8), 1,3-butanediol (9), 1,2-butanediol (10), tert-butanol (11), ethanol (12). (d) Conversion of the main substrates 12-HSA, 10-HSA, 3-pentanol, 2-pentanol, and 2-propanol in U/mL.
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Figure 5. pH optima were analyzed using a multimode microplate reader, and temperature optima using GC-FID. Relative activities were given to the maximum of each respective curve. Experiments were conducted in triplicates; melting temperature analyses were conducted in quadruplicates. (a) pH optimum and (b) temperature optimum of secADH using 12-HSA as substrate. (c) Comparison of the pH optima of two cascade reactions with secADH and either OhyRe or OhyEm. (d) Comparison of the temperature optima of two cascade reactions with secADH and either OhyRe or OhyPp. (e) Analysis of the melting temperatures of secADH with and without secondary alcohols at pH 9 using 100 mM TAPS buffer. There was no statistically relevant difference between without sec-alcohol and 2- and 3-pentanol as calculated using a two-sided t-test. (f) Comparison of melting temperatures of secADH with and without 12-HSA at different pH values. p < 0.01. All variants were compared with each other using a t-test and show a p-value < 0.001, except for the comparison of pH 6.5 to 7.5, with a p-value < 0.01, as shown in the figure.
Figure 5. pH optima were analyzed using a multimode microplate reader, and temperature optima using GC-FID. Relative activities were given to the maximum of each respective curve. Experiments were conducted in triplicates; melting temperature analyses were conducted in quadruplicates. (a) pH optimum and (b) temperature optimum of secADH using 12-HSA as substrate. (c) Comparison of the pH optima of two cascade reactions with secADH and either OhyRe or OhyEm. (d) Comparison of the temperature optima of two cascade reactions with secADH and either OhyRe or OhyPp. (e) Analysis of the melting temperatures of secADH with and without secondary alcohols at pH 9 using 100 mM TAPS buffer. There was no statistically relevant difference between without sec-alcohol and 2- and 3-pentanol as calculated using a two-sided t-test. (f) Comparison of melting temperatures of secADH with and without 12-HSA at different pH values. p < 0.01. All variants were compared with each other using a t-test and show a p-value < 0.001, except for the comparison of pH 6.5 to 7.5, with a p-value < 0.01, as shown in the figure.
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Figure 6. (a) Evaluating the effects of the four additives of ethanol, Tween 80, dimethyl sulfoxide, and 1,5-pentanediol at concentrations of 1, 2, 5, 10, and 20% (v/v) on a cascade reaction with OhyRe and secADH using oleic acid as substrate. (b) Fold improvement from reactions with no additives to those with additives for each of the optimal concentrations was used. (c) Evaluating different light intensities for the reduction of FAD. The same concentrations and volumes of reaction components were used as in (a) except for the additives, where the buffer volume was adjusted accordingly. Reactions were conducted for 3 h.
Figure 6. (a) Evaluating the effects of the four additives of ethanol, Tween 80, dimethyl sulfoxide, and 1,5-pentanediol at concentrations of 1, 2, 5, 10, and 20% (v/v) on a cascade reaction with OhyRe and secADH using oleic acid as substrate. (b) Fold improvement from reactions with no additives to those with additives for each of the optimal concentrations was used. (c) Evaluating different light intensities for the reduction of FAD. The same concentrations and volumes of reaction components were used as in (a) except for the additives, where the buffer volume was adjusted accordingly. Reactions were conducted for 3 h.
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Figure 7. Evaluating different concentrations for secADH and OhyRe in a coupled assay. Dark grey curve: fixed secADH concentration (0.01 g/L). Red curve: fixed OhyRe concentration (1 g/L). The response for OhyRe was fitted using y = Vmax·xn/(kn + xn); for secADH, using y = Vmax·x/(Km + x·(1 + x/Ki)).
Figure 7. Evaluating different concentrations for secADH and OhyRe in a coupled assay. Dark grey curve: fixed secADH concentration (0.01 g/L). Red curve: fixed OhyRe concentration (1 g/L). The response for OhyRe was fitted using y = Vmax·xn/(kn + xn); for secADH, using y = Vmax·x/(Km + x·(1 + x/Ki)).
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Figure 8. (a) Conversion of oleic acid into 10-HSA and 10-KSA using OhyRe and secADH over the course of 16 h. Reactions were performed in triplicates. Oleic acid was mixed with 50 mM MES buffer pH 6.5 to an emulsion of 10 mM, and 20 µL of it was added to a final concentration of 1 g/L OhyRe (with an equimolar concentration of FAD) and 0.01 g/L secADH, 20 µL NAD+ (10 mM), 1% (v/v) 1,5-pentanediol, and 50 mM MES buffer pH 6.5 to a total volume of 200 µL. Reactions were stopped with 1 mL ethyl acetate. Samples were analyzed using GC-FID. (b) Conversions (%) of 10-HSA and 10-KSA of in (a) shown reactions.
Figure 8. (a) Conversion of oleic acid into 10-HSA and 10-KSA using OhyRe and secADH over the course of 16 h. Reactions were performed in triplicates. Oleic acid was mixed with 50 mM MES buffer pH 6.5 to an emulsion of 10 mM, and 20 µL of it was added to a final concentration of 1 g/L OhyRe (with an equimolar concentration of FAD) and 0.01 g/L secADH, 20 µL NAD+ (10 mM), 1% (v/v) 1,5-pentanediol, and 50 mM MES buffer pH 6.5 to a total volume of 200 µL. Reactions were stopped with 1 mL ethyl acetate. Samples were analyzed using GC-FID. (b) Conversions (%) of 10-HSA and 10-KSA of in (a) shown reactions.
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Table 1. Fatty acid profiles of the oils used for the cascade reaction of a lipase (from Candida rugosa) and OhyRe in (%).
Table 1. Fatty acid profiles of the oils used for the cascade reaction of a lipase (from Candida rugosa) and OhyRe in (%).
Fatty AcidHigh-Oleic Sunflower OilOlive OilRapeseed OilMicrobial Oil from C.o.Linseed Oil
C16:02.6411.824.1436.485.40
C16:1-0.650.200.490.11
C18:01.502.751.6617.503.92
C18:1 (oleic acid)93.0775.0262.5743.1420.71
C18:22.218.5121.501.2815.44
C18:3-0.719.24-54.27
C20:00.180.540.580.150.16
C20:10.40-0.110.96-
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Prem, S.A.; Haack, M.; Melcher, F.; Ringel, M.; Garbe, D.; Brück, T. Two Cascade Reactions with Oleate Hydratases for the Sustainable Biosynthesis of Fatty Acid-Derived Fine Chemicals. Catalysts 2023, 13, 1236. https://doi.org/10.3390/catal13091236

AMA Style

Prem SA, Haack M, Melcher F, Ringel M, Garbe D, Brück T. Two Cascade Reactions with Oleate Hydratases for the Sustainable Biosynthesis of Fatty Acid-Derived Fine Chemicals. Catalysts. 2023; 13(9):1236. https://doi.org/10.3390/catal13091236

Chicago/Turabian Style

Prem, Sophia A., Martina Haack, Felix Melcher, Marion Ringel, Daniel Garbe, and Thomas Brück. 2023. "Two Cascade Reactions with Oleate Hydratases for the Sustainable Biosynthesis of Fatty Acid-Derived Fine Chemicals" Catalysts 13, no. 9: 1236. https://doi.org/10.3390/catal13091236

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