Johnson Matthey Technol. Rev., 2020, 64, (4), 529
Biocatalytic Reduction of Activated Cinnamic Acid Derivatives
Asymmetric reduction of C=C double bonds using Johnson Matthey enzymes
The asymmetric reduction of C=C double bonds is a sought-after chemical transformation to obtain chiral molecules used in the synthesis of fine chemicals. Biocatalytic C=C double bond reduction is a particularly interesting transformation complementary to more established chemocatalytic methods. The enzymes capable of catalysing this reaction are called ene-reductases (ENEs). For the reaction to take place, ENEs need an electron withdrawing group (EWG) in conjugation with the double bond. Especially favourable EWGs are carbonyls and nitro groups; other EWGs, such as carboxylic acids, esters or nitriles, often give poor results. In this work, a substrate engineering strategy is proposed whereby a simple transformation of the carboxylic acid into a fluorinated ester or a cyclic imide allows to increase the ability of ENEs to reduce the conjugated double bond. Up to complete conversion of the substrates tested was observed with enzymes ENE-105 and *ENE-69.
The use of enzymes for the asymmetric reduction of activated C=C double bonds can be a viable and straightforward alternative to asymmetric hydrogenation. Traditionally, whole cell microorganisms were used for this purpose but a recent increase in the number of isolated and characterised ENEs means that recombinantly-expressed enzyme preparations are now generally favoured over whole cells, as a number of recent publications demonstrate (1–10).
Double bond ‘activation’ to facilitate ENEs mediated reduction can be achieved in many cases by alpha substituted functional groups including aldehydes, ketones or nitro moieties. Carboxylate derivatives (such as esters, lactones and anhydrides) can also act as activating groups but their ability to sufficiently activate the C=C bond in the absence of other groups is less evident (11, 12). The traditional approach in these cases is to turn to chemocatalytic hydrogenation (see (13–15) for reviews focused on industrial applications). Herein we describe a new approach to activate α,β-unsaturated carboxylic acids for the reduction with ENEs using a substrate engineering approach.
All reagents and solvents were purchased from Sigma-Aldrich and Alfa Aesar, Thermo Fisher Scientific. They were of the highest available purity and were used without further purification. 1H nuclear magnetic resonance (NMR) spectra were recorded using a Bruker 400 MHz Avance III HD equipped with SMART probe (Bruker Corporation, USA) where spectra are referenced to deuterated chloroform (CDCl3) 7.26 ppm, shifts are recorded in parts per million and J values in hertz. The NMR results can be found in the Supplementary Information.
2.2 Enzyme Preparations
Genes coding for Johnson Matthey, ENEs (ENE-101, ENE-102, ENE-103, ENE-104, ENE-105, *ENE-69 and GDH-101) were ordered codon-optimised from GeneArt (Thermo Fisher Scientific) and cloned into T5 vector pJEx401 (ATUM). Enzymes were expressed recombinantly in Escherichia coli BL21 in both shake flasks and fed batch fermentations, whereby induction was carried out with isopropyl β-D thiogalactopyranoside (IPTG) at 30°C. Harvested biomass was resuspended in 100 mM potassium phosphate buffer (pH 7) and cells were broken up either by sonication or homogenisation. The so-obtained cell lysate was clarified by centrifugation and filtrated prior to lyophilisation. Protein expression was assessed by sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) and chromatographic activity assays.
Enzymes ERED-103, ERED-110, ERED-112, ERED-207, ERED-P1-A04, ERED-P1-E04 and ERED-P1-H09 were purchased from Codexis.
2.3 2,2,2-Trifluoroethyl Cinnamate (3a) and 3-Phenyl-Acrylic Acid 2,2,2-Trifluoro-1-Trifluoromethyl-Ethyl Ester (5a)
Cinnamic acid 1a (5 g, 33.75 mmol) and oxalyl chloride (2.85 ml, 33.75 mmol) in dichloromethane (5 ml) were stirred at 25°C for 2 h before adding the fluorinated alcohol-trifluoro ethanol for 3a (2.47 ml, 33.75 mmol) and 1,1,1,3,3,3-hexafluoropropan-2-ol for 5a (3.50 ml, 33.75 mmol). The reaction was then stirred at room temperature overnight before being quenched by addition of saturated aqueous NaHCO3 (20 ml) and extracted with dichloromethane (2 × 20 ml), dried over MgSO4, filtered and concentrated under reduced pressure to afford the corresponding fluorinated esters 3a and 5a in quantitative yield.
2.4 3-Phenyl-Acrylic Acid 2,2,3,3,4,4,4-Heptafluoro-Butyl Ester (6a) and (Perfluorophenyl)Methyl Cinnamate (7a)
Cinnamoyl chloride (0.75 g, 4.50 mmol) and the corresponding fluorinated alcohols – 2,2,3,3,4,4,4-heptafluorobutan-1-ol for 6a (0.98 g, 4.50 mmol) and pentafluroro benzyl alcohol for 7a (0.89 g, 4.50 mmol) – in dichloromethane (2.5 ml) were stirred at room temperature overnight. The reaction was then quenched by addition of saturated aqueous NaHCO3 (20 ml) and extracted with dichloromethane (2 × 20 ml), dried over MgSO4, filtered and concentrated under reduced pressure to afford the corresponding fluorinated esters 6a and 7a in 95% to 99% yield.
2.5 1-Cinnamoylpyrrolidin-2-one (9a)
Cinnamoyl chloride (5 g, 30.01 mmol), pyrrolidinone (2.3 ml, 36.01 mmol) and triethylamine (13 ml, 90.03 mmol) in dichloromethane (50 ml) were stirred at room temperature overnight. The reaction was quenched by addition of water (20 ml), the organic layer was separated and washed with saturated aqueous NaCl (20 ml), dried over MgSO4, filtered and concentrated under reduced pressure to afford 9a in 81% yield.
2.6 3-Cinnamoyloxazolidin-2-one (8a)
Cinnamic acid 1a (5 g, 33.56 mmol) and oxalyl chloride (2.85 ml, 33.56 mmol) in dichloromethane (5 ml) were stirred at room temperature overnight before removing the solvent under reduced pressure. The reaction crude was dissolved in anhydrous tetrahydrofuran (THF) (20 ml) and n-butyllithium (1.6 M in hexane, 21 ml, 33.56 mmol, one equivalent) was added dropwise over 30 min. The cinnamoyl chloride solution was then added dropwise to a solution of oxazolidinone (2.92 g, 33.56 mmol) in anhydrous THF (100 ml) at 0°C before stirring at room temperature overnight. The reaction was quenched with water (50 ml), extracted with ethyl acetate (EtOAc) (2 × 100 ml), washed with saturated aqueous NaHCO3 (20 ml) and saturated aqueous NaCl (20 ml). The solvent was removed under reduced pressure and the solid was recrystallised from a 1:1 mixture EtOAc:heptane (20 ml). The solid was filtered and washed with hexane (10 ml) to give crystals of 8a in 80% yield.
2.7 (E)-1-(2-Methyl-3-Phenylacryloyl)Pyrrolidin-2-one (10a) and (E)-1-(2,3-Diphenylacryloyl)Pyrrolidin-2-one (11a)
(E)-2-methyl-3-phenylacrylic acid (5 g, 30.86 mmol) was converted to the corresponding acid chloride by addition of oxalyl chloride (1.4 ml, 30.86 mmol) in dichloromethane (5 ml). The reaction was stirred at room temperature for 3 h. Pyrrolidinone (2.82 ml, 37.03 mmol) and triethylamine (13 ml, 92.58 mmol) were added before stirring the reaction overnight. The reaction was quenched by addition of water (20 ml) and saturated aqueous NaCl (20 ml). The solvent was removed under reduced pressure and the solid was dissolved in EtOAc and treated with activated charcoal (1 g), filtered through Celite® and concentrated. The solid was recrystallised from heptane (10 ml) to give 10a in 55% yield.
Following an identical procedure, 11a was synthesised in 53% yield from (E)-2,3-diphenylacrylic acid (10 g, 44.64 mmol).
2.8 Small Scale Screening Reactions
Substrates 1a–9a (0.025 mmol) and enzymes ENE-101, ENE-102, ENE-103, ENE-104, ENE-105 or *ENE-69 (2.5 mg), were added to reaction vials containing 500 μl of aqueous media at pH 7 (250 mM potassium phosphate buffer pH 7, 1.1 mM NAD(P)+, 100 mM D-glucose, 10 U ml−1 GDH-101) to give a final concentration of substrate of 50 mM. The vials were shaken at 400 rpm, 30°C for 18 h. For high-performance liquid chromatography (HPLC) analysis, the reactions were quenched with acetonitrile (MeCN) (1 ml), vortexed, centrifuged and aliquoted. For gas chromatography (GC) analysis, samples were extracted with EtOAc (2 × 0.5 ml), dried over MgSO4 and analysed directly. For NMR analysis, the reactions were extracted with CDCl3 and analysed directly.
2.9 Preparative Scale Screening Reactions
Reactions were scaled up using three-neck round bottom flask equipped with stir bar and pH titrator (10 M NaOH). To the flask was weighed 100–500 mg substrate (40–100 mM final concentration) and 5 mg ml−1 enzyme which was suspended in aqueous media at pH 7 (250 mM potassium phosphate buffer pH 7, 1.1 mM NAD(P)+, 100–200 mM D-glucose (two equivalent), 10 U ml−1 GDH-101) the reactions were stirred at 30°C, 400 rpm for 18 h.
2.10 Analytical Methods
HPLC analysis of conversion was conducted on an 1260 Infinity II LC system (Agilent, USA) using a C18 SunFire Column (Waters Corporation, USA, 150 × 4.6 mm, 3.5 μm) with an isocratic method (MeCN:Water, 30:70 + 0.1% trifluoroacetic acid) and a flow rate of 1 ml min−1.
Chiral HPLC analysis was performed on a Varian ProStar series (Agilent) with a CHIRALCEL® OD-H column (Chiral Technologies, USA, 250 × 4.6 mm, 5 μm) with an isocratic method A (heptane:isopropyl alcohol (IPA), 88:12) and a flow rate of 1 ml min−1 or isocratic method B (heptane:IPA, 98:2).
GC analysis of conversion was performed on a Varian CP-3800 (Agilent) using γ-DEX™ 225 capillary column (Sigma-Aldrich, 30 m × 0.25 mm × 0.25 μm) and using helium as carrier gas. Percentage conversion was measured by integration of the product peak in the GC (uncorrected area under curve (AUC)), values below 100% indicate that unreacted starting material was detected. No side products were detected in any of the reported reactions. GC program parameters: injector 250°C, flame ionization detector (FID) 250°C, 80°C for 3 min then 5°C min−1 up to 160°C, hold 1 min (total time 20 min), constant flow 5 ml min−1.
3. Results and Discussion
It has been found that a particular ENE in Johnson Matthey’s collection, a homologue from the tobacco ENE reductase fold (16), ENE-105, was capable of reducing methyl ester 2a (Figure 1), albeit in a very low yield of 3% (Entry 2, Table I). By comparison, cinnamic acid 1a was a poor substrate and showed no conversion to the reduced product 1b at pH 7.0 (Entry 1, Table I). The pKa of cinnamic acid 1a is 4.4 and therefore, at pH 7.0, the carboxylic acid should be deprotonated affecting its ability to bind to the enzyme active site. This observation is in line with other literature examples where carboxylates were found to be poor activating groups (17). Encouraged by this initial result, we turned our efforts towards the use of more activated esters. It was envisaged that converting the alkyl chain in the ester moiety to a more EWG could lead to an increase in double bond activation. A similar approach has been reported previously by BASF SE for the lipase-catalysed kinetic resolution of racemic amines and alcohols, where the choice of acylating agent proved critical (18). We chose trifluoroethyl ester 3a as a starting point which was reduced by ENE-105 and *ENE-69 in 6% and 12% conversion respectively (Entry 3, Table I) suggesting that the addition of an EWG had a positive activating-effect on the reduction. To consolidate this theory, ethyl ester 4a was tested with the novel ENEs; only a trace of reduction was observed <0.5% (Entry 4, Table I).
aIntegration of the product peak in the GC (uncorrected AUC), values below 100% indicate that unreacted starting material was detected; no side products were detected for these reactions
bIntegration of the product peak in the HPLC (achiral method, uncorrected AUC), values below 100% indicate that unreacted starting material was detected; no side products were detected for these reactions
Other EWGs such as hexafluoroethyl in compound 5a, heptafluorobutyl in 6a and pentafluorobenzyl in 7a could also activate the double bond in the same way, so 5a, 6a and 7a were prepared by reacting cinnamoyl chloride with the corresponding fluorinated alcohols and these substrates were subsequently tested with the ENEs. Hexafluoro 5a was not reduced by ENE-105 or *ENE-69 (Entry 5, Table I), instead, a significant amount of hydrolysis product (cinnamic acid 1a, 10%) was observed. Heptafluorobutyl 6a and pentafluoro 7a were poor activating groups with 6a showing only a trace amount of product 6b (Entry 6, Table I) and 7a giving no conversion (Entry 7, Table I).
With only limited success with the fluorinated activating groups, our efforts turned towards cyclic imides since activated substrates 8a and 9a have been shown to be highly activated towards Michael addition reactions (19, 20, 21) (Figure 2). Compounds 8a and 9a were synthesised and tested with enzymes ENE-105 and *ENE-69. Pleasingly, oxazolidinone 8a was successfully reduced by both ENEs (51% and 39% conversion to 8b, Entry 1, Table II) and pyrrolidinone 9a was reduced to 9b in >95% conversion (Entry 2, Table II), proving to be an excellent activating group. The 1H NMR shift of the alkene proton alpha to the carbonyl for pyrrolidinone 9a is shifted down field (7.92 ppm) compared to cinnamic acid 1a (6.46 ppm), therefore supporting the electron-withdrawing nature of the activating group.
The enzymes were then tested for their ability to reduce α-substituted cinnamic acid derivatives such as α-methyl 10a and α-phenyl 11a (Figure 3). Encouragingly, the tri-substituted double bond in 10a was reduced to 10b in >95% conversion by 1H NMR analysis (Entry 2, Table III). However, bulkier substrate 11a, was not tolerated so well on an analytical scale due to solubility issues causing mass-transfer limitations (Entry 3, Table III). The reaction was repeated on a larger scale with stirring (Entry 4, Table III) and >95% conversion was achieved. 10b and 11b were obtained as racemic mixtures.
With a successful activating group found, the reaction was repeated on a preparative scale to test reproducibility and scalability (Table IV). Pyrrolidinone 9a was successfully reduced using enzyme ENE-105 at 130 mg scale with the desired product 9b being obtained in 95% conversion by 1H NMR (Entry 1, Table IV). 72% conversion to 10b was achieved after 20 h (Entry 3, Table IV) on the reduction of pyrrolidinone 10a at 500 mg scale.
|Entry||Substrate||Scale, mg||Concentration, mM||Time, h||Conversion, %a|
Having found enzymes in Johnson Matthey’s collection that could successfully reduce masked carboxylic acids, other commercially available enzymes were tested as a comparison on the reduction of 10a (Table V). Six enzymes from Johnson Matthey collection (Entries 3 to 6, Table V) and seven enzymes purchased from Codexis (Entries 7 to 13, Table V) were compared with ENE-105 and ENE-69* (Entries 1 and 2, Table V). It was found that, despite the extra activation of the C=C double bond, none of the tested enzymes could reduce cinnamic acid derivative 10a, highlighting the unique ability of ENE-105 and *ENE-69 within the focused library (13 enzymes) screened.
In summary, we have shown that cinnamic acid derivatives activated as fluorinated esters or as cyclic imides can be reduced using Johnson Matthey enzymes ENE-105 or *ENE-69. The concept of ‘substrate engineering’ as opposed to ‘enzyme engineering’, offers a complimentary and faster approach to developing a bioprocess, making difficult transformations possible. The reduced products can be subsequently converted to the parent carboxylic acids by LiOH hydrolysis (22, 23) and the potential re-use of these activating groups will be investigated in the future. It is envisaged that the work will lead to further examples of activated acids or esters being reduced by ENEs.
The biocatalysed reduction of the double bond of cinnamic acid derivatives is strongly influenced by the nature of the EWG. While no conversion was observed on the biocatalysed reduction of cinnamic acid 1a, an enzyme in Johnson Matthey’s collection, ENE-105, was capable of reducing methyl ester derivative 2a in low conversion. By replacing the alkyl chain in the ester moiety by a more EWG, such as fluorinated alkanes, and in the presence of enzymes ENE-105 and *ENE-69, we were able to significantly increase conversion to the reduced product. Furthermore, other electronegative derivatives such as cyclic imides proved to be even better activating groups, allowing the reduction of challenging substituted double bonds such as substrates 10a and 11a.
In summary, by ‘masking’ the carboxylic acid moiety into a fluorinated alkyl ester or a cyclic imide, following a straightforward synthetic procedure, and in combination with the right enzyme, it was possible to biocatalytically reduce the conjugated double bond of cinnamic acid and substituted derivatives.
B. Dominguez, U. Schell, S. Bisagni and T. Kalthoff, Johnson Matthey Technol. Rev., 2016, 60, (4), 243 LINK https://www.technology.matthey.com/article/60/4/243-249/
M. Hall, C. Stueckler, H. Ehammer, E. Pointner, G. Oberdorfer, K. Gruber, B. Hauer, R. Stuermer, W. Kroutil, P. Macheroux and K. Faber, Adv. Synth. Catal., 2008, 350, (3), 411 LINK https://doi.org/10.1002/adsc.200700458
M. Hall, C. Stueckler, W. Kroutil, P. Macheroux and K. Faber, Angew. Chem., Int. Ed., 2007, 46, (21), 3934 LINK https://doi.org/10.1002/anie.200605168
J. F. Chaparro-Riggers, T. A. Rogers, E. Vazquez-Figueroa, K. M. Polizzi and A. S. Bommarius, Adv. Synth. Catal., 2007, 349, (8–9), 1521 LINK https://doi.org/10.1002/adsc.200700074
A. Müller, B. Hauer and B. Rosche, Biotechnol. Bioeng., 2007, 98, (1), 22 LINK https://doi.org/10.1002/bit.21415
M. A. Swiderska and J. D. Stewart, J. Mol. Catal. B: Enzym., 2006, 42, (1–2), 52 LINK https://doi.org/10.1016/j.molcatb.2006.06.023
D. Dobrijevic, L. Benhamou, A. E. Aliev, D. Méndez-Sánchez, N. Dawson, D. Baud, N. Tappertzhofen, T. S. Moody, C. A. Orengo, H. C. Hailes and J. M. Ward, RSC Adv., 2019, 9, (63), 36608 LINK https://doi.org/10.1039/c9ra06088j
H. S. Toogood and N. S. Scrutton, ACS Catal., 2018, 8, (4), 3532 LINK https://doi.org/10.1021/acscatal.8b00624
G. Brown, T. S. Moody, M. Smyth, S. J. C. Taylor, ‘Almac: An Industrial Perspective of Ene Reductase (ERED) Biocatalysis’, in “Biocatalysis: An Industrial Perspective”, eds. G. de Gonzalo and P. Domínguez de María, ch. 8, Royal Society of Chemistry, London, UK, 2018, pp. 229–256 LINK https://doi.org/10.1039/9781782629993-00229
D. Mangan, I. Miskelly and T. S. Moody, Adv. Synth. Catal., 2012, 354, (11–12), 2185 LINK https://doi.org/10.1002/adsc.201101006
R. Stuermer, B. Hauer, M. Hall and K. Faber, Curr. Opin. Chem. Biol., 2007, 11, (2), 203 LINK https://doi.org/10.1016/j.cbpa.2007.02.025
Y. Kawai, M. Hayashi, Y. Inaba, K. Saitou and A. Ohno, Tetrahedron Lett., 1998, 39, (29), 5225 LINK https://doi.org/10.1016/s0040-4039(98)01027-2
H. U. Blaser, B. Pugin, and F. Spindler, ‘Asymmetric Hydrogenation’, in “Topics in Organometallic Chemistry: Organometallics as Catalysts in the Fine Chemical Industry”, eds., M. Beller, H. U. Blaser, Vol. 42, Springer-Verlag, Berlin, Germany, 2012, pp. 65–102 LINK https://doi.org/10.1007/3418_2011_27
D. J. Ager, A. H. M. de Vries and J. G. de Vries, Chem. Soc. Rev., 2012, 41, (8), 3340 LINK https://doi.org/10.1039/c2cs15312b
C. S. G. Seo and R. H. Morris, Organometallics, 2018, 38, (1), 47 LINK https://doi.org/10.1021/acs.organomet.8b00774
D. J. Mansell, H. S. Toogood, J. Waller, J. M. X. Hughes, C. W. Levy, J. M. Gardiner and N. S. Scrutton, ACS Catal., 2013, 3, (3), 370 LINK https://doi.org/10.1021/cs300709m
J. Waller, H. S. Toogood, V. Karuppiah, N. J. W. Rattray, D. J. Mansell, D. Leys, J. M. Gardiner, A. Fryszkowska, S. T. Ahmed, R. Bandichhor, G. P. Reddy and N. S. Scrutton, Org. Biomol. Chem., 2017, 15, (20), 4440 LINK https://doi.org/10.1039/c7ob00163k
M. Breuer, K. Ditrich, T. Habicher, B. Hauer, M. Keßeler, R. Stürmer and T. Zelinski, Angew. Chem., Int. Ed., 2004, 43, (7), 788 LINK https://doi.org/10.1002/anie.200300599
D. Monge, H. Jiang and Y. Alvarez-Casao, Chem. Eur. J., 2015, 21, (12), 4494 LINK https://doi.org/10.1002/chem.201405552
V. A. Soloshonok, C. Cai, V. J. Hruby, L. Van Meervelt and T. Yamazaki, J. Org. Chem., 2000, 65, (20), 6688 LINK https://doi.org/10.1021/jo0008791
T. Inokuma, Y. Hoashi and Y. Takemoto, J. Am. Chem. Soc., 2006, 128, (29), 9413 LINK https://doi.org/10.1021/ja061364f
D. A. Evans, T. C. Britton, R. L. Dorow and J. F. Dellaria, J. Am. Chem. Soc., 1986, 108, (20), 6395 LINK https://doi.org/10.1021/ja00280a050
T. F. Woiwode and T. J. Wandless, J. Org. Chem., 1999, 64, (20), 7670 LINK https://doi.org/10.1021/jo990916s
Download the Supplementary Information for this article (PDF 127KB)
Samantha Staniland graduated from The University of Manchester, UK, in 2011 with an MChem in Chemistry with Industrial Experience, while carrying out her industrial placement at Pfizer, UK, in Medicinal Chemistry. In 2011–2015, Sam did a PhD in the groups of Professor Jonathan Clayden and Professor Nicholas Turner on the biocatalytic asymmetric synthesis of atropisomers. Sam joined Johnson Matthey in 2015 as a research chemist in catalysis.
Tommaso Angelini completed his PhD in Chemical Science in 2010 from University of Perugia, Italy, working on the development of environmentally friendly synthetic protocols. During his postdoctoral studies, he finalised his work designing new continuous flow devices for the use of solid supported catalyst in low E-Factor transformations. Later, he gained experience in developing active pharmaceutical ingredient (API) production process at Procos (Italy). In 2015, he joined Johnson Matthey as Research Chemist, designing new enantioselective synthetic process for the preparation of APIs. He is now a Research Expert at Evotec Verona (Italy), working on the production of preclinical and Phase 1 API candidates.
Ahir Pushpanath obtained his PhD in Birkbeck College (University of London, UK) working on the engineering of enzymes for industrial biofuel production. With a biochemistry background, he specialises in the use of bioinformatics and computational biology in the rational design of new enzyme variants. Ahir joined Johnson Matthey in 2013 as a Senior Biologist and was instrumental in demonstrating the utility of computational techniques for rapid enzyme discovery through genome mining, in silico design and targeted enzyme engineering. He currently leads the enzyme development arm of biocatalysis, continuing to develop faster, more effective methods for ‘predictive biocatalysis’.
Amin Bornadel studied chemical engineering and received a PhD in biotechnology from Lund University in Sweden. For postdoctoral work, Amin went to Germany, where he carried out research within biocatalysis at University of Dresden and Technical University of Hamburg. In 2016, Amin joined Johnson Matthey to work as a biocatalysis researcher. He is currently a senior scientist working in the Biotech team.
Elina Siirola completed her PhD in 2012 from the University of Graz, Austria, where she worked on biocatalytic C=C bond hydrolysis. After a postdoctoral position in enzyme engineering at the Max Planck Institute for Coal Research, Germany, she joined Johnson Matthey in 2013, where she worked on biocatalysis research and development (R&D). Since 2017 she is a Principal Scientist in the Bioreactions group at Novartis Pharma in Basel, Switzerland.
Serena Bisagni completed her MSc in Industrial Biotechnology from the University of Pavia, Italy, in 2010 and then moved to Lund University, Sweden, for her postgraduate studies. In 2014 she obtained her PhD in Biotechnology in which she focused on the identification of new Baeyer-Villiger monooxygenases for fine chemicals synthesis within the Marie Curie Innovative Training Networks (ITN) ‘Biotrains’. In 2015 Serena joined Johnson Matthey. Her main interests are enzyme screening for synthesis of active pharmaceutical ingredients and fine chemicals and identification of novel biocatalysts.
Antonio Zanotti-Gerosa studied in Milano, Italy, completing his PhD in 1994 (organometallic chemistry). His academic experience include secondments to Imperial College, UK (Professor S. V. Ley), Nagoya University, Japan (Professor R. Noyori) and postdoctoral research at the University of Lausanne, Switzerland (Professor C. Floriani). Since 1997 he has been working on industrial applications of homogeneous catalysis. In 2003 he joined Johnson Matthey and, as R&D Director, he is leading the chemocatalysis group in the Cambridge laboratories.
Beatriz Domínguez gained her PhD in Synthetic Organic Chemistry from the University of Vigo, Spain, and then moved to the UK where she worked with Professor Tom Brown at the University of Southampton, UK, and with Professor Guy Lloyd-Jones at the University of Bristol, UK. In 2002 she joined Synetix, soon to become Johnson Matthey Catalysts and Chiral Technologies and has worked at Johnson Matthey’s facilities in Cambridge since. Beatriz has gained broad experience in the application of metal catalysis and biocatalysis, working closely with fine chemicals companies to deliver optimal catalysts for chemical processes.