Article pubs.acs.org/OPRD
Efficient Biochemical Cascade for Accessing Green Leaf Alcohols Fredi Brühlmann* and Bojan Bosijokovic Firmenich SA, Corporate R&D, Route des Jeunes 1, CH-1211 Geneva 8, Switzerland ABSTRACT: The biotechnological production of six-carbon membered green leaf aldehydes and alcohols appreciated for their fruity, green-grassy odors in fragrances and flavors exploits the 13-lipoxygenase pathway found in higher plants. Homogenized seeds, fruits, or wild-type microorganisms have traditionally served as catalysts used in a one-pot reaction. However, many of these catalysts can be advantageously replaced by enzymes produced in engineered microorganisms for improved process performance. Substituting guava fruits by an engineered 13-hydroperoxide lyase produced in E. coli was recently shown to provide increased efficiency for cleaving fatty acid 13-hydroperoxides into green leaf aldehydes. Replacing baker’s yeast by a cosubstrate-dependent recombinant ketoreductase was found to offer superior productivity and chemo-selectivity in the reduction of hexenals to the corresponding alcohols. Finally, combining both cellular catalysts at reasonable loading in a simple one-pot cascade reaction offered the green leaf alcohol (Z)-3-hexenol at high isomeric purity (>99%) and high titers (8 g L−1) in spite of challenging, nonphysiological but otherwise benign reaction conditions. KEYWORDS: cascade reaction, biochemical, lipoxygenase, hydroperoxide lyase, ketoreductase
■
INTRODUCTION The use of biocatalysts, both as enzymes and also as engineered whole cells, for producing a wide range of chemicals is appealing for several reasons. Such processes are often carried out under benign conditions using nontoxic biocatalysts. Furthermore, biochemical conversions not based on living organisms can be carried out under nonphysiological reaction conditions in favor of high space time yields. If needed, biocatalysts can be engineered using a wide array of protein and strain engineering tools to fit process requirements.1−3 Consequently, biocatalysis has become an established complementary tool for the chemical industry.4,5 While synthetic chemistry offers access to a vast array of chemicals, exceptions remain particularly for complex molecular structures or in case of regulatory constraints (e.g. natural products). Similarly, isolation from natural sources may not always be feasible due to difficult sourcing of raw materials or due to low concentrations of target molecules that would inflict high costs and strong ecological footprints. Biocatalysis has a good chance to address many of these shortcomings. Leaf aldehydes and alcohols and their esters, also known as green notes, represent an interesting group of fatty acid derived chemicals appreciated for their green-grassy odors in flavors and fragrances.6,7 Six-carbon membered green leaf aldehydes and alcohols are formed in plants via a branch of the 13lipoxygenase pathway (Figure 1). Many of these chemicals play important roles in the plant’s defense against pathogens or are involved in chemical signaling between plants, or between plants and insects among others.8,9 The lipoxygenase branch toward green leaf aldehydes and alcohols starts from easily available polyunsaturated fatty acids such as α-linolenic acid or linoleic acid. It requires three enzymes including a 13lipoxygenase belonging to the nonheme-iron containing dioxygenases for regio- and stereospecific oxygenation,10 a 13hydroperoxide lyase (CYP74B) belonging to the superfamily of P450 enzymes for cleavage of fatty acid hydroperoxides,11 and finally an oxidoreductase for reducing the aldehydes.12 © XXXX American Chemical Society
Mechanistic work revealed that fatty acid hydroperoxide lyases isomerize fatty acid hydroperoxides into short-lived hemiacetals, thus could be considered as hemiacetal synthases.13,14 Different enzyme sources can be exploited. Early methods relied on the use of selected homogenized seeds, fruits, and wild type microorganisms as biocatalysts. For example, the 13lipoxygenase has been traditionally used in form of easily available ground soy beans, whereas homogenized plants such as guava, bell pepper, sugar beet, and so forth offered the 13hydroperoxide lyase.15−17 Baker’s yeast has traditionally served as catalyst for reducing carbonyl compounds. However, despite good availability these biocatalysts do not always provide high performance for several reasons. The enzyme of interest can be too diluted or may lack storage and process stability. Consequently, high biomass loading is required, which can negatively impact the process in addition to added costs. Furthermore, different enzyme activities are normally present in such biocatalyst competing for the same substrate leading to side products and reduced yields. Such drawbacks can be circumvented by the use of recombinant microorganisms as alternative enzyme sources.6 For example the application of a recombinant 13-hydroperoxide lyase expressed in E. coli was successfully applied for the cleavage of fatty acid hydroperoxides at preparative scale.18 In a more recent example it was shown that an engineered 13-hydroperoxide lyase offered improved process stability due to an increased total turnover number (TTN) and thermal stability that translated into lower catalyst loading.19 Similarly, the use of a ketoreductase overproduced in a microorganism instead of baker’s yeast might offer higher selectivity and increased volumetric yields at reduced catalyst loading. In this work we show that baker’s yeast can indeed be advantageously replaced by a recombinant ketoreductase relying on cosubstrate-dependent cofactor recycling for producing green leaf alcohols. Of particular Received: September 7, 2016
A
DOI: 10.1021/acs.oprd.6b00303 Org. Process Res. Dev. XXXX, XXX, XXX−XXX
Organic Process Research & Development
Article
Figure 1. 13-Lipoxygenase pathway toward six carbon membered, unsaturated aldehydes and alcohols starting from α-linolenic acid. Isomerization of (Z)-3-hexenal can be optionally carried out to access (E)-2-hexenal or the corresponding alcohol, respectively. A similar pathway from linoleic acid leads to hexanal and n-hexanol. Undesired side reactions are boxed that would also include oxidations of aldehydes into carboxylic acids (not shown). Enzyme abbreviations relate to 13-lipoxygenase (13-LOX), 13-hydroperoxide lyase (13-HPL), ketoreductase (KRED), isomerase (ISO), and enereductase (ENR).
interest was the concomitant use of the engineered 13hydroperoxide lyase and a cosubstrate-dependent ketoreductase in a one-pot cascade reaction. Cascade reactions are particularly appealing since they allow a sequence of reactions without the need of isolating the intermediate(s).20−22
■
RESULTS AND DISCUSSION Since ground soybeans offer an easily available, inexpensive, and relative robust catalyst showing high 13-lipoxygenase activity for producing fatty acid 13-hydroperoxides,15,23 priorities were given to the 13-hydroperoxide lyase mediated cleavage of the fatty acid hydroperoxide and the reduction of aldehydes to alcohols. The 13-hydroperoxide lyase present in guava fruits was known to be limiting due to its low concentration, seasonal variations, but also low storage and process stability as seen by its rapid inactivation during the reaction. Mechanism-based inactivation via adducts and destruction of the heme group by reaction intermediates, also termed “suicide inactivation”, is also common to many other P450s.24 These limitations motivated the cloning of the 13-hydroperoxide lyase of guava and its engineering. The enzyme was subsequently improved by directed evolution for increased process stability.19 Immobilization of the enzyme as recently shown for the 13-hydroperoxide lyase of A. tricolor could also be considered.25 For in situ reduction of (Z)-3-hexenal a commercially available isopropanol-accepting ketoreductase (ADH005 from Codexis, Redwood City, USA) was tested. This enzyme showed excellent activity toward (E)-2-hexenal in a buffered system offering high space time yields of about 100 g L−1 h−1 in batch reactions (Figure 2). The excess of the cosubstrate isopropanol and the removal of acetone under reduced pressure drove the reduction to completion. Such high space time yields would have been impossible with baker’s yeast as the catalyst, which showed partial conversions at much lower substrate concentrations but also the formation of n-hexanol due to the presence of an enereductase (data not shown). We then tested whether this isopropanol-accepting ketoreductase could also accept (Z)-3-hexenal as substrate formed during the cleavage reaction. Because it was uncertain whether
Figure 2. Reduction of (E)-2-hexenal in a buffer solution with the ketoreductase ADH005 using isopropanol as the hydride donor under reduced pressure for acetone removal. Symbols: (E)-2-hexenol (red ■), (E)-2-hexenal (−), acetone (green ●).
the 13-hydroperoxide lyase would function in the presence of isopropanol, we first tested the sequential cleavage and reduction by the timely separated addition of enzymes. In this system the cleavage reaction was allowed to proceed in the absence of isopropanol. After completion of the cleavage reaction, isopropanol, the cofactor (NADP+), and the ketoreductase ADH005 were then added for reducing the aldehydes (Figure 3A). In this system the (Z)-3-hexenal accumulated rapidly during the cleavage phase enabled by the high turnover number (kcat) of the 13-hydroperoxide lyase estimated at >2000 s−1.26 Some isomerization of (Z)-3-hexenal into the more stable (E)-2-hexenal occurred, though less than 0.2 g L−1 of (E)-2-hexenal were usually formed under these conditions. The subsequent reduction of the unstable (Z)-3hexenal with the isopropanol-dependent ketoreductase proceeded also very well. Therefore, the presence of nonreacted fatty acid hydroperoxides and free fatty acids had no detrimental effect on the ketoreductase. Some residual (Z)-3hexenal remained at the end of the reaction, although complete reduction could be expected after further improvements of the reaction conditions. We then attempted the synthesis of (Z)-3-hexenol in a concomitant cascade reaction. In this case both enzymes were present to allow formation of the aldehydes and their B
DOI: 10.1021/acs.oprd.6b00303 Org. Process Res. Dev. XXXX, XXX, XXX−XXX
Organic Process Research & Development
Article
Figure 4. Inhibitory effect of isopropanol during the cleavage of the 13-hydroperoxide of α-linolenic acid by the 13-hydroperoxide lyase.
hydroperoxide of α-linolenic acid over the 13-hydroperoxide of linoleic acid, which is also present in the fatty acid hydroperoxide made from crude α-linolenic acid, as previously reported.26 Other endogenous enzymes present in soybeans and in E. coli such as ene-reductases, aldehyde dehydrogenases, or alcohol oxidases did not play a noticeable role since the corresponding side products were not detected. The amounts of recombinant 13-hydroperoxide lyase and of the cosubstratedependent ketoreductase in form of bacterial cell suspensions were low and represented a few percent of the reaction volume only. Such low loadings have not been possible with homogenized fruits or baker’s yeast as catalyst requiring up to 50% of the reactor volume. Despite low catalyst loadings high space time yields (>8 g L −1 h−1 ) were obtained in nonoptimized batch reactions. These productivities were significantly higher than what has been reported thus far.15,18,23 The molar yield was 41%. Considerably higher yields were observed at lower substrate concentrations (data not shown) that is in line with previous reports.18,27 The difficulties associated with the use of baker’s yeast (low space time yields, side reactions, etc.) were all greatly avoided by the highly active, cosubstrate-dependent, cell-based recombinant ketoreductase. A comparison of pros and cons of traditionally used wild-type biocatalysts compared to recombinant microorganisms overproducing one or more target enzymes is illustrated for carbonyl reductions in Table 1. Furthermore, the biocatalysts used in form of cell suspensions showed excellent storage stabilities easily exceeding more than one year when stored frozen. From a processing point of view the concomitant cleavage and reduction appears easier and more reliable than the sequential cleavage and reduction by the timely segregated addition of biocatalysts. In the latter, minor variations in the timing of the addition of biocatalysts and reactants could lead to variability in yield and isomeric purity of (Z)-3-hexenol. The concomitant cleavage and reduction also reduces the incidence of isomerization of the unstable (Z)-3-hexenal. The described use of single-enzyme-biocatalysts has so far outperformed more complex catalysts based on the coexpression of more than one enzyme in the same host.28 Cytotoxic effects of fatty acids and of fatty acid hydroperoxides have also been reported when working with catalysts based on living cells.29 However, the extent of cytotoxic effects may depend on the host organism used.30 The coexpression of biosynthetic enzymes will require careful fine-tuning to avoid accumulation of inhibitory or cytoxic intermediates. Nevertheless, such cell factories show high potential for accessing “green notes” and many other chemicals from a simple carbon source.
Figure 3. Comparison of sequential (A) and concomitant cleavage and reduction (B) involving the engineered 13-HPL and the ketoreductase ADH005. In experiment (A) the cleavage reaction had already proceeded for 5 min prior to the addition of the ketoreductase, isopropanol and cofactor at t = 0 min. Symbols: (Z)-3-hexenol (blue ◆), (E)-2-hexenol (red ■), (Z)-3-hexenal (green ▲), (E)-2-hexenal (−).
concomitant reduction. Thus, the 13-hydroperoxide lyase and the ketoreductase were basically added at the same time to the reaction containing the fatty acid hydroperoxides, isopropanol, and the cofactor. The cofactor was added in minute amounts only (0.1 mM) relying on a TTN of at least 1600. This low concentration of the cofactor is not irrelevant when considering its high costs. This reaction proceeded also very well as seen by the rapid accumulation of (Z)-3-hexenol (Figure 3B). The (Z)3-hexenal was transiently formed at low concentrations thus keeping the risk of its isomerization low. The biochemical reduction went to completion with no aldehydes left and with very little (E)-2-hexenol as a side product ( 80 g L−1 and 0.2 mL of lysed cells of E. coli (cell suspension equivalent to OD600 = 10) containing the 13HPL. After 5 min 400 μL of isopropanol, 25 μL of NADP+ (50 mM), and 120 μL of the enzyme ADH005 (775 U mL−1) of Codexis (Redwood City, California) were added under stirring at room temperature. Samples of 100 μL were withdrawn from the reaction after 2 min, 5 min, 10 min, 20 min, 40 min, 60 min, and immediately diluted with 900 μL of water. Extraction was with 1 volume of ethyl acetate containing 1 g L−1 of octanol as the internal standard for analysis by gas chromatography. Concomitant Cleavage of 13-HPOT/D and Reduction of (Z)-3-Hexenal. Into a 10 mL vial were added: 1800 μL of 13-HPOT/D of 80 g L−1, 4 μL of 1 M MgSO4, 25 μL of 50 mM NADP+, 400 μL of isopropanol, 120 μL of the ketoreductase ADH005 (775 U mL−1) of Codexis (Redwood City, California), and 250 μL of an E. coli lysate (cell suspension equivalent to OD600 = 10) containing the 13-HPL (variant GC7). The reaction was stirred at room temperature. Samples of 100 μL were withdrawn from the reaction at fixed time points during 1 h, diluted with 900 μL of H2O, and extracted with 1 mL of ethyl acetate prior to analysis by gas chromatography as described in the previous example. Octanol served as the internal standard. Effect of Isopropanol on the Cleavage Kinetics of the 13-HPL. Cleavage reactions were set up containing 5, 10, 20, and 40 v/v% of isopropanol. In a separate series, reactions were run containing the same amount of added water instead of isopropanol with the volumes for the substrate and the enzyme adjusted accordingly. Reactions were left under magnetic stirring at room temperature. Aliquots of 100 μL were taken after 2 min, diluted with 900 μL of NaBH4 (10 g L−1), and left under stirring for 5 min prior to extraction with ethyl acetate for analysis by GC. Individual reactions with and without isopropanol were compared for estimating the inhibitory effect of isopropanol. Preparation of (Z)-3-Hexenol. Into a 1 L flask were added 692 mL of 13-HPOT/D > 80 g L−1, 1.5 mL of 1 M of MgSO4, 155 mL of isopropanol, 20 mL of 25 mM of NADP+, 46 mL of the ketoreductase ADH 005 (775 U mL−1) of Codexis, and 97 mL of lysed cells of E. coli containing the 13-HPL variant GC7 (cell suspension equivalent to OD600 = 7). The reaction was agitated with a magnetic stirrer at 800 rpm at room temperature for 40 min. Aliquots of 200 μL were withdrawn after 2, 5, 10, 20, and 40 min to monitor the reaction via gas chromatography. The entire reaction was then extracted 3× with MTBE. The organic extract was washed with water, dried over Na2SO4, and filtered with a PTFE membrane of 0.45 μm (Sartorius AG, Germany) prior to the removal of the organic solvent using the
Table 1. Comparison of Process Relevant Criteria for Carbonyl Reductions in Vitro with Baker’s Yeast and Cell Based Recombinant Ketoreductases As Catalysts baker’s yeast (wild type) space time yield chemo-selectivity regio-selectivity stereoselectivity cofactor regeneration biomass/protein loading viscosity process control scale-up catalyst availability down stream processing catalyst costs biosafety level
recombinant KRED (cell based)
often low (inhibitions, low substrate loading etc.) often low (competing enzymes) often low often low generally in vivo (endogenous cofactor regeneration) often high
high (>100 g L−1 h−1)
often high (high biomass loading) often difficult (competing enzymes etc.) can be tricky unlimited
low
affected by viscosity, side products etc.
requires cosubstrate and coproduct removal (often by distillation) low
often high (high biomass loading, low yield) 1
high high high in vivo or in vitro (cosubstratedependent regeneration preferred) low
simple and reproducible simple and reproducible unlimited (if freedom to operate)
1
■
CONCLUSION The green leaf alcohol (Z)-3-hexenol was obtained in a simple one-pot cascade reaction by enzyme catalyzed cleavage of a fatty acid 13-hydroperoxide using an engineered 13-hydroperoxide lyase (CYP74B) followed by concomitant reduction of the unstable aldehyde with a cosubstrate-dependent ketoreductase. Transiently formed unstable (Z)-3-hexenal was kept low to avoid its isomerization thanks to concomitant reduction into the stable (Z)-3-hexenol. In spite of harsh reaction conditions both enzymes performed astonishingly well. The method described is simple, robust, and scalable and offers the desired green leaf alcohol (Z)-3-hexenol at high isomeric purity at economically viable product titers. The methods also illustrates one of the still few examples for the preparative use of an engineered biocatalyst of the highly versatile cytochrome P450 superfamily.
■
EXPERIMENTAL SECTION Reduction (E)-2-Hexenal with the Ketoreductase ADH005. 952 μL of 400 mM Tris pH 8.2, 2500 μL of isopropanol, 620 mg of (E)-2-hexenal, 250 μL of 10 mM NADP+, 25 μL of 1 M MgCl2, and 666 μL of ADH005 (1550 U mL−1) of Codexis (Redwood City, California) were added to a rotating flask connected to an evaporator (Rotavapor R210, Büchi, Switzerland). The reaction was left at 26 °C and under 60 mbar for removing the acetone formed. At given time intervals aliquots were taken and extracted with ethyl acetate for analysis by GC. The mass of the reaction was followed gravimetrically. Synthesis of 13-Fatty Acid Hydroperoxides (13-HPOT/ D). The synthesis of crude 13-fatty acid hydroperoxides was prepared from a linseed hydrolysate containing 48.3% of αlinolenic acid and 16.3% of linoleic acid as described.19 Production of Cell-Free Crude 13-Hydroperoxide Lyase. 0.5 mL of an overnight culture of cells of E. coli harboring a recombinant expression plasmid with the D
DOI: 10.1021/acs.oprd.6b00303 Org. Process Res. Dev. XXXX, XXX, XXX−XXX
Organic Process Research & Development
Article
(20) Ricca, E.; Brucher, B.; Schrittwieser, J. H. Adv. Synth. Catal. 2011, 353, 2239. (21) Garcia-Junceda, E.; Lavandera, I.; Rother, D.; Schrittwieser, J. H. J. Mol. Catal. B: Enzym. 2015, 114, 1. (22) Riva, S.; Fessner, W.-D. Cascade Biocatalysis; John Wiley & Sons: New York, 2014. (23) Whitehead, I.; Muller, B. L.; Dean, C. Cereal Foods World 1995, 194, 193. (24) Matsui, K.; Miyahara, C.; Wilkinson, J.; Hiatt, B.; Knauf, V.; Kajiwara, T. Biosci., Biotechnol., Biochem. 2000, 64, 1189. (25) Liu, Q.; Kong, X.; Zhang, C.; Chen, Y.; Hua, Y. J. Sci. Food Agric. 2013, 93, 1953. (26) Tijet, N.; Wäspi, U.; Gaskin, D. J. H.; Hunziker, P.; Muller, B. L.; Vulfson, E. N.; Slusarenko, A.; Brash, A. R.; Whitehead, I. M. Lipids 2000, 35, 709. (27) Jacopini, S.; Mariani, M.; Brunini-Bronzini de Caraffa, V.; Gambotti, C.; Vincenti, S.; Desjobert, J.-M.; Muselli, A.; Costa, J.; Berti, L.; Maury, J. Appl. Biochem. Biotechnol. 2016, 179, 671. (28) Buchhaupt, M.; Guder, J. C.; Etschmann, M. M. W.; Schrader, J. Appl. Microbiol. Biotechnol. 2012, 93, 159. (29) Thanh, H. T.; Vergoignan, C.; Cachon, R.; Kermasha, S.; Gervais, P.; Nguyen, T. X. S.; Belin, J.-M.; Husson, F. J. Mol. Catal. B: Enzym. 2008, 52−53, 146. (30) Bourel, G.; Nicaud, J. M.; Nthangeni, B.; Santiago-Gomez, P.; Belin, J. M.; Husson, F. Enzyme Microb. Technol. 2004, 35, 293.
Rotavapor R-210 (Büchi, Switzerland). The residue was then distilled in a micro distillation apparatus at 70−85 °C under vacuum (15 mbar). The identity of the product was verified by GC-MS and NMR. Gas Chromatography. A gas chromatograph HP6890 (Hewlett-Packard, USA) equipped with a flame ionization detector and a DB-WAX column (L = 30 m, ID = 0.25 mm, coating = 0.25 μm) was used following the temperature program: 80 °C (2 min), 160 °C (4 °C min−1), 230 °C (30 °C min−1), 230 °C (6 min). The helium flux was 1.4 mL min−1 using a split ratio of 1:50.
■
AUTHOR INFORMATION
Corresponding Author
*E-mail: fredi.bruhlmann@firmenich.com. Notes
The authors declare no competing financial interest.
■
ACKNOWLEDGMENTS We are thankful to Michiel Hacking of Codexis, current address: European Patent Office, The Hague, The Netherlands, for having initially provided the ketoreductase ADH005.
■
REFERENCES
(1) Bornscheuer, U. T.; Huisman, G. W.; Kazlauskas, R. J.; Lutz, S.; Moore, J. C.; Robins, K. Nature 2012, 485, 185. (2) Opgenorth, P. H.; Korman, T. P.; Bowie, J. U. Nat. Chem. Biol. 2016, 12, 393. (3) Denard, C. A.; Ren, H.; Zhao, H. Curr. Opin. Chem. Biol. 2015, 25, 55. (4) Torrelo, G.; Hanefeld, U.; Hollmann, F. Catal. Lett. 2015, 145, 309. (5) Whitthall, J.; Sutton, P. W. Practical Methods for Biocatalysis and Biotransformations, Vol. 1 & 2; John Wiley & Sons: New York, 2012. (6) Gigot, C.; Ongena, M.; Fauconnier, M. L.; Wathelet, J. P.; Du Jardin, P.; Thonart, P. Biotechnol. Agron. Soc. Environ. 2010, 14, 451. (7) Hatanaka, A. Food Rev. Int. 1996, 12, 303. (8) Santino, A.; Bonsegna, S.; De Domenico, S.; Poltronieri, P. Curr. Top. Plant Biol. 2010, 11, 103. (9) Griffiths, G. Free Radical Res. 2015, 49, 565. (10) Feussner, I; Kühn, H. Enzymes in Lipid Modification: Application of lipoxygenases and related enzymes for the preparation of oxygenated lipids; Wiley-VCH Verlag GmbH & Co: Weinheim, 2000. (11) Hughes, R. K.; De Domenico, S.; Santino, A. ChemBioChem 2009, 10, 1122. (12) De Wildeman, S. M. A.; Sonke, T.; Schoemaker, H. E.; May, O. Acc. Chem. Res. 2007, 40, 1260. (13) Grechkin, A. N.; Hamberg, M. Biochim. Biophys. Acta, Mol. Cell Biol. Lipids 2004, 1636, 47. (14) Grechkin, A. N.; Brühlmann, F.; Mukhtarova, L. S.; Gogolev, Y. V.; Hamberg, M. Biochim. Biophys. Acta, Mol. Cell Biol. Lipids 2006, 1761, 1419. (15) Muller, B. L.; Dean, C.; Whitehead, I. M. Bioflavour 95: The industrial use of plant enzymes for the production of natural ‘green note’ flavor compounds; INRA: Paris, 1995. (16) Németh, Á . S.; Márczy, J. S.; Samu, Z.; Háger-Veress, Á .; Szajáni, B. Enzyme Microb. Technol. 2004, 34, 667. (17) Rabetafika, H. N.; Gigot, C.; Fauconnier, M.-L.; Ongena, M.; Destain, J.; du Jardin, P.; Wathelet, J.-P.; Thonart, P. Biotechnol. Lett. 2008, 30, 1115. (18) Noordermeer, M.; van der Goot, W.; van Kooij, A. J.; Veldsink, J. W.; Veldink, G. A.; Vliegenthart, J. F. G. J. Agric. Food Chem. 2002, 50, 4270. (19) Brühlmann, F.; Bosijokovič, B.; Ullmann, C.; Auffray, P.; Fourage, L.; Wahler, D. J. Biotechnol. 2013, 163, 339. E
DOI: 10.1021/acs.oprd.6b00303 Org. Process Res. Dev. XXXX, XXX, XXX−XXX
