Chemoenzymatic Resolution of Selected Vicinal Diols Faculty of Natural Resources School of Business and Natural Science Universi

Viðskipta- og Raunvísindasvið Auðlindadeild 2014

Chemoenzymatic resolution of selected vicinal diols

Sean Michael Scully Final Project in Biotechnology

Faculty of Natural Resources School of Business and Natural Science University of Akureyri 2014

Háskólinn á Akureyri Viðskipta- og Raunvísindasvið

Viðskipta- og Raunvísindasvið Auðlindadeild

Námskeið LOK 1123 og LOK 1223 Heiti verkefnis Chemoenzymatic resolution of selected vicinal diols Verktími August 2011 – April 2014 Nemandi Sean Michael Scully Leiðbeinandi Sigþór Pétursson, PhD Upplag Blaðsíðufjöldi 104 Fjöldi viðauka Útgáfu- og notkunarréttur Fylgigögn ISSN-númer

Lokaverkefni til 90 eininga B.Sc.-prófs á líftæknibraut

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I hereby attest that I am the author of this dissertation herein and that it is a product of my own academic research from the autumn of 2012 to the spring of 2014.

Sean Michael Scully

Signature, date and place

I hereby certify that the dissertation successfully fulfills the requirements to complete a Bachelor of Science in Biotechnology at the University of Akureyri.

Signature, date and place

Abstract

The use of chemoenzymatic resolutions for the separation of racemic mixtures is a common route that affords enantiomerically pure compounds. Finding novel and selective protecting group strategies to facilitate the resolution of 1,2-diol systems remains challenging. The use of tin(II) halide-catalyzed reactions with diazo[bis(4- methylphenyl)]methane with racemic vicinal 1,2-diols gives a mixture of the corresponding 1- and 2-benzhydryl ethers with a regiopreference for the primary hydroxyl group in good overall yield. Tin(II) chloride and Tin(II) bromide were evaluated as catalyst for the regioselective introduction of bis(4-methylphenyl)methane ethers from the corresponding diazocompound. Following partial protection, five commercially available lipases were evaluated for their ability to resolve 1-benzhydryl ethers of 1,2-diol systems. Subsequent kinetic resolution with immobilized lipase from Pseudomonas cepacia afforded the corresponding (2R) acetate and (2S) alcohol. De-acylation of the 2R diol affords the corresponding free alcohol.

Keywords: Biocatalysis, lipase, kinetic resolution, 1,2-diols, tin(II) halides, stereochemistry .

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Acknowledgements

This work was carried out at the University of Akureyri. I would like to thank Dr. Sigríður Jónsdóttir for her assistance with obtaining NMR and MS data over the past 4 years. I would also like to kindly think Eva María Ingvadóttir for her helpful comments regarding this manuscript.

I would like to express my deep and eternal gratitude to Dr. Sigþór Pétursson for opening the door for the continuation of my career in the physical sciences in 2005 and helping me restart my career in science in 2008.

I would also like to extend my gratitude to my lovely wife, Hugrún, for her nearly unlimited patience with my work and academic habits over the past seven years. The finalization of this manuscript would not have been possible without her support, dedication and a willingness to overlook my continued absence.

„I do not know what I may appear to the world, but to myself I seem to have been only like a boy playing on the sea-shore, and diverting myself in now and then finding a smoother pebble or a prettier shell than ordinary, whilst the great ocean of truth lay all undiscovered before me.“ Sir Issac Newton

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Útdráttur

Aðskilnaður handhverfa með efnafræðilegum aðferðum og notkun ensíma eru vel þekktar aðferðir við einangrun hreinna handhverfa. Dæmi um slíkar handhverfur eru 1,2-diol, sem innihalda hendið kolefni, og afleiður þeirra. Staðvendin myndun eter afleiða af eingreindum alkohól hóp slíkra díóla, sérstaklega með stórum alkyl hópum, hentar vel fyrir myndun afleiða fyrir ensím hvörfin. Tin(II) halíð hvata hvörf diaryldiazometan efnasambanda við hliðlæg díól og mynda eingöngu einetera þar sem eingreindi eterinn myndast í meira magni. Heildarheimtur einetranna eru góðar. Samanburður var gerður á tin(II) klóríði og tin(II) brómíði sem hvata fyrir myndun bis(4-methylphenyl)methyl etera með notkun tilheyrandi diazoefnsambands. Eftir einangrun eingreinda etersins, voru fimm aðkeyptir lípasar rannsakaðir við aðskilnað handhverfa þessara 1-diarylmethyl etera díólanna. Lípasinn frá Pseudomonas cepacia var síðan notaður við myndun (2R)-asetatsins sem var skilið frá (2S)-alkohólinu. Afasetylerun (2R)-asetatsins gaf síðan (2R)-alkohólið.

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Table of Contents

Abstract ...... iii

Acknowledgements ...... iv

Útdráttur ...... v

List of Figures ...... ix

List of Tables ...... xi

Abbreviations ...... xii

1 Background and Research Objectives ...... 1

1.1 Stereochemistry...... 2

1.1.1 Basic terminology in stereochemistry ...... 4

1.1.2 Nomenclature of stereoisomers ...... 5

1.1.3 Metrics in Stereochemistry ...... 9

1.1.4 Asymmetric strategies in synthesis ...... 11

1.2 Biocatalysis with lipases ...... 13

1.2.1 General Aspects of Biocatalysis ...... 14

1.2.2 Lipases ...... 16

1.2.3 Biocatalytic routes to enantiomerically pure diols ...... 29

1.3 Benzhydryl Derivatives as Protecting Groups ...... 33

1.3.1 Overview of Protecting Group Strategy for Hydroxyl Groups ...... 33

1.3.2 Benzhydryl Ethers as Hydroxyl Protecting Groups ...... 36

2 Experimental ...... 41

2.1 General methods ...... 41

2.2 Preparation of diazo compounds ...... 43

2.2.1 Preparation of diazo[bis(4-methylphenyl)]methane ...... 43

2.2.2 Preparation of diazo[bis(4-methoxyphenyl)]methane ...... 43

2.3 Protection studies towards primary benzhydrylation of vicinal diols ...... 44

2.3.1 Benzhydralation experiments on 1,2-propanediol ...... 44

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rac-[1-O-bis(4-methylphenyl)methyl]–propan-2-ol (SnCl2 catalyzed) ...... 44

rac-[1-O-bis(4-methylphenyl)methyl]–propan-2-ol (SnBr2 catalyzed) ...... 45

2.3.2 Benzhydrylation experiments on 3-phenoxy-1,2-propanediol ...... 47

3-O-phenoxy-1-O-(bis(4-methylphenyl)methyl)-propan-2-ol (SnCl2 catalyzed) ...... 47

3-O-phenoxy-1-O-(bis(4-methylphenyl)methyl)-propan-2-ol (SnBr2 catalyzed) ...... 47

Effort towards 1-O-bis(4-methoxyphenyl)dimethyldiphenyl –propan-2-ol (SnCl2 catalyzed) ...... 48

Effort towards 1-O-bis(4-methoxyphenyl)dimethyldiphenyl –propan-2-ol (SnBr2 catalyzed) ...... 48

2.3.3 Primary benzyhydrylation of 3-chloro-1,2-propanediol ...... 49

1-O-bis(4-methylphenyl)-3-chloro-propan-2-ol (SnCl2 catalyzed) ...... 49

Separation of 1-dimethyldipheny 3-chloro-propane-2-ol ether from 2 ether (pg 40) .. 50

2.4 Lipase-catalyzed resolution of partially protected diols ...... 51

2.4.1 Comparison of lipases for 2(R) acetylation of rac-1[bis(4- methylphenyl)]methoxypropane-2-ol ...... 51

2.4.2 Kinetic Resolution of rac-1[bis(4-methylphenyl)]methoxypropane-2-ol ...... 52

2.4.3 Kinetic Resolution of 1-dimethyldiphenyl ether of rac-3-phenoxy-1[bis(4- methylphenyl)]methoxypropane-2-ol ...... 53

2.4.4 Kinetic Resolution of rac-3-chloro-1[bis(4-methylphenyl)]methoxypropane-2-ol 54

2.5 Deprotection of (2R)-Acetyl Derivatives...... 55

2.5.1 Deprotection of (2R)-1[bis(4-methylphenyl)]methoxy-2-acetoxypropane ...... 55

2.5.2 Deprotection of (2R)-3-phenoxy-1[bis(4-methylphenyl)]methoxy 2- acetoxypropane ...... 55

2.5.3 Deprotection of (2R)-3-chloro-1[bis(4-methylphenyl)]methoxy 2-acetoxypropane ...... 56

3 Results ...... 57

3.1 Preparation of diazocompounds ...... 57

3.2 Protection studies towards primary benzhydrylation of vicinal diols ...... 57

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3.3 Kinetic Resolution of Partially Protected Diols ...... 58

3.3.1 Comparison of lipases for 2(R) acetylation of rac-1[bis(4- methylphenyl)]methoxypropane-2-ol ...... 58

3.4.2 PCL-mediated chemoenzymatic resolution and deprotection of rac-1[bis(4- methylphenyl)]methyl ether derivatives ...... 60

4 Discussion ...... 62

4.1 Preparation of diazo compounds ...... 62

4.2 Protection studies towards primary benzhydrylation of vicinal diols ...... 62

4.3 Kinetic resolution of partially protected diols using lipases ...... 65

4.3.1 Comparison of lipases for 2(R) acetylation of rac-1[bis(4- methylphenyl)]methoxypropane-2-ol ...... 66

4.4.2 Chemoenzymatic resolution and deprotection of rac-1[bis(4- methylphenyl)]methoxy derivatives of 1,2-propanediol...... 67

4.4 Future directions ...... 68

5 Conclusions ...... 70

6 References ...... 71

Appendix A Kinetic Data ...... 79

Appendix B 1H and 13C NMR Spectra ...... 84

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List of Figures

Figure 1 – Relationships between isomers...... 3 Figure 2 – Enantiomers arise as a result of a stereogenic atom bearing four different substitutions while the molecule is non-superposable on its mirror image ...... 4 Figure 3 - The Fischer conventions for relative stereo configuration (adapted from Carey & Sundberg, 2001; pg 81) ...... 6 Figure 4 - Chirality rule for the assignment of absolute configuration. This is the R- configuration if the priority is A → B → C → D (modified from Eliel & Wilen, 1994) ...... 7 Figure 5 - Erythro threo (modified from Smith & March, 2007) ...... 8 Figure 6 - Syn anti system for adjacent stereocenters (modified from Smith & March, 2007) . 8 Figure 7 - Stereoisomers of 2,3-dihydroxybutanoic acid...... 9 Figure 8 - Calculation of enantiomeric excess as a percentage from the mole fraction of two enantiomers when the R enantiomer is the major species (Eliel & Wilen, 1994) ...... 10 Figure 9 - Asymmetric modification strategies (modified from Ager, 2006)...... 11 Figure 10 - Hydrolysis of triacylglycerides by lipases ...... 16 Figure 11 – Types of reactions catalyzed by lipases (Halldorsson, 2003; pg 6) ...... 17 Figure 12 - Double Displacement “ping-pong” Mechanism (From Proteins pg 389 by Creighton) ...... 23 Figure 13 – Mechanism of Lipase-catalyzed transesterification of an alcohol substrate (modified from Anthorsen, 2001); active site residues are depicted in black, acyl donor is pink, alcohol substrate is orange ...... 24 Figure 14 - Stereochemical numbering of glycerides (modified from Bornscheuer & Kazlauskas, 2006) ...... 26 Figure 15 - The relative substrate specificity of commercially available lipases (modified from Faber, 2011) ...... 27 Figure 16 - Kazlauskas's rule for the prediction of the faster reacting enantiomer among secondary alcohols ...... 28 Figure 17 - Yeast-mediated preparation of (S)-1,2-propanediol by oxidation (Kometani et al., 1993) ...... 30 Figure 18 - Resolution of 1,2-propanediol by Baker's yeast-mediated redox reactions (modified from Kometani, et al, 1996) ...... 31 Figure 19 – Application of Kazlauskas’s Rule to 1,2-propanediol derivatives; “small” group = blue, “large” group = red ...... 32

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Figure 20 - Structures of four common diarylmethyl ethers; (A- benzhydryl, B-, C-, D-) ..... 36 Figure 21 - Mechanism of benzhydryl ether formation from corresponding diazo compound via thermolysis ...... 37 Figure 22 - Formation of electrophilic metal-carbene complexes (Carey & Sundberg, 2001a) ...... 39 Figure 23 - Proposed partial mechanism of stannous chloride-mediated reaction of diaryldiazo compounds with vicinal diols (modified from Pétursson, 2001) ...... 40 Figure 24 – Pseudomonas cepacia Lipase (PCL) catalyzed kinetic resolution of 1-bis(4- methylphenyl)methyl-propan-2-ol ...... 59 Figure 25 – Pseudomonas fluorescens Lipase (PFL) catalyzed kinetic resolution of 1-bis(4- methylphenyl)methyl-propan-2-ol ...... 59 Figure 26 – Mucor mehei Lipase (MML) catalyzed kinetic resolution of 1-bis(4- methylphenyl)methyl-propan-2-ol ...... 59 Figure 27 – Candida antarctica Lipase (CAL) catalyzed kinetic resolution of 1-bis(4- methylphenyl)methyl-propan-2-ol ...... 59 Figure 28 - PCL-catalyzed resolution of 3-phenoxy-1[bis(4-methylphenyl)]methoxy 2- acetoxypropane ...... 61 Figure 29 - PCL-catalyzed resolution of rac-3-chloro-1[bis(4- methylphenyl)]methoxypropane-2-ol ...... 61 Figure 30 – PCL-catalyzed reaction of 1-bis(4-methylphenyl)methyl-propan-2-ol with vinyl acetate ...... 79 Figure 31 -PFL-catalyzed reaction of 1-bis(4-methylphenyl)methyl-propan-2-ol with vinyl acetate ...... 80 Figure 32 – MML-catalyzed reaction of 1-bis(4-methylphenyl)methyl-propan-2-ol with vinyl acetate ...... 81 Figure 33 – CCL-catalyzed reaction of 1-bis(4-methylphenyl)methyl-propan-2-ol with vinyl acetate ...... 82 Figure 34 – CAL-catalyzed reaction of 1-bis(4-methylphenyl)methyl-propan-2-ol with vinyl acetate ...... 83

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List of Tables

Table 1 - Comparison of enzyme and whole cell biocatalysis (modified from Faber, 2011) .. 14 Table 2 – Use of lipases for industrial-scale production of compounds (Liese et al., 2006) ... 18 Table 3 - The structures of some commercially available lipases ...... 21 Table 4 - Tin(II) halide benzhydrylation of vicinal diols using bis(4-methyl phenyl)diazomethane ...... 57 Table 5 – Summary of yields from PCL-catalyzed resolutions of rac-1-[bis(4- methylphenyl)]methyl ether derivatives of selected diols ...... 60

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Abbreviations

°C – degrees Celsius 12PD – 1,2-propanediol Ac – Acetate Aq – Aqueous Bz – Benzoyl CAL – Candida antartica lipase CRL – Candida rugosa lipase d - days d.r. = diastereometric ratio DCM-dichloromethane DMAP – dimethylamino pyridine DME- 1,2-dimethoxyethane ee – enantiomeric excess eq- equivalence EtOAc – Ethyl acetate GC – gas chromatography Gem- geminal h - hours Hex - Hexane HPLC – high performance liquid chromatography min – minute(s) mol - mole NMR – nuclear magnetic resonance PCL – Pseudomonas cepacia (Burkholderia cepacia) PPL – Porcine pancreatic lipase R – rectus rac- racemic

Rf – retention factor

Rt – retention time rt – room temperature S – sinister

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1 Background and Research Objectives

The increasing awareness of the need for green chemistry and enantiomerically pure building blocks has driven interest in the use of biotransformations in synthetic organic and process chemistry. The demand for building blocks high in enantiomeric purity for sectors such as the pharmaceutical and pesticide industry has intensified as a result of awareness of the biological differences which can be largely attributed to developing an understanding that an increased incidence of birth defects in the middle of the 20th century were caused by use of the non-therapeutic enantiomer of thalidomide (Solomons & Fryhle, 2007). The minimum optical purity for a chiral building block is 95% although commercial processes must typically produce building blocks of 98% or greater enantiomeric excess (Pollard & Woodley, 2007). As such, a highly-selective process for producing or resolving enantiomers is needed.

Small vicinal diols, such as 1,2-propanediol and closely related compounds, present challenging targets for chemoenzymatic resolution due to their small size and similar reactivity of their adjacent hydroxyl groups. While the use of enzymes such as lipases (Section 1.3) for the resolution of secondary alcohols for the production of fine chemicals has been well establish, the selective introduction of large protecting groups has not received much attention. Thus, the need for novel protecting group strategies coupled with a sufficiently discriminating enzyme is needed for successful kinetic resolution. Facile routes to small, optically enriched vicinal diols such as 1,2- propanediol and its derivatives are a particular challenge. Due to the similarity in reactivity and the close proximity of the hydroxyl groups, small diols are difficult to resolve using traditional chemoenzymatic methods.

The purpose of the work presented in this thesis is twofold: to explore the regio-selective introduction of novel benzhydryl ether derivatives as protecting groups for the primary protection of vicinal diols through regioselective introduction using a tin(II) halide catalyst to kinetically resolve partially protected diols using commercially available immobilized lipases

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1.1 Stereochemistry

Although the phenomenon of optical activity of crystals and organic solutions had been previously investigated by Arago and Biot in the early 19th century, it is generally recognized that discovery of molecular chirality began with Pasteur’s resolution of the sodium ammonium salts of tartaric acid which he reported in 1848 and resulted in the separation of two forms of tartaric acid, each rotating plane polarized light in the opposite direction from the other (Flack, 2009). For this reason, Pasteur is generally credited with the discovery of chirality. Decades later, Jacobus van’t Hoff and Joseph Le Bel proposed that the optical behavior of organic molecules was due to the presence of an asymmetric carbon atom in which the carbon is tetravalent bearing four different ligands thus bringing chirality into the realm of structural organic chemistry (Gasteiger, Gillespire, Marquarding, & Ugi, 1974).

Isomerism results from the different connectivity of atoms within a molecule; constitutional (structural) isomerism is concerned with the molecular connectivity of atoms whereas stereoisomerism describes molecules with the same connectivity but differing in the arrangement of atoms in three-dimensional space (Solomons & Fryhle, 2007). Stereoisomers can be divided into diastereomers and enantiomers. Enantiomers are molecules that are non-superimposable mirror images of one another whereas diastereomers are stereoisomers that are not mirror images of each other (Solomons & Fryhle, 2007). Figure 1 describes the relationships between different types of isomerism.

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Isomers

Connectivity

Differing Identical

Constitutional Isomers Stereoisomers

Rotation about σ bond

Identical Non-Identical

Confomers Configurational Isomers

Mirror plane

Nonsuperposable Multiple stereocenters, Mirror images Not mirror images

Enantiomers Diastereomers

Figure 1 – Relationships between isomers

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In the synthesis of organic molecules, the control of the configuration of stereocenters is the key; failure to do so can result in the formation of mixtures of enantiomers or diastereomers that will, at best, lower the yield of the desired product, or produce a mixture of isomers that cannot be separated by conventional separation methods.

1.1.1 Basic terminology in stereochemistry Among molecules that differ in their arrangement in three-dimensional space, asymmetric molecules can be further divided into enantiomers and diastereomers. Any molecule that has a non-superposable reflection, as demonstrated in Figure 2, is an enantiomer. Diastereomers are asymmetric molecules that contain more than one chiral center and are not mirror images. Any molecule possessing a plane of symmetry, even if stereocenters are present, is achiral (Carey & Sundberg, 2001; pg 79).

Figure 2 – Enantiomers arise as a result of a stereogenic atom bearing four different substitutions while the molecule is non-superposable on its mirror image

Chirality can occur for several reasons: the presence of one or more quadrivalent stereogenic atoms, restricted bond rotation, or the presence of tertiary structure such as a helical shape (Smith & March, 2007). Trivalent molecules possessing a lone pair of electrons may also be chiral however these are difficult to isolate due to rapid pyramidal inversion(Smith & March, 2007). A chiral molecule may have an axis of rotation but not planes of symmetry, an axis of reflection or inversion centers (Wolf,

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2008). Thus, molecules bearing multiple stereogenic centers may lack optical activity due to the presence of planes of symmetry (diastereomers).

Stereogenic (chiral) centers can occur when an atom bearing at least four different groups is present; a tetrahedral molecule with only one stereogenic center will always be optically active (Smith & March, 2007). Stereogenic centers may be sp3 hybridized carbon atoms or other heteroatoms, such as sulfur or phosphorous (Davies & Teng, 2003). The possible number of stereoisomers for a given molecule cannot exceed 2n where n is the number of chiral centers; for a molecule with multiple stereogenic centers to be an enantiomer, it must have the opposite configuration at all stereogenic centers (Carey & Sundberg, 2001a).

Stereoisomers possess a number of unique physical and biochemical properties including the ability to rotate plane polarized light and a differing reactivity in biological systems; for example, enantiomers often have different tastes and smells. In the absence of an external chiral influence, pairs of enantiomers have the same physical and chemical properties, except the rotation of a plane polarized light. However, enantiomeric composition can have an effect on all physical properties of a mixture of enantiomers (Eliel & Wilen, 1994; pg 162). Enantiomers operating under physiological conditions, a chiral environment may differ in their reactivity.

Enantiomers have opposite configurations at all stereogenic centers, while diastereomers have the same configuration at one or more stereogenic centers but opposite configurations at others.

1.1.2 Nomenclature of stereoisomers

Stereochemistry is often described in terms of either relative or absolute configuration. Relative configuration relates the arrangement of molecules to other chiral molecules, such as glyceraldehyde, whereas absolute configuration is determined by the precise location of atoms in space.

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Relative configuration, commonly used in the biological sciences and glycochemistry, is denoted by D and L and report a compounds arrangement of atoms relative to a compound of known configuration. Enantiomers that rotate plane polarized light clockwise or anti-clockwise are denoted dextrorotatory (d, (+)) or levorotary (l, (-)). Absolute configuration describes the arrangement of a stereogenic center as determined by the Cahn, Ingold, and Preolog system described below. There is no direct relationship between D/L, R/S, or +/- assignments (Carey & Sundberg, 2001a). Once the relative configuration of the reference material is known, the absolute configuration is known.

Fischer Conventions for Relative Configuration

The relative configurations of biomolecules such as carbohydrates and amino acids are defined with reference to the Fischer convention. The Fischer convention relates the configuration of the highest numbered chiral carbon on the compound in question to the glyceraldehyde’s chiral carbon as demonstrated in Figure 3.

HO H HO H

HOH2C CHO HOH2C CHO (D)-(+)-glyceraldehyde (L)-(-)-glyceraldehyde

CHO CHO H C OH HO C H

CH2OH CH2OH

CHO CHO H C OH HO C H

CH2OH CH2OH Figure 3 - The Fischer conventions for relative stereo configuration (adapted from Carey & Sundberg, 2001; pg 81)

The enantiomer, where the hydroxyl group on the chiral carbon in the Fischer projection points to the right, is D and the opposite structure is L. The D and L classification of carbohydrates with more than one chiral atom refers only to the

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highest numbered chiral atom. A Fischer projection always has the most oxidized carbon nearest the top. The numbering is from top to bottom. Fischer projection should be viewed such that all horizontal bonds distend from the viewing plane (i.e. towards the viewer) whereas all vertical bonds project behind the plane of the page. Fischer projections can only be manipulated by rotating them 180° and IUPAC rules state that all hydrogen atoms should be displayed (Solomons & Fryhle, 2007). The use of Fisher projections as well as relative configurations is now discouraged due to ambiguities that can arise.

Cahn-Ingold-Prelog Nomenclature for Absolute Configuration

Absolute configuration, which uses the Cahn-Ingold-Prelog (CIP) nomenclature rules (Carey & Sundberg, 2001a) describes chiral molecules in terms of R (Latin: rectus) and S (Latin: sinister). Under the CIP system, the ligand of lowest priority is placed opposite the viewer and the path from the highest priority (highest atomic number) ligand to the third highest priority ligand is traced as summarized in Figure 4; if the resultant rotation is counter-clockwise, the absolute configuration is S whereas if the resultant trace is clockwise, the absolute configuration about the stereocenter is R.

Figure 4 - Chirality rule for the assignment of absolute configuration. This is the R-configuration if the priority is A → B → C → D (modified from Eliel & Wilen, 1994)

Conventions in naming diastereomers

Threo-erythro is sometimes used to define the relative relationship of two adjacent stereocenters bearing a common functional group such as a hydroxyl (Smith & March, 2007); A Fischer projection on which the substituents on two adjacent stereocenters are on the same side are termed erythro isomers whereas when the

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substituents are on opposite sides are termed threo isomers (Carey & Sundberg, 2001a).

X Y Y X X Y Y X X Y Y X Y X X Y

Erythro pair of Threo pair of enantiomers enantiomers Figure 5 - Erythro threo (modified from Smith & March, 2007)

Another system for denoting the configuration of diastereomers with two adjacent stereocenters is the syn/anti system. Figure 6.

Y Y Y Y

X X X X

syn pair of anti pair of enantiomers enantiomers Figure 6 - Syn anti system for adjacent stereocenters (modified from Smith & March, 2007)

The presence of multiple stereocenters within a molecule can sometimes result in the formation of planes of symmetry and rotation about this plane results in the identical configuration. Compounds bearing multiple stereogenic centers, which contain a plain of symmetry are superimposable with their mirror image, that is are the same as their mirror image, are known as meso compounds. Examples include R,S-butane-2,3-diol and R,S-tartaric acid.

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Enantiomers

CO2H HO2C HO OH OH HO

(2R,3S)-2,3-dihydroxybutanoic acid s

(2S,3R)-2,3-dihydroxybutanoic acid t

e r

r o

r r

o i

m m HO2C CO H e 2 r

s OH HO HO OH

(2S,3S)-2,3-dihydroxybutanoic acid (2R,3R)-2,3-dihydroxybutanoic acid

Enantiomers Figure 7 - Stereoisomers of 2,3-dihydroxybutanoic acid

1.1.3 Metrics in Stereochemistry

The enantiomeric composition can be measured. Optical activity has been the common metric for measuring enantiomeric purity through the use of optical rotary dispersion (ORD) and circular dichroism (Eliel & Wilen, 1994). These methods rely on measurements of optical activity although more modern methods commonly used rely upon chiral chromatography and the use of chiral derivative spectroscopy, including NMR (Beesley & Scott, 1998).

The optical activity refers to a chiral compounds ability to rotate plane-polarized light; optically pure enantiomers rotate light to the same magnitude but opposite. If the enantiomer rotates the light to the right, it is dextrorotatory (Latin: dexter), “d” or “(+)”. Optical isomers that rotate light to the left are levorotatory (Latin: laevus), “l” or “(-)”. Optical rotation (α) is typically measured at 589 nm corresponding to the emission line of a sodium lamp. The specific rotation, defined below, can be calculated from the density of a solution, the observed rotation, and the pathlength of light in decimeters (Solomons & Fryhle, 2007).

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The optical purity can be calculated from the measured optical rotation of a solution divided by the rotation of the pure enantiomer as shown below.

Another common metric to quantify the enantiomeric purity of a mixture is enantiomeric excess (ee) as described in Figure 8. Enantiomeric excess is defined such that it corresponds to optical purity (Eliel & Wilen, 1994). For mixtures of diastereomers, the definition can be modified and presented as diastereomeric excess. The contribution of each chiral component to the overall optical rotation of the mixture is proportional to the mole fraction of each component.

Figure 8 - Calculation of enantiomeric excess as a percentage from the mole fraction of two enantiomers when the R enantiomer is the major species (Eliel & Wilen, 1994)

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1.1.4 Asymmetric strategies in synthesis

The need for chiral bioactive molecules has driven interest in the large-scale synthesis of optically active compounds (Ager, 2006). Techniques fall into two broad categories as summarized in Figure 9: chiral resolution and asymmetric synthesis. Asymmetric synthesis focuses on starting with chiral materials or enantioselectively creating new chiral centers. Chiral resolutions deal with separating enantiomers or diastereomers by taking advantage of specific chemical or physical properties or by using resolving agents, such as enzymes (Section 1.2), to achieve separation.

Asymmetric Manipulations

Resolution Methods Synthetic Methods Separation of stereoisomers Selective modification or creation of stereocenters

Crystallization Physical Separation Enzymatic Methods Chiral Pool Chiral Catalysts Chiral Auxillaries

Organo catalysts Biocatalysts

Figure 9 - Asymmetric modification strategies (modified from Ager, 2006)

Reactions involving stereoisomers can be regarded as either stereospecific or stereoselective; the latter involve stereoisometric starting materials producing stereoisometric products whereas the former involve reactions in which one stereoisomer is formed preferentially (Eliel & Wilen, 1994; pg 97). Asymmetric (or stereoselective) synthesis involves the de novo synthesis of a chiral molecule from an achiral precursor (Eliel & Wilen, 1994); these techniques can be broadly classified into synthetic methods that selectively create or manipulate a chiral center or

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approaches designed to resolve (separate) stereoisomers. Amongst synthetic methods, there are three general strategies: chiral pool synthesis, the use of chiral auxiliaries to transfer chirality, or asymmetric catalysis (Smith & March, 2007). Enantioselective organocatalysis reviewed in Dalko (2007) and the references therein.

Asymmetric induction involves the creation of a new stereogenic center in a molecule already possessing a stereocenter leading to the formation of diastereomers in equal quantities except when steric or electronic factors favor the formation of one diastereomer (Smith & March, 2007). Alternately, chiral pool synthesis involves the successive stereoselective manipulation of inexpensive, naturally occurring chiral building blocks such as sugars or amino acids (Faber, 2011).

Chiral resolution methods, can be divided into crystallization methods, physical separations, and catalytic methods. Chiral pool synthesis draws upon starting materials that are available as chiral compounds; these building blocks are typically natural products such as monosaccharides, amino acids, terpenes, and steroids (Smith & March, 2007). The use of chiral auxiliaries, which includes the use of chiral templates, is reviewed in the reference herein (Schaad, 2006). Optical resolution by crystallization is reviewed in (Nohira & Sakai, 2001). Chiral separations on both analytical and preparative scales rely on either chiral derivatization, or the direct use of a chiral selector in the form of either a chiral mobile phase or a chiral stationary phase (Gubitz & Schmid, 2004a). Cyclodextrins (cyclic oligosaccharides of six to eight residues) are commonly used chiral selectors in chiral chromatography (Gubitz & Schmid, 2004a). Chromatographic chiral separations are reviewed in (Beesley & Scott, 1998; Gubitz & Schmid, 2004b; Subramanian, 2001).

Each methodology is not universally appropriate and often has significant drawbacks. The use of enantioselective catalysis is generally more efficient than the use of chiral auxiliaries and chiral pool synthesis which are both more efficient than chiral resolution (Carey & Sundberg, 2001b). The theoretical yield of a chiral resolution from a racemic mixture is only 50% whereas the use of chiral auxiliaries often requires stoichiometric quantities of a chiral reagent. The chiral pool is limited based upon what nature provides.

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1.2 Biocatalysis with lipases

Biocatalysis involves the use of whole cells or cell components such as enzymes to perform the chemical transformation of organic molecules (Faber, 2011; Glazer & Nikaido, 2007). Recent interest in green chemistry and white (industrial) biotechnology have led to an increase in the use of biocatalysts due to their substrate specificity, chemo-, regio-, and stereoselectivity, reusability, and their production from renewable, biological sources. Biocatalysts have already been used to replace or supplement the large scale synthesis of commercial compounds, often reducing the number of steps required such as in lactic acid production (Lancaster, 2002) and the microbial or chemoenzymatic transformation of steroids (Glazer & Nikaido, 2007).

Biocatalysts have numerous applications ranging from food production, pulp and paper, pharmaceuticals and synthetic organic chemistry. Within synthetic organic chemistry, enzyme systems have been used for the introduction and removal of protecting groups (review: Kadereit, et al., 2002), asymmetric synthesis, kinetic resolutions, and other applications. General reviews of biocatalysis in organic synthesis can be found at the citations herein (Drauz & Waldmann, 2002; Faber, 2011; Liese, Seelbach, & Wandrey, 2006; Nair, Tang, Eriksen, & Zhao, 2010; Whitesides & Wong, 1985; Wong & Whitesides, 1994).

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1.2.1 General Aspects of Biocatalysis

Types of biocatalysts

The use of whole-cells (resting or growing), cell extracts, crude or purified enzymes all constitute biocatalysts any of which can be potentially immobilized or used in solution.

Table 1 - Comparison of enzyme and whole cell biocatalysis (modified from Faber, 2011)

The enzymes responsible for individual reactions of interest are classified according to the International Union of Biochemistry and Molecular Biology enzyme classification (EC) system with the organism which produces a specific enzyme typically noted. Enzymes are organized into six classes: Oxidoreductases (EC 1), Transferases (EC 2), Hydrolases (EC 3), Lyases (EC 4), Isomerases (EC 5), and Ligases (EC 6). Each classification is further divided into subclasses and sub- subclasses based upon the specific motifs used, the intended substrate, or required

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cofactors and ultimately the specific reaction catalyzed. The current classification system contains over 3000 unique enzymes. (Buxbaum, 2007)

Advantages and pitfalls of biocatalysis

Biocatalysts offer a number of advantages over traditional chemical catalysts. Enzymes often operate under mild reaction conditions, are biodegradable, can be immobilized for repeated use, offer high catalytic efficiency, and are often commercially available (Rozzell, 1999). Many enzyme systems can function in non- aqueous environments so long denaturation and dehydration do not occur (Faber, 2011).

The most attractive feature of many biocatalysts is their substrate selectivity. Many enzymes exhibit highly specific substrate, functional group, stereo- and regioselectivity; this allows the extremely specific manipulations of complex molecules (Rozzell, 1999). Alternately, other enzymes exhibit a high degree of catalytic promiscuity and will accept non-native substrates with hydrolases and other digestive enzymes being particularly accommodating towards artificial substrates (Bornscheuer & Kazlauskas, 2006a; Faber, 2011).

While many enzymes require cofactors such as NAD(P)H and FADH2, systems, such as coupled cofactor regeneration schemes or whole-cell systems, have been developed to regenerate in order to mitigate cost (Rozzell, 1999). Despite popular misconceptions, enzymes frequently display high stability at elevated temperatures and in organic media (Bornscheuer & Kazlauskas, 2006); immobilized enzyme catalysts have shown half-lives of weeks or months under industrial process conditions (Rozzell, 1999; pg 2257).

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1.2.2 Lipases Hydrolases (EC 3), such as lipases and esterases, are ubiquitous in nature being produced by bacteria, archaea and eukaryotes alike. Lipases (EC 3.1.1.3), distinguishable from esterases by their natural preference for water-insoluble esters such as long chain fatty acyl groups, are a class of hydrolases which catalyze the hydrolysis of esters with a preference for triacylglycerides. Lipases have received a great deal of attention due to the applications in synthesis and biotechnology, the ease with which they can be isolated, high enantio- and regioselectivity, and because cofactors are not required for hydrolysis. Under physiological conditions, lipases catalyze the hydrolysis of triacylglycerides, as shown in Figure 10 as well as other esters. (Bornscheuer & Kazlauskas, 2006)

O R1 O O Lipase OH O R O 2 R2 O H O O 2 + HO R1 pH 7 O O R3 R3 O O Figure 10 - Hydrolysis of triacylglycerides by lipases

Enantioselective lipase activity has also been developed among catalytic antibodies (Janda, Benkovic, & Lerner, 1989). Lipases have been demonstrated to exhibit varying degrees of substrate and catalytic promiscuity. In addition to the hydrolysis of esters, lipases are typically credited with catalyzing the hydrolysis and acylation of esters as well as transesterifications as summarized in Figure 10.

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1) Hydrolysis of Esters

O O + H2O + R1OH R OR1 R OH 2) Direct Esterification

O O + R1OH + H2O R OH R OR1 3) Transesterification A) Alcoholysis

O O + R2OH + R1OH R OR1 R OR2 B) Acidolysis

O O O O + + R OR1 R2 OH R OH R2 OR1 C) Interestification

O O O O + + R OR R OR 1 2 3 R OR3 R2 OR1 Figure 11 – Types of reactions catalyzed by lipases (Halldorsson, 2003; pg 6)

Lipases have been heavily employed as industrial catalysts such as in the production of chiral building blocks and pharmaceuticals as summarized in Table 2.

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Table 2 – Use of lipases for industrial-scale production of compounds (Liese et al., 2006)

Product Use Lipase source Reaction Type Company (R)-phenylethylamine Intermediate for B. plantarii Carboxylic ester BASF pharmaceuticals and hydrolysis pesticides (3R,4S)-cis-azetidinone acetate Paclitaxel (taxol) P. cepacia Carboxylic ester Bristol-Meyers hydrolysis Squibb F

OH Anti-cholesterol P. cepacia Acetylation Bristol-Meyers drug Squibb O O

F N N N N O Intermediate M. miehei Carboxylic ester Celltech Group t n Bu Bu hydrolysis HN Ph O (R)-oxiranyl-methanol (S)-beta blocker Porcine Carboxylic ester DSM intermediate hydrolysis (-)-endo-hydroxy-2-oxabicyclo[3.3.0]oct-7- Pharmaceutical P. cepacia Carboxylic ester Celltech Group en-3-one intermediate hydrolysis (S)-ibuprofen Anti-inflammatory drug C. cylindracea Carboxylic ester Pfizer hydrolysis Acetic acid 4-(2,4-difluoro-phenyl)-2- Anti-fungal C. antarctica Carboxylic ester Schering Plough hydroxymethyl-pent-4-enyl ester hydrolysis Lotrafiban Fibrinogen receptor C. antarctica Carboxylic ester GlaxoSmithKline antagonist hydrolysis Pharmaceuticals (+)-(1S,2S)-2-methoxycyclohexanol β-lactam building block C. antarctica Transesterification GlaxoSmithKline Pharmaceuticals

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Additionally, Lipase B from Candida antartica has been demonstrated to catalyze other reactions including Michael addition (Faber, 2011), epoxidations, and aldol condensations (Bornscheuer & Kazlauskas, 2006b). These reactions, however, do not require a serine nucleophile and proceed better in its absence (Loo & Van Hollfelder, 2010).

Beyond their natural substrates, a broad range of substrates that can be hydrolyzed by lipases allows for their use in many sectors. Lipases are currently used in the production of cheeses and in washing detergents to remove fat stains (Bornscheuer & Kazlauskas, 2006). Lipases have also been used extensively in synthetic work including the kinetic resolution of enantiomers (Section 1.2.3), the introduction and removal of protecting groups from hydroxylated compounds such as sugars (Kadereit et al., 2002; Wong & Whitesides, 1994) and the production of biodiesel (Robles- Medina, González-Moreno, Esteban-Cerdán, & Molina-Grima, 2009). Lipases have also been used in the preparation of structured and modified lipids (Halldorsson, 2003; Haraldsson & Hjaltason, 2001).

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General Structural Features of lipases

Lipases are single unit proteins that range in size from 30 to 65 kDa and are commonly classified according to sequence alignment falling into three families: mammalian, bacterial, and fungal lipases (Bornscheuer & Kazlauskas, 2006a). A number of structural features play a key role in the catalytic activity of lipases, namely the active site, hydrophobic lid, and the α/β-hydrolase fold (Bornscheuer & Kazlauskas, 2006b).

Many lipases have a “lid” or “flap” composed of helical segment that blocks access to the active site. A lipid-induced conformational change occurs (interfacial activation) which also places the oxyanion-stabilizing residues in a catalytic position (Bornscheuer & Kazlauskas, 2006). For this reason, the use of lipases for acylglyceride hydrolysis is typically performed in a biphasic system including an aprotic nonpolar solvent and an aqueous phase (Faber, 2011) although the nonpolar nature of solvents used in lipase-catalyzed acylations are typically sufficient to induce interfacial activation. It should, however, be noted that CAL-B, CVL and Pseudomonas aeruginosa do not show interfacial activation and other lipases only show interfacial activation with some substrates (Bornscheuer & Kazlauskas, 2006).

The structures of commercially available lipases commonly used in synthetic and industrial applications are shown in Table 3.

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Table 3 - The structures of some commercially available lipases

Candida antarctica lipase Candida antarctica lipase Candida rugosa lipase (CAL-A; Novozyme) (CAL-B; Novozyme) (CRL) PDB 3GUU (Brandt et al., 2009) PDB 3ICV(Qian, et al., 2010) PDB 1TRH (Grochulski,et al., 1994)

Rhizomucor miehei lipase Pseudomonas fluorescens lipase Porcine pancreatic lipase (MML) (PCL, Amano-P) (PPL) PDB 3TGL (Brzozowski, et al., 1992) 1OIL (Kim, et al., 1997) PDB 1ETH (Hermoso et al., 1996)

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Lipase active site and mechanism of action

The architecture of the substrate binding pocket and active site of lipases (and esterases) appears to be highly conserved (Cygler et al., 1994). The substrate pockets of lipases and esterases are a tunnel of variable depth through which the substrate enters and gains access to the active site. Within the tunnel, there are regions in which the various moieties of the substrate interact, namely the alcohol and acidic portion of the ester. One of these pockets is of variable size and hydrophobic while the other is termed the stereoselective pocket. The specific interaction of the groups of the substrate with regards to the orientation of these pockets is the basis for the enantioselectivity as discussed in (Bornscheuer & Kazlauskas, 2006a; Cygler et al., 1994)

The active site of most lipases contains a serine catalytic triad consisting of serine, which acts as a nucleophile, histidine, and aspartate; this configuration is highly similar to other serine hydrolases including serine proteases (Bornscheuer & Kazlauskas, 2006a) although variations in which the aspartate residue has been replaced by glutamate have been reported (Schrag, Li, Wu, & Cygler, 1991). The oxyanion hole, which is responsible for stabilizing the highly reactive deprotonated serine via hydrogen bonding, typically consists of glycine and alanine residues (Bornscheuer & Kazlauskas, 2006a). While lipases differ from esterases in their amino acid sequence, the serine-histidine-aspartate catalytic triad and residues responsible for the stabilization of the oxyanion hole are highly conserved. Lipases, like many other hydrolases, use an α/β-hydrolase fold motif which positions the catalytic residues within the active site (Bornscheuer & Kazlauskas, 2006b). The position of the serine residue is implanted in a turn between a β-sheet sandwiched by an α-helix and a β-strand with the histidine residue and the oxyanion hole on either side (Cygler et al., 1994).

Mechanism of Action

Lipases catalyze the hydrolysis of esters in which water acts as a nucleophile; other nucleophiles, such as vinyl acetate, can be used instead of water in which case a transesterification occurs (Loo & Van Hollfelder, 2010). Both reactions involve a

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double displacement (“bi-bi” or “ping-pong”) mechanism involving two successive reactions. In the case of the esterification reaction, the formation of the acyl donor- serine intermediate is followed by the nucleophilic attack of the substrate as summarized in Figure 12.

A P B E EA E' E'B E+Q Figure 12 - Double Displacement “ping-pong” Mechanism (From Proteins pg 389 by Creighton)

The mechanism by which lipases catalyze the acylation of alcoholic substrates is shown in Figure 13. In the hydrolysis reaction, the substrate (e.g. vinyl acetate) is replaced by water.

23 Háskólinn á Akureyri Viðskipta- og Raunvísindasvið 1) Formation of activated nucleophile 2) Stabilization of nucleophile 3) Formation of lipase-acyl donor intermediate

His

Ser Asp O H N N H O O O H N N H O O O H N N H O H N O R2 O R1 H N O Acyl Donor (Vinyl Acetate) "Oxyanion hole" 4) Tetrahedral Intermediate I 5) Formation of Serine-ester 6) Attack of substrate

O O O -R1OH H N N H O H N N H O H N N O O R1 O O R1 O O R O H N R2 O 2 H OR' R2 O Tetrahedral H N Intermediate I (Td1) 7) Tetrahedral Intermediate II Release of ester and active site regeneration Regenerated catalytic triad

O O O R' O H N N H O H N N H O O R2 H N N H O O R' O O R' O (R)-Ester O R2 O R2 O H N H N

Figure 13 – Mechanism of Lipase-catalyzed transesterification of an alcohol substrate (modified from Anthorsen, 2001); active site residues are depicted in black, acyl donor is pink, alcohol substrate is orange

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Within the active site, the nucleophilic serine is activated by the removal of a proton from its hydroxyl group by the imidazole ring of histidine. The resultant serine oxyanion is stabilized by the oxyanion hole; the serine oxyanion is a powerful nucleophile which can then attack the substrate, in this case an acyl donor (vinyl

acetate) resulting in the formation of a tetrahedral intermediate (Td1). An internal rearrangement leads to the formation of a serine-ester and a release of acetaldehyde. The alcohol substrate then attacks the serine-ester after losing a proton to the imidazole ring of histidine. The lone pair of electrons on the serine’s oxygen atom then abstract a proton from the histidine residue resulting in the release of the acylated alcohol substrate followed by the regeneration of the active site.

The generation of acetaldehyde from acylation using vinyl esters can lead to the irreversible inactivation of lipases; a study by Weber, et al. (1995) demonstrated that the activity of lipases is stable in the presence of acetaldehyde with the exception of Candida rugosa and Geotrichum candidum.

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Substrate specificity

The native substrate of lipases is glycerol and its acyl derivatives; each of glycerol’s hydroxyl groups is assigned a stereonumber (sn) as shown in Figure 14.

sn-1 position

CH2OH sn-2 position HO * H CH OH 2 sn-3 position

Figure 14 - Stereochemical numbering of glycerides (modified from Bornscheuer & Kazlauskas, 2006)

Some lipases are regioselective for the sn-1 and sn-3 positions. Individual lipase sn- selectivity varies with interfacial tension within the active site and can thus vary with variation of the groups attached to the other sn-positions (Bornscheuer & Kazlauskas, 2006a).

Lipases and esterases natively catalyze the reversible hydrolysis of esters and the acylation of alcohols. Generally, lipases and esterases are somewhat chemoselective in that they will react preferentially with hydroxyl groups of higher acidity with primary hydroxyl groups being the most reactive (1˚>2˚>3˚). Lipases have been shown to react regiospecifically with primary hydroxyl groups of some substrates such as 1,2-alkyldiols (Parmar et al., 1993).

Lipases can also catalyze the enantioselective acylation of other heteroatoms including nitrogen and sulfur. The N-acylation of amines is generally slower than O- acylations; alcohols can be selectively acylated in the presence of more reactive amides by using less reactive acylating agents such as esters or carbonate (Bornscheuer & Kazlauskas, 2006). The hydrolysis of amides, however, does not typically proceed or proceeds slowly (Bornscheuer & Kazlauskas, 2006). The use of lipases for the enantioselective hydrolysis or alcoholysis of primary and secondary thiols has also been reported although the acylation of thiols does not proceed using lipases; carboxylic acids and amine, may however, be reactive in some cases (Bornscheuer & Kazlauskas, 2006).

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Steric factors are the largest determinant of lipase selectivity although electronic factors such as the presence of electron withdrawing groups on the substrate lower selectivity (Bornscheuer & Kazlauskas, 2006). Commercially available enzymes differ in the size of substrate that they will react with as shown in Figure 15. Generally, lipases from digestive organs (such as bovine and porcine lipase) and fungi can accommodate bulkier substrates whereas bacterial lipases have narrower active sites.

Figure 15 - The relative substrate specificity of commercially available lipases (modified from Faber, 2011)

Empirical rules for predicting the stereopreferences at secondary hydroxyl groups are difficult as enantioselectivity has been demonstrated to vary with temperature, solvent, water activity, immobilization method and the purity of the enzyme preparations. These rules have been developed through extensive experimentation with substrates and were verified by X-ray studies (Cygler et al., 1994). Enantioselective lipases typically react with the R-enantiomer according to Kazlauskas’s Rule although S-selective enzymes have been reported in the literature (Bornscheuer & Kazlauskas, 2006a; Kazlauskas, et al., 1991).

Kazlauskas’s Rule, an extension of Prelog’s rule, predicts that when a hydroxyl group (or ester) is present on a chiral carbon, the enantioselectivity of the reaction will be governed by the size of the groups immediately adjacent to the chiral carbon atom as shown in Figure 16 (Kazlauskas et al., 1991). In lipase-catalyzed acylations, the faster reacting substrate will contain a medium-sized substituent (M) such as a methyl group

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in a specific orientation relative to the reacting hydroxyl group and a larger substituent (such as a phenyl group).

(R) H OH M L

Figure 16 - Kazlauskas's rule for the prediction of the faster reacting enantiomer among secondary alcohols

For the majority of lipases and esterases, this rule governs that the (R)-enantiomer will be the faster reacting enantiomer. The enantioselectivity can also be reversed by interchanging the medium substituent for a larger substituent (Bornscheuer & Kazlauskas, 2006). It has also been determined that the slower reacting enantiomer disrupts the hydrogen bonding of the histidine’s imidazole ring to the oxygen atom of the ester (Silverman, 2002). In addition to steric factors, it has been suggested by a number of studies (Bornscheuer & Kazlauskas, 2006a) that electronic effects of the substrate play a role in the substrate selectivity of the catalyst.

Beyond the normal acyl glyceride substrates, lipases are capable of catalyzing the acylation (and corresponding hydrolysis) of many different alcohol-bearing substrates. Lipases have largely been used for the chemoenzymatic resolution of secondary alcohols although secondary alcohols with only a small difference in their size are difficult to resolve (Baumann, Hauer, & Bornscheuer, 2000), including 1,2- propanediol. The use of lipases to resolve 1,2-propanediol and closely related substrates is examined in the following section.

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1.2.3 Biocatalytic routes to enantiomerically pure diols

Resolution of 1,2-propanediol and its derivatives

While several methods have been successfully applied to the resolution of racemic mixtures of 1,2-propanediol, most have focused on biotransformations rather than chemoenzymatic resolutions. The resolution of derivatives of 1,2-propanediol, however, has been more broadly applied and includes chemoenzymatic routes facilitated by lipases.

Generally, three synthetic routes using lipases are applicable to the resolution of 1,2- diols: selective esterification, transesterification and ester hydrolysis. The ability to resolve (R)- and (S)-isomers is related to the steric bulk of the molecule where bulkier groups attached to the primary hydroxyl group tend to produce better resolutions. (Goergens & Schneider, 1991)

1,2-propanediol

Relatively few studies using lipases have been reported on the resolution of racemic 1,2-propanediol. Goergens and Schnieider (1991) reported the resolution of 1,2- propanediol by the silylation of the primary hydroxyl group using a tert- butyldimethylsilyl (O-TBDMS) ether on the primary hydroxyl group followed by Pseudomonas fluorescens (SAM-1) lipase-catalyzed hydrolysis of the TBDM group in 50% conversion of the desired enantiomer with ee’s greater than 95%. While this methodology produces good results and protected (S)-esters may be of direct synthetic value, silyl ester groups are expensive, corrosive, and typically require global deprotection to remove. For these reasons, such a route is not viable to the large-scale production of chiral 1,2-diols.

Another approach used by Petursson (2009) involved the partial protection of the primary hydroxyl group via a reaction with bis-(4-methoxyphenyl)diazomethane to yield the 1-(4,4’-dimethoxybenzhydryl) ether as the primary product. This was followed by the subsequent removal of small quantities of the unwanted 2-ether with

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tritylation; the mixture was then resolved using a lipase from Pseudomonas cepacia (PCL) using vinyl acetate as the acyl donor. (R)-and (S)-1,2-propanediol were isolated in low overall yield with high enantiomeric purity (Petursson, 2009). 1-fluorenyl ether of 1,2-propanediol was also reportedly resolved using PCL (Petursson & Jonsdottir, 2012).

A more facile approach reported by (Sigthor Petursson & Jonsdottir, 2011) including the use of triphenylmethyl (trityl, Trt), with its highly regioselective preference for primary hydroxyl groups, has also been used to resolve racemic 1,2-propanediol using a more promiscuous lipase (Mucor meihei, MML). MML has a much more accepting active site than other commercially available lipases, such as PCL and PFL, and can thus presumably accomadate the bulky trityl ether.

Various biocatalytic routes have also been employed for the resolution of 1,2- propanediol. Lee and Whitesides (1986) reported the enantioselective reduction of 1- hydroxy-2-propanone to (R)-1,2-propanediol by immobilizing commercially available glycerol dehydrogenase (GDH) from Enterobacter aerogens using FDH and NAD+ over 19 days under argon in 50% yield. Studies of GDH from both E. aerogens and Cellomonas sp. (also commercially available) have shown that the GDHs have high selectivity for glycerol and 1,2-propanediol as substrates but suffer from both noncompetitive and mixed product inhibition which necessitates the continuous removal of product when performed on a large scale (Lee & Whitesides, 1986)

Yeast-mediated bioreductions and oxidations have presented simple and potentially industrially feasible routes to both the (R)- and (S)-isomers of 1,2-propanediol. Kometani, et al (1993) reported an aerobic baker’s yeast-mediated preparation of (S)- 1,2-propanediol from a racemic mixture in 51% yield with an ee of 82.3% while a second resolution yielded the (S)-isomer in 71% yield with 98.0% ee.

OH Baker's yeast O OH OH OH + OH O2 (S)-1,2-propanediol 51%, 82.4% ee Figure 17 - Yeast-mediated preparation of (S)-1,2-propanediol by oxidation (Kometani et al., 1993)

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When performed on a large scale (10L), the production of (S)-1,2-propanediol was produced in approximately 25% chemical yield with an enantiomeric excess of 79% which could be further enriched to greater than 96% ee. The (R)-isomer was oxidized to hydroxyacetone (acetol) which was isolated as an aqueous distillate and then reduced to (R)-1,2-propanediol by yeast-mediated reduction in about 60% yield and over 98% enantiomeric excess. (Kometani, et al., 1995)

OH OH Bakers' Yeast-mediated R OH Enantioselective oxidation OH (S)-diol R O2

O OH OH OH R R Bakers' Yeast-mediated 1-hydroxy alkanone (R)-diol Enantioselective reduction

Figure 18 - Resolution of 1,2-propanediol by Baker's yeast-mediated redox reactions (modified from Kometani, et al, 1996)

A fed-batch yeast mediated bioreduction of acetol to 1,2-propanediol using yeast under aerobic conditions with ethanol as an energy source to produce (S)-1,2- propanediol in 62% yield with an ee of 96% (Kometani, Toide, Kaikaiji, & Goto, 2001).

1,2-Propanediol Derivatives

The chemoenzymatic resolution of 1,2-propanediol derivatives some studies in the literature using various protecting group strategies and lipases for either hydrolysis or acylation. Overall, the use of larger groups such as halides on the small position (Figure 19) seems to decrease the enantioselectivity of both hydrolysis and esterification reactions in a study using CAL-B whereas the nature of the “large” position, corresponding to group on the primary hydroxyl group, is more complex (Anthonsen & Hoff, 1998). When a phenyl group is used as the “large” group, the

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number of methylene spaces alters the resultant enantiomeric ratio (Anthonsen & Hoff, 1998).

OR2 R1

Figure 19 – Application of Kazlauskas’s Rule to 1,2-propanediol derivatives; “small” group = blue, “large” group = red

Additionally, the enantiomeric ratios of lipase-catalyzed hydrolysis of acylated diols tend to show higher E values than lipase-catalyzed acylation (Anthonsen & Hoff, 1998).

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1.3 Benzhydryl Derivatives as Protecting Groups

1.3.1 Overview of Protecting Group Strategy for Hydroxyl Groups

Protecting (or “blocking”) groups are required to prevent the unwanted reaction of specific functional groups within a molecule; this is of particular importance during multi-step syntheses. A common criticism of many protecting group strategies is that multistep synthesis often suffers a decrease in yield with each subsequent reaction and decrease the atom economy for a synthesis. For a protecting group strategy to be successful, several conditions must be met:

protecting groups must be introduced in a chemo- and regioselective manner they must be introduced and removed in good yield they must be stable under conditions of subsequent manipulations they must possess minimum additional functionality conditions for removal (deprotection) must be selective for specific protecting groups

Common problems with protecting group strategies often involve a lack of selectivity and stability after introduction. The presence of multiple functional groups of similar reactivity presents a constant challenge and is particularly problematic for molecules containing hydroxyl groups and amines (such as diols and carbohydrates). Many protecting groups are sensitive to the presence of acids (such as cyclic ethers) or bases (such as esters). Thus, there is interest in developing protecting group strategies that involve mild reaction conditions for both the protection and subsequent deprotection step.

The most common protecting group strategies for hydroxyl groups, including diols, typically employ a combination of esters and ethers. Hydroxyl groups have a nucleophilic character and will react with electrophiles and oxidizing agents. Generally, primary hydroxyl groups are reactive and thus more easily protected than secondary and tertiary hydroxyl groups. The differences in reactivity of these hydroxyl groups can allow for selective protections under some circumstances. The

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presence of other heteroatoms, which are also nucleophilic in nature, are often a complicating factor in hydroxyl group protection.

Esterification is commonly used for the protection of primary and secondary hydroxyl groups. Esters are easily prepared from an acyl chloride, anhydride, acid or by lipase- catalyzed acylations Selective preparations are possible although the results are highly dependent upon the substrate. The simplest acyl protecting group is the acetyl moiety. Other commonly used ester protecting groups for alcohols, such as benzoyl groups, are base labile and readily migrate to other hydroxyl groups. During regioselective acylations, such as the lipase-catalyzed trans-acylations discussed in Section 1.2, primary hydroxyl groups must be protected to avoid inadvertent reactions necessitating the need for non-ester protecting groups.

Acyl group migrations have been widely reported among substrates bearing one or more free hydroxyl groups including partial glycerides and carbohydrates (Furutani, et al., 1996; Li, et al., 2010; Turon, et al., 2003). Studies on the acyl migration of partial glycerides bearing acyl groups at the sn-1 and sn-3 positions in the presence of lipases has been shown to be water activity and temperature dependent with low water activities favoring migration (Li, et al., 2010) and the decreasing polarity of a solvent system increasing the rate of acyl migration (Li, et al., 2010). For this reason, the use of acyl groups for the protection of primary hydroxyl groups during kinetic resolutions should be avoided.

Ethers are also a commonly used alternative for the protection of hydroxyl groups. They are more acid and base-stable than esters and are less prone to migration. Methyl ethers are introduced to hydroxyl groups by the Williamson ether synthesis using methyl halides and a strong base (Robertson, 2003) or the use of other methyl transfer reagents such as diazomethane. (Wuts & Greene, 2007). Methyl ethers are highly stable in acidic and basic conditions making their removal problematic. Methoxymethyl (MOM) protecting groups can be introduced via the corresponding chloride or ammonium halide salt and removed by either a Lewis acid or base catalyst. Reagents commonly used to introduce MOM groups have the distinct disadvantage of being highly carcinogenic.

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A number of bulky ether protecting groups can be introduced into molecules with high regioselectivity. Ethers such as triphenylmethyl (trityl), and silyl ethers such as trimethylsilyl (TMS) are commonly used to protect hydroxyl groups; these groups can be introduced with good selectivity onto the least sterically hindered alcohol in good yield (Wuts & Greene, 2007). The triphenylmethyl group, while providing excellent regioselectivity for primary hydroxyl groups, is extremely acid labile while silyl ethers are prone to a variety of migrations under some reaction conditions although they are typically more resistant to acid (or base) hydrolysis, depending on the specific silyl ether (Wuts & Greene, 2007).

Benzyl ether protecting groups are also widely employed, however, their introduction into a molecule requires acidic or basic conditions that may disturb other protecting groups (Wuts & Greene, 2007). Like methyl ethers, benzyl groups are often introduced using a Williamson reaction although other alternatives are available such as the use of diphenyldiazomethane (Carey & Sundberg, 2001b).

Benzhydryl ethers, discussed in the next section, have been used to protect hydroxyl groups under mild conditions (thermal decomposition, tin-halide catalysis) using a diazocompound as the benzhydryl-donor (Carey & Sundberg, 2001b; Petursson, 2008; Petursson, 1979). Benzhydryl ethers can be introduced selectively and are easily removed under mild conditions.

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1.3.2 Benzhydryl Ethers as Hydroxyl Protecting Groups

Fluorenyl, benzhydryl, and other diarylmethyl derivatives can be used as protecting groups for carboxylic acids (Wuts & Greene, 2007) and alcohols including vicinal diols; these groups can be introduced to a molecule through either photolysis (Carey & Sundberg, 2001b), thermal degradation (Best et al., 2008), or through a tin halide- mediated reaction of diaryldiazo derivatives of the parent group (Petursson, 2008; Petursson, 1979). The use of benzhydryl ethers as protecting groups is advantageous and can be introduced under mild conditions, which reduces the risk of modifications to other protecting groups, racemizerization, or other unwanted side reactions. Benzhydryl ethers, being secondary ethers, are not as acid labile as tertiary ethers, such as trityl groups and not prone to migration as is the case with some acyl protecting groups. Additionally, benzhydryl ethers can be removed under mild conditions such as catalytic hydrogenolosysis over palladium.

Protections using benzhydryl protecting groups

The diarylmethyl ethers (Figure 20), such as benzhydryl and related derivatives, have been used for alcohol protection using a variety of methods for their introduction. Several methodologies involve the use of concentrated acids and exotic catalysts (Wuts & Greene, 2007) which may not be appropriate for molecules with sensitive moieties.

A B C D MeO OMe Cl Cl

O O O O

Figure 20 - Structures of four common diarylmethyl ethers; (A- benzhydryl, B-, C-, D-)

Diarylmethyl ethers are labile under strongly acidic conditions. However, the conditions under which these ethers are labile has not been as broadly defined for more widely employed protecting groups. Diarylmethyl ethers have been reportedly

36 Háskólinn á Akureyri Viðskipta- og Raunvísindasvið

removed by relatively mild conditions such as hydrogenolysis over 10% palladium on carbon (Petursson, 2008).

The use of mild conditions such as the thermolysis or tin(II) halides catalyzed decomposition of diazo compounds can be used to introduce benzhydryl ethers to hydroxyl groups (Petursson, 2008). The diphenyldiazomethane is more stable than diazomethane due to the diazo group being attached to a secondary carbon and can be used to introduce a benzhydryl moiety to alcohols. Diphenyldiazomethane is inconvenient to handle due to its low melting point (~25 ˚C). Diphenyldiazomethane derivatives, such as dimethyl-, dichloro-, dinitro-, and di-methoxydiphenyl diazomethane are more convenient to handle due to their higher melting point.

The thermolysis of the diazo diarylmethyl compounds can be achieved by refluxing with the alcohol substrate in a high-boiling solvent which leads to the generation of a highly reactive carbene which will react non-selectively with hydroxyl groups present. The mechanism, shown in Figure 21, involves the formation of the reactive carbene, subsequent attack of an intramolecular hydride transfer resulting in the corresponding ether.

Ph Ph or h C N N C + N2 Ph Ph Carbene

Ph Ph Ph R O C + C O R CH O R Ph Ph Ph H H Ether Carbene Ylide Figure 21 - Mechanism of benzhydryl ether formation from corresponding diazo compound via thermolysis

The use of palladium(II) salts as a catalyst for the diphenyldiazomethane or dimethyldiphenyldiazomethane ether protection and deprotection for some primary and some secondary hydroxyl groups has been reported (Bikard et al., 2008). The

37 Háskólinn á Akureyri Viðskipta- og Raunvísindasvið

appeal of this methodology is the use of mild conditions. More recently, the use of a copper(II) bromide catalyst has been reported to catalyze the protection of primary and some secondary alcohols with the derivatives of benzhydryl ether group and the corresponding deprotection in good yields at room temperature (Mezaache et al., 2009). The regioselectivity of the parent alcohol for primary or secondary hydroxyl groups was not explored for either catalyst.

Selective introduction of benzhydryl protecting groups using tin(II) halide-mediated reactions

The use of tin and tin derivatives to facilitate the regioselective protection of hydroxyl groups is well documented. While such methods are useful, stoichiometric quantities of the tin derivative are required. Tin(II) halides can be used to introduce benzhydryl groups to vicinal diol systems using diaryldiazo derivatives with good regioselectivity under milder (room temperature) conditions using diaryldiazomethane compounds. The tin(II) halide-catalyzed reaction works only with cis-vicinal systems in furanoses but both cis- and trans- react in pyranoses (Petursson, 1979; Pétursson, 2001).

In primary 1,2-diol systems, regioselectivity was dependent upon the structure of the diazo compound being introduced with bis(4-methylphenyl)-diazomethane and bis(4- methoxyphenyl diazomethane showing the highest selectivity for the primary hydroxyl group.

The mechanism of tin(II) halide-catalyzed alkylations with diaryldiazomethanes is not well understood. Three potential mechanisms involve the formation of an electrophilic tin-carbene complex, the formation of a diol-tin complex and the formation of a diol-tin intermediate. It is possible that a combination of several mechanisms are involved.

It is known that transition metals react with diazo compounds leading to the formation of a carbene and molecular nitrogen, which is a good leaving group due to the favorable thermodynamics of lysis, being a gas, and having a neutral character. Alternatively, transition metals, such as rhodium, palladium, molybdenum, copper,

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and tin, can form metal-carbene complexes which are electrophilic in character (Carey & Sundberg, 2001) as shown in Figure 22.

R R R M + C N N M C M C - N R 2 R R Figure 22 - Formation of electrophilic metal-carbene complexes (Carey & Sundberg, 2001a)

Both the free carbine and the metal-carbene complex are reactive species and present two potential pathways for the introduction of moieties into substrates. It has been observed that highly reactive diaryldiazo compounds, notably bis(4- methylpheny)diazomethane and bis(4-methoxyphenyl)diazomethane, tend to be much more regioselective diazo-bis(phenyl)-, bis(4-chlorophenyl)-, and bis(4- nitrophenyl)methane and diazafluroene when used in combinations with tin(II) halide catalysts.

Alternately, it has been proposed that the stannous halide catalyzed reaction proceeds via the formation of a stannous halide-diol complex as opposed to a metal-carbene complex, as shown in Figure 23 (Petursson, 2001).

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Figure 23 - Proposed partial mechanism of stannous chloride-mediated reaction of diaryldiazo compounds with vicinal diols (modified from Pétursson, 2001)

Petursson (2001) suggested that the metal-carbene complex is less likely due to the destruction of the catalyst and the kinetics of the model system appeared to proceed through a typical pseudo-first order mechanism.

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2 Experimental

2.1 General methods

All reactions were performed at ambient temperature in the absence of light without special precautions unless otherwise noted. All melting points were determined using a glass capillary and a Theile tube; values are uncorrected. 1H and 13C NMR spectrometry was performed on a Bruker AV400 using tetramethylsilane (TMS) as an external standard. 3Å molecular sieves were oven dried at 250 °C prior to use. 1,2- Dimethoxyethane (DME) was refluxed over sodium for two hours, distilled, and stored over sodium prior to use. Dichloromethane was dried over 3Å molecular sieves prior to use. Diisopropyl ether was stored over blue-indicating 3Å molecular sieves prior to use. Anhydrous methanol was prepared by drying over 3Å molecular sieves for 6 hours and distilling prior to use. Petroleum ether was of the 40 -60˚C boiling range, all reagents were obtained from Sigma-Aldrich unless otherwise noted; solvents were reagent grade or better. Optical rotations were obtained using an Eloptron (Schmidt-Haenson, Germany) for solutions dissolved in chloroform at a pathlength of 1 dm at ambient temperature relative to the Sodium-D line.

The following immobilized lipases were obtained from Fluka; Pseudomonas cepacia lipase (PCL,) immobilized on ceramic (Fluke 17261, 15156 U/g), Pseudomonas fluorescens (PFL) immobilized on Sol-Gel-AK (>40 U/g), Mucor miehei (MML, Lipozyme®) immobilized on ceramic (Fluka 62350, 40 U/g), Candida cylindracea (CCL) immobilized on Sol-Gel-AK (Fluka 62278, 13 U/g), and Candida antarctica (CAL) immobilized on ceramic (Fluka 73940, 2.5 U/g).

Thin layer chromatography (TLC) plates were Merck silica-60 containing F254 fluorescent indicator on aluminum backing; plates were visualized using Cerium (IV) ammonium molybdenum sulfate (CAM). Preparative flash chromatography was performed using Silica-60-packed 2.2 by 20 cm columns unless otherwise noted.

HPLC analysis was performed using a Shimadzu UFLC system equipped with a SPD-

20 UV-Vis detector at 254 nm, DGU-20A3 on-line degasser, LC-20A solvent pump,

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Sigma Supelco Discovery C18 column (150x4.6 mm, 3 µm packing) equipped with equivalent pre-column, and a manual injector equipped with a 20 µL sample loop. The solvent system consisted of 20% un-buffered Type I Laboratory water and 80% methanol; mobile phase solvents were degassed by sonication prior to use.

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2.2 Preparation of diazo compounds

Hydrazones were prepared from the corresponding ketone as described by Petursson (1979).

2.2.1 Preparation of diazo[bis(4-methylphenyl)]methane 29.6 g (132 mmol) of bis-(4-methylphenyl)methyl hydrazone was dissolved in

600 mL of diethyl ether; 30 g of dry Na2SO4 and 60.0 g (277 mmol, 2.1 eq.) of mercury(II) oxide was added followed by 10 mL of KOH saturated ethanol. The reaction was stirred magnetically for 1.5 hour and followed by TLC (hexane/ethyl acetate 4:1). The mixture was filtered through Whatman #1 filter paper and the solvent was reduced to approximately 150 mL. The solution was crystallized by cooling to -40 ˚C; the resultant purple crystals were isolated by vacuum filtration yielding 29.33 g of the title compound in 79.3% yield, mp = 104˚C.

2.2.2 Preparation of diazo[bis(4-methoxyphenyl)]methane 19.0 g of bis-(4-methoxyphenyl)methyl hydrazone (74.2 mmol) and 50 g of

dry Na2SO4 was dissolved in 400 mL of diethyl ether; 40.0 g (184.6 mmol, 2.5 eq) of mercury(II) oxide was added followed by 10 mL of KOH saturated ethanol. The mixture was stirred magnetically for 5 hours and followed by TLC (Hexane/Ethyl acetate 4:1). The solution was then filtered through two pieces of Whatman #1 filter paper; the volume of the solution was then reduced to approximately 200 ml by a rotary evaporator (30˚C, -1 bar) . The solution was then crystallized by cooling to -40˚C; the resultant shiny purple flakes were collected by vacuum filtration yielding 15.32 g of the title compound in 81.3% yield, mp = 101-103 ˚C.

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2.3 Protection studies towards primary benzhydrylation of vicinal diols

2.3.1 Benzhydralation experiments on 1,2-propanediol

rac-[1-O-bis(4-methylphenyl)methyl]–propan-2-ol (SnCl2 catalyzed)

OH O OH 1.5 eq bis(4-methylphenyl)diazomethane + OH SnCl2 O DME OH

(Major) (Minor)

0.7609 g (10 mmol) of 1,2-propanediol was dissolved in 200 mL of DME to which

250 mg of SnCl2 were added followed by 3.3640 g (15 mmol, 1.5 eq) of diazo[bis(4- methylphenyl)]methane. After 45 minutes, TLC examination (hexane/ethyl acetate 4:1) revealed that all of the starting material had reacted. The material was chromatographed on a 4.4x20 cm column of silica gel packed with 9:1 petroleum ether/ethyl acetate. 21x50 mL fractions were collected by eluting 9:1 hexane/ethyl acetate. Fraction s20-50 were pooled and the solvent was removed using a rotary evaporator (45˚C, -1 bar) yielding 2.3736 g (87.8%) of a light yellow oil of an unresolved mixture of the 1- and 2-ether in ratio of 4:1.

1 H NMR (400 MHz, CDCl3 J = 12.79, 6.38, 6.38, 3.19, 3.19 Hz, 1H), 3.37 (dd, J = 9.40, 3.10 Hz, 1H), 3.19 (dd, J = 9.32, 8.17 Hz, 1H), 2.25 (d, J = 6.64 Hz, 1H), 1.06 (t, J = 5.91, 5.91 Hz, 1H), 7.16-7.03 (m, 1H), 2.41-2.29 (m, 13 1H); C NMR (101 MHz, CDCl3 74.68 (s), 66.72 (s), 18.66 (s), 21.16 (s), 129.16 (s), 126.88 (d), 137.24 (d), 139.17 (s).

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rac-[1-O-bis(4-methylphenyl)methyl]–propan-2-ol (SnBr2 catalyzed)

OH O OH 1.5 eq bis(4-methylphenyl)diazomethane + OH SnBr2 O DME OH

(Major) (Minor) 760.9 mg (10 mmol) of 1,2-propanediol were dissolved in 200 mL of DME; 360.0 mg

of SnBr2 (1.29 mmol) followed by 3.3285 g (15 mmol, 1.5 eq) of diazo[bis(4- methylphenyl)]methane. The reaction was followed by TLC; after 30 minutes the solvent was removed using a rotary evaporator (40 ˚C, -1 bar). The resultant oil was chromatographed on a 4.4 x 24 cm of silica gel packed with 9:1 hexane/ethyl acetate; 98x20 mL fractionswere collected by eluting 9:1 hexane/ethyl acetate. 802.1 mg (30.0% yield) of the 1-ether were isolated in addition to 372 mg of an unresolved mixture of the corresponding 1- and 2-ethers; the overall yield of 1.1741 g (43.4%).

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Separation of 1-dimethyldipheny propane-2-ol ether from 2 ether

OH OH O O TrtCl (1.2 eq) + O + O TEA Ph DCM O OH RT Ph Ph

1.5618 g (5.8 mmol) of the mixture of 1 and 2 ether was dissolved in 25 mL of dichloromethane to which 2.0 mL of triethylamine (TEA) were added. 0.5122 g (1.837 mmol, 1.2 eq) of trityl chloride was added followed by 6.0 mg (0.049 mmol) of DMAP. The reaction was stirred magnetically for 96 hour and followed by TLC (4:1 hexane/ethyl acetate). The solvent was removed on the rotary evaporator (45˚C, -1 bar) and the mixture was chromatographed on a 4.4x20 cm column of silica gel packed with 9:1 petroleum ether/ethyl acetate containing 2% TEA (v/v). 21x50 mL fractions were collected by eluting 9:1 petroleum ether/ethyl acetate. Fractions 11-21 were pooled and the solvent removed to afford 0.9446 g of the 1-ether (82.2% recovery) as a light yellow oil.

1 H NMR (400 MHz, CDCl3) δ ppm 5.39 (s, 1H), 4.08 (dqd, J = 12.86, 6.40, 6.40, 6.39, 3.18 Hz, 1H), 3.50 (dd, J = 9.40, 3.10 Hz, 1H), 3.33 (dd, J = 9.31, 8.15 Hz, 1H), 2.38 (s, 1H), 1.20 (t, J = 8.05, 8.05 Hz, 1H), 7.31-7.23 (m, 1H), 7.19 (d, J = 13 7.71 Hz, 1H), 2.52 (d, J = 10.00 Hz, 1H); C NMR (101 MHz, CDCl3) δ ppm 139.19 (s,1C), 137.24 (d, J = 2.70 Hz,1C), 129.17 (s,1C), 126.89 (d, J = 3.23 Hz,1C), 83.89 (s,1C), 74.69 (s,1C), 66.73 (s,1C), 21.17 (s,1C), 18.68 (s,1C)

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2.3.2 Benzhydrylation experiments on 3-phenoxy-1,2-propanediol Two benzhydrylations of the 3-phenoxy-1,2-propanediol were conducted as described by SP.

3-O-phenoxy-1-O-(bis(4-methylphenyl)methyl)-propan-2-ol (SnCl2 catalyzed)

bis(4-methylphenyl)diazomethane (1.5 eq) PhO OH O PhO OH PhO O DME + OH OH SnCl2

(Major) (Minor) 672.8 mg (4 mmol) of 3-phenoxy-1,2-propanediol were dissolved in 80 mL of DME;

25 mg of SnCl2 were added followed by 1.342 g (1.5 eq, 6 mmol) of diazo[bis(4- methylphenyl)]methane. The reaction was followed by TLC (hexane/ethyl acetate 4:1).After 45 minutes, the DME was removed on the rotary evaporator and the resultant mixture was chromatographed on a 4.4 x 20 cm column of silica gel packed with 19:1 hexane/ethyl acetate. The overall yield was 88.4%; 334.9 mg of corresponding title compound (49.8% yield), 158.2 mg (23.5%) of an unresolved mixture, and 101.6 mg (15.1) of the 2-ether. 1H NMR and 13C were identical to previously published spectra.

3-O-phenoxy-1-O-(bis(4-methylphenyl)methyl)-propan-2-ol (SnBr2 catalyzed)

bis(4-methylphenyl)diazomethane (1.5 eq) PhO OH PhO OH O PhO O + DME OH OH SnBr2

(Major) (Minor)

672.8 mg (4 mmol) of 3-phenoxy-1,2-propanediol was dissolved in 80 mL of DME;

36 mg of SnBr2 were added followed by 1.332 g (1.5 eq, 6 mmol) of diazo[bis(4- methylphenyl)]methane. The reaction was followed by TLC (hexane/ethyl acetate 4:1).After 1 hour, the DME was removed on the rotary evaporator and the resultant mixture was chromatographed on a 4.4 x 20 cm column of silica gel packed with 19:1 hexane/ethyl acetate. The overall yield was 69.0%; 889.6 mg of corresponding title

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compound (61.4% yield), 28.9 mg of an unresolved mixture, and 82.1 mg of the 2- ether. 1 H NMR (400 MHz, CDCl3) d ppm 5.57 (s, 1H), 4.01 (t, J = 5.09, 5.09 Hz, 1H), 2.24 (d, J = 6.10 Hz, 1H), 3.85 (dt, J = 9.80, 9.67, 5.56 Hz, 1H), 3.79-3.63 (m, 1H), 6.91- 6.74 (m, 1H), 7.05 (dd, J = 14.25, 7.95 Hz, 1H), 7.20-7.14 (m, 1H), 1.80 (t, J = 5.74, 5.74 Hz, 1H).

Effort towards 1-O-bis(4-methoxyphenyl)dimethyldiphenyl –propan-2-ol (SnCl2 catalyzed)

OMe

bis(4-methoxyphenyl)diazomethane PhO OH O PhO OH PhO O DME + OH OH SnCl2 MeO OMe OMe (Major) (Minor) 373.1 mg (2 mmol) of 3-O-phenoxy-1,2-propanediol was dissolved in 20 mL of DME

to which 6 mg of SnCl2 were added followed by 3.26 g (12.8 mmol, 6.4 eq) of diazo[bis(4-methoxyphenyl)]methane . The reaction was followed by TLC as previously described. DME was removed as previously described and the resultant mixture was chromatographed on a 2.5 x 25 cm column of silica gel packed with 9:1 hexane/ethyl acetate. Fractions containing suspected product were pooled and the mixture was re-chromatographed using a 19:1 to 9:1 gradient of hexane/ethyl acetate. Pooled fractions yielded 127.3 mg (~16.1%) of an impure ether.

Effort towards 1-O-bis(4-methoxyphenyl)dimethyldiphenyl –propan-2-ol (SnBr2 catalyzed)

OMe

PhO OH 1.5 eq bis(4-methylphenyl)diazomethane O PhO OH PhO O + DME OH OH SnBr2 MeO OMe OMe

372.8 mg (2 mmol) of 3-phenoxy-1,2-propanediol and 72 mg (mmol) of SnBr2 was dissolved in 80 mL of DME. 3.75 g (14.7 mmol, 7.5 eq.) of diazo[bis(4- methoxyphenyl)]methane was added incrementally until all of the starting diol had been consumed. The reaction was monitored by TLC (hexane/ethyl acetate 4:1). The

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reaction mixture was separated on silica gel (4.4 cm x 15 cm) packed with 9:1 hexane/ethyl acetate. Products were eluted by running a gradient of 9:1 to 4:1 hexane ethyl acetate. Pooled fractions yielded 172.6 mg of an impure ether derivative (~21.8% yield).

2.3.3 Primary benzyhydrylation of 3-chloro-1,2-propanediol

1-O-bis(4-methylphenyl)-3-chloro-propan-2-ol (SnCl2 catalyzed)

OH Cl O OH 1.5 eq bis(4-methylphenyl)diazomethane + Cl OH SnCl2 O DME Cl OH

(Major) (Minor)

453.6 mg (4 mmol) of 3-chloro-1,2-propanediol was dissolved in 80 mL of DME. 100

mg of SnCl2 were added followed by 1333.3 mg (6 mmol, 1.5 eq) of diazo(bis[4- methylphenyl)methane. After 1 hour and a TLC examination (4:1 petroleum ether/ethyl acetate), the starting material had been consumed. The products were chromatographed on a 4.4x12 cm column of silica gel using 4:1 petroleum ether/ethyl acetate and collected at fifty 50 mL fractions. Fractions 5-8 yielded 1.2016 g (98.5% overall yield) of an unresolved mixture.

1 H NMR (400 MHz, CDCl3 ppm 5.26 (s, 1H), 4.05 (q, J = 7.14, 7.14, 7.14 Hz, 1H), 3.98-3.91 (m, 1H), 3.61 (dd, J = 11.10, 5.33 Hz, 1H), 3.57-3.51 (m, 1H), 3.51- 3.44 (m, 1H), 2.41 (t, J = 5.13, 5.13 Hz, 1H), 2.24 (s, 1H), 1.97 (s, 1H), 1.18 (t, J = 7.15, 7.15 Hz, 1H), 7.07 (dd, J = 14.47, 8.31 Hz, 1H), 7.12 (t, J = 7.04, 7.04 Hz, 1H), 13 7.20-7.15 (m, 1H); C NMR (101 MHz, CDCl3 ppm 129.19 (d, J = 2.78 Hz,1C), 126.86 (s,1C), 77.39 (s,1C), 77.08 (s,1C), 76.76 (s,1C), 21.13 (d, J = 6.38 Hz,1C)

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Separation of 1-dimethyldipheny 3-chloro-propane-2-ol ether from 2 ether (pg 40)

OH OH Cl O Cl O TrtCl (1.2 eq) + O + O TEA Ph DCM Cl O Cl OH Ph Ph

1.2 g (4.1 mmol) of the mixture of 1 and 2 ethers of 3-chloro-1,2-propanediol was dissolved in 20 mL of DCM to which 10 mL of TEA, 557 mg of trityl chloride (2 mmol) and 30.0 mg of DMAP was added. The reaction was allowed to proceed overnight. The DCM was removed by rotary evaporator (25°C, -1 bar); the mixture was loaded onto a 4.4x12 cm column of silica gel packed with 9:1 petroleum ether/ethyl acetate containing 1% (v/v) TEA. The products were eluted using 4:1 petroleum ether/ethyl acetate. Fractions were pooled yielding 677.1 mg of the 1-ether (56.1% yield).

1H NMR (400 MHz, Solvent ppm 5.27 (s, 1H), 3.94 (td, J = 10.78, 5.38, 5.38 Hz, 1H), 3.65-3.52 (m, 1H), 3.52-3.44 (m, 1H), 2.40 (d, J = 5.91 Hz, 1H), 2.25 (s, 1H), 7.08 (dd, J = 19.55, 8.96 Hz, 1H), 7.16-7.10 (m, 1H), 7.21-7.16 (m, 1H), 7.27-7.21 (m, 1H), 7.36 (dd, J = 5.33, 3.36 Hz, 1H);

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2.4 Lipase-catalyzed resolution of partially protected diols

2.4.1 Comparison of lipases for 2(R) acetylation of rac-1[bis(4- methylphenyl)]methoxypropane-2-ol Five lipases (PCL, PFL, MML, CCL, and CAL) were screened for their ability to resolve a racemic mixture of rac-1[bis(4-methylphenyl)]methoxypropane-2-ol as a model compound for subsequent resolutions.

OAc OH OH O O O Lipase +

Diisopropyl ether Vinyl Acetate

(R)-1-(dip-tolylmethoxy)propan-2-yl acetate (S)-1-(dip-tolylmethoxy)propan-2-ol

33.8 mg (0.125 mmol) of rac-1[bis(4-methylphenyl)]methoxypropane-2-ol was dissolved in 5 mL of diisopropyl ether and 23 µL (0.25 mmol, 2 eq) of vinyl acetate in a 5 mL conical flask. 5 mg of lipase immobilized on ceramic beads were added and the reaction stirred magnetically. Samples were removed periodically and analyzed by HPLC after diluting 24:1 with mobile phase as described in Section 2.1.

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2.4.2 Kinetic Resolution of rac-1[bis(4-methylphenyl)]methoxypropane-2-ol

OAc OH OH O O O PCL +

Diisopropyl ether Vinyl Acetate

Product I Product II

284.4 mg (1.05 mmol) of the title compound was dissolved in 40 mL of diisopropyl ether to which 184 µL (2.3 mmol, 2 eq) of vinyl acetate was added. 40.0 mg of immobilized PCL was added and stirred magnetically. The reaction was followed by HPLC. After 96 hours, the solvent was carefully transferred to a round bottom flask and the solvent removed (35°C, -1 bar). The mixture was chromatographed on a column of silica gel (2.2x20 cm) using 9:1 petroleum ether/ethyl acetate and collected as 30x15 mL fraction. Fractions 5-11 (acetate) were pooled to yield 162.1 mg (99.0%) of the corresponding (2R)-acetate (Product I) and 138.3 mg (97.3%) of the (2S)- alcohol (Product II).

(2R)-1[bis(4-methylphenyl)]methoxy-2-acetoxypropane (Product I) 1H NMR (400 MHz, CDCl3) ppm 5.24 (s, 1H), 5.06 (d p, J = 6.37, 6.36, 6.36, 6.36, 4.69 Hz, 1H), 3.39 (dq, J = 10.23, 10.23, 10.23, 5.30 Hz, 1H), 2.25 (d, J = 9.37 Hz, 1H), 1.96 (d, J = 2.53 Hz, 1H), 1.17 (t, J = 5.39, 5.39 Hz, 1H), 7.13 (dd, J = 8.02, 1.51 Hz, 1H), 7.04 (d, 13 J = 7.98 Hz, 1H); C NMR (101 MHz, CDCl3 ppm 129.06 (s,1C), 126.88 (d, J = 6.09 Hz,1C), 77.37 (s,1C), 77.06 (s,1C), 76.74 (s,1C).

(2S)-1[bis(4-methylphenyl)]methoxypropane-2-ol (Product II) 1H NMR (400 MHz, CDCl3 ppm 5.26 (s, 1H), 3.95 (dqd, J = 12.82, 6.40, 6.40, 6.40, 3.13 Hz, 1H), 3.37 (dd, J = 9.41, 3.10 Hz, 1H), 3.19 (dd, J = 9.37, 8.13 Hz, 1H), 2.35 (s, 1H), 2.22 (d, J = 18.77 Hz, 1H), 1.06 (d, J = 6.39 Hz, 1H), 7.06 (t, J = 6.93, 6.93 Hz, 1H), 7.14 (d, J = 6.96 Hz, 1H);

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2.4.3 Kinetic Resolution of 1-dimethyldiphenyl ether of rac-3-phenoxy-1[bis(4- methylphenyl)]methoxypropane-2-ol

PCL PhO O PhO O + PhO O OH Vinyl acetate OAc OH Diisopropyl ether

Product I Product II

Kinetic experiments: 90.9 mg (0.25 mmol) of the title compound was placed in a 5 mL round bottom flask and dissolved in 2.5 mL of diisopropylether. 46 µL (0.500 mmol, 2 eq) of vinyl acetate were added followed by 10 mg of lipase from Pseudomonas cepacia (Fluka 17261). The mixture was stirred magnetically and followed by HPLC.

The reaction was repeated on a larger scale by placing 362.4 mg (1 mmol) of the title compound in a round bottom flask to which 10 mL of diisopropyl ether followed by 184 µL (2.3 mmol, ~2 eq) of vinyl acetate and 200 mg of lipase from P. cepacia (Fluka 17261). The mixture was magnetically stirred and followed by HPLC. After 7 days, the peak ratio of the two products was 50:50; the solvent was removed using a rotary evaporator (45 ˚C, -1 bar) and chromatographed on a 2.2 x 20 cm column of silica gel packed with 4:1 hexane/ethyl acetate. 4:1 hexane/ethyl acetate was run and collected as 10 mL fractions. Fraction 7-11 were pooled yielding 201.3 mg (99.5%) of Product I, a clear, colorless oil. Fraction 12 contained 16.8 mg of a mixture of Product I and II. Fractions 13-22 were pooled yielding 141.0 mg (94.8%) of Product II, a clear colorless oil.

(2R)-3-phenoxy-1[bis(4-methylphenyl)]methoxy 2-acetoxypropane (Product I) (Acetate) 1H NMR (400 MHz, CDCl3 ppm 5.38 (s, 1H), 4.24 (dd, J = 4.78, 3.14 Hz, 1H), 3.77-3.72 (m, 1H), 2.36 (s, 1H), 2.12 (s, 1H), 5.42 (dd, J = 10.09, 5.03 Hz, 1H), 6.94 (d, J = 8.50 Hz, 1H), 7.04-6.98 (m, 1H), 7.15 (dd, J = 8.04, 3.86 Hz, 1H), 7.27-7.21 (m, 1H), 7.32 (dd, J = 8.52, 7.82 Hz, 1H); 13C NMR (101 MHz, CDCl3 ppm 129.10 (s,1C), 126.89 (d, J = 4.45 Hz,1C), 77.38 (s,1C), 77.07 (s,1C), 76.75 (s,1C), 21.16 (s,1C). (2S)-3-phenoxy-1[bis(4-methylphenyl)]methoxy methoxypropane-2-ol (Product 1 II) H NMR (400 MHz, CDCl3 ppm 5.41 (s, 1H), 4.15-4.07 (m, 1H), 3.73-3.65 (m, 1H), 2.38 (d, J = 9.12 Hz, 1H), 7.36-7.30 (m, 1H), 7.28-7.23 (m, 1H), 7.18 (t, J = 10.36, 10.36 Hz, 1H), 7.01 (t, J = 7.35, 7.35 Hz, 1H), 6.95 (d, J = 8.10 Hz, 1H), 2.58 (d, J = 3.90 Hz, 1H), 4.29-4.22 (m, 1H); 13C NMR (101 MHz, Solvent) d ppm 129.17 (s,1C), 126.90 (s,1C), 114.64 (s,1C), 77.39 (s,1C), 77.07 (s,1C), 76.75 (s,1C)

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2.4.4 Kinetic Resolution of rac-3-chloro-1[bis(4- methylphenyl)]methoxypropane-2-ol

PCL Cl O + Cl O Cl O OAc OH Vinyl acetate OH Diisopropyl ether

Product I Product II

314.3 mg of the title compound was dissolved in 40 mL of diisopropyl ether to which 184 µL of vinyl acetate was added. 40.0 mg of ceramic immobilized PCL was added. The reaction was monitored by HPLC as previously described. After 6 days, the ceramic beads were removed by filtration and the resultant mixture was evaporated in vacuo prior to being chromatographed as described in previous sections. 168.0 mg of Product I (98.9%) and 146.5 mg of Product II (93.2% yield).

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2.5 Deprotection of (2R)-Acetyl Derivatives

The acetate groups of benzhydryl derivatives were removed using a Zemplén de-O- acetylation as described by Wuts & Greene (2007) using catalytic quantities of sodium in anhydrous methanol.

2.5.1 Deprotection of (2R)-1[bis(4-methylphenyl)]methoxy-2-acetoxypropane

OAc OH O Na O MeOH

101.7 mg (0.33 mmol) of the title compound was dissolved in 10 mL anhydrous methanol containing 1 mg of sodium. The reaction was stirred under positive pressure of nitrogen for 6 hours and followed by TLC (ethyl acetate). The solvent was removed and the reaction mixture was chromatographed on a column of silica gel packed with hexane/ethyl acetate (1:1). Pooled fractions yielded 77.1 mg (97.6%) of the resultant 2R-alcohol. 1H and 13C NMR was identical to the previously described 2S-alcohol.

2.5.2 Deprotection of (2R)-3-phenoxy-1[bis(4-methylphenyl)]methoxy 2- acetoxypropane

Na PhO O PhO O OAc MeOH OH

104.4 mg (0.26 mmol) of the title compound was handled as previously described in Section 2.5.1 for 16 hours. The reaction mixture was chromatographed as described in the previous section except using hexane/ethyl acetate (3:1). Pool fractions yielded 93.0 mg (99.5%) of the 2R-alcohol with 1H and 13C NMR was identical to the previously described 2S-alcohol.

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2.5.3 Deprotection of (2R)-3-chloro-1[bis(4-methylphenyl)]methoxy 2- acetoxypropane

Na Cl O Cl O OAc MeOH OH

98.8 mg (28.5 mmol) of the title compound was dissolved in anhydrous methanol containing 10 mg of sodium. The reaction was stirred under positive pressure of nitrogen for 12 hours and followed by TLC (ethyl acetate). The solvent was removed and the reaction mixture was chromatographed on a column of silica gel packed with ethyl acetate. Collected as 10 mL fractions. 83.6 mg (96.3%) of the resultant 2R alcohol was obtained. 1H and 13C NMR was identical to the previously described 2S- alcohol.

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3 Results

3.1 Preparation of diazocompounds

Bis(4-methylphenyl)diazomethane and bis(4-methoxyphenyl)diazomethane were prepared in 79.3 and 81.3% yield, respectively. Melting point values for these two diazo compounds were comparable to values previously published in the literature.

3.2 Protection studies towards primary benzhydrylation of vicinal diols

The initial protection of partially protected diols was carried out on bis(4-

methylphenyl)diazomethane using either SnCl2 or SnBr2 as a catalyst; the resultant yields and selectivity for the primary or secondary hydroxyl group of 1,2-propanediol and related compounds are reported in Table 4. Overall yields ranged from 43.4 to 98.5% with reaction times under 60 minutes in all cases. Selectivity for the primary hydroxyl group varied both as a function of the diol substrate and the tin(II) halide catalyst used.

Table 4 - Tin(II) halide benzhydrylation of vicinal diols using bis(4-methyl phenyl)diazomethane

Diol substrate Catalyst Total yield 1-ether : 2-ether Reaction (%) ratio time (min) 1,2-propanediol SnCl2 87.8 2.8:1 45 SnBr2 43.4 7.5:1 30 3-phenoxy-1,2-propanediol SnCl2 88.4 4.1:1 45 SnBr2 69.0 9.1:1 <30 3-chloro-1,2-propanediol SnCl2 98.5 10.2:1 (HPLC) 60

The use of diazo(bis[4-methoxyphenyl)methane was only attempted for 1,2-

propanediol and 3-phenoxy-1,2-propanediol using SnBr2 as a catalyst. While reaction times were short, a much larger excess of the diazo compound was required to

completely react with the diol substrates. Yields were 16.1% and 21.8% for the SnCl2

and SnBr2-catalyzed reactions, respectively. The reaction and the presence of two co- eluting spots complicated separation and subsequent characterization.

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3.3 Kinetic Resolution of Partially Protected Diols

The ability of commercially available lipases to resolve rac-1[bis(4- methylphenyl)]methoxypropane-2-ol was screened and the data presented in Section 3.3.1. The PCL-catalyzed resolution of rac-1[bis(4-methylphenyl)]methyl ether derivatives of 1,2-propanediol, 3-chloro-1,2-propanediol, and 3-phenoxy-1,2- propanediol and subsequent deprotections of the (2R)-ester are discussed in Section 3.3.2.

3.3.1 Comparison of lipases for 2(R) acetylation of rac- 1[bis(4-methylphenyl)]methoxypropane-2-ol

1-bis(4-methylphenyl)methyl-propan-2-ol was used as a substrate to screen for lipases with sufficiently accommodating active sites to allow the entry of similar partially protected diols. Kinetic plots were generated for each of the reactions using the five lipases over the 7 day experiment period from the HPLC data. The data from PCL, PFL, MML, and CAL are presented in Figure 24 to Figure 27, respectively. The use of (CCL) did not result in the formation of the corresponding acetate over the 7 day reaction period (data not shown). Summary chromatographs are presented as Figure 30 to Figure 34 in Appendix A. All lipase catalyzed reactions showed pseudo first- order kinetics (data not shown).

PCL catalyzed the acylation of 50% of 1-bis(4-methylphenyl)methyl-propan-2-ol within 24 hours and PFL within 48 hours. Neither reaction proceeded beyond 50% of the starting material over the experimental period. CAL rapidly acylated the substrate but consumed more than half of the starting material whereas MML slowly proceeded reacting with about half of the starting material after approximately 5 days.

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100,0 100,0 90,0 90,0

Starting Material 80,0 80,0 Starting Material 70,0 Product (2-R-Ac) 70,0 Product (2-R-Ac) 60,0 60,0 50,0 50,0 40,0 40,0

30,0 30,0 Concentration(%) 20,0 Concentration(%) 20,0 10,0 10,0 0,0 0,0 0 2000 4000 6000 8000 10000 0 2000 4000 6000 8000 10000 Time (min) Time (min)

Figure 24 – Pseudomonas cepacia Lipase (PCL) catalyzed kinetic resolution Figure 25 – Pseudomonas fluorescens Lipase (PFL) catalyzed kinetic resolution of 1-bis(4- of 1-bis(4-methylphenyl)methyl-propan-2-ol methylphenyl)methyl-propan-2-ol 100,0 100,0

90,0 90,0

80,0 Starting Material 80,0 70,0 Product (2-R-Ac) 70,0 60,0 60,0 50,0 50,0 Starting Material 40,0 40,0 Product (2-R-Ac)

30,0 30,0 Concentration(%) 20,0 Concentration(%) 20,0 10,0 10,0 0,0 0,0 0 2000 4000 6000 8000 10000 0 2000 4000 6000 8000 10000 Time (min) Time (min)

Figure 26 – Mucor mehei Lipase (MML) catalyzed kinetic resolution of 1- Figure 27 – Candida antarctica Lipase (CAL) catalyzed kinetic resolution of 1-bis(4- bis(4-methylphenyl)methyl-propan-2-ol methylphenyl)methyl-propan-2-ol

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3.4.2 PCL-mediated chemoenzymatic resolution and deprotection of rac- 1[bis(4-methylphenyl)]methyl ether derivatives

The yields of the PCL-catalyzed acylation of the partially protected diol substrates are presented in the table below. The (R) enantiomer was isolated as the corresponding acetate whereas the (S) enantiomer was not acylated by the lipase.

Table 5 – Summary of yields from PCL-catalyzed resolutions of rac-1-[bis(4-methylphenyl)]methyl ether derivatives of selected diols

Partially protected compound Enantiomer Yield (%) rac-1[bis(4-methylphenyl)]methoxypropane-2-ol (R) 99.0 (S) 97.3 rac-3-phenoxy-1[bis(4-methylphenyl)]methoxy 2- (R) 99.5 acetoxypropane (S) 94.8 rac-3-chloro-1[bis(4-methylphenyl)]methoxy 2-acetoxypropane (R) 98.9 (S) 93.2

The reaction kinetics of the enantioselective acylation of rac-3-phenoxy-1[bis(4- methylphenyl)]methoxy 2-acetoxypropane and 3-chloro-1[bis(4- methylphenyl)]methoxy 2-acetoxypropane are presented in the figures. The reaction kinetics of the 1,2-propanediol reaction were obtained but were nearly identical to those presented in the previous section. Both resolutions presented below follow pseudo-first order kinetics (data not shown).

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100,0 90,0 Starting Material 80,0

70,0 Product (2-R-acetate) 60,0 50,0 40,0

%Concentration 30,0 20,0 10,0 0,0 0 5000 10000 15000 20000 Time (min) Figure 28 - PCL-catalyzed resolution of 3-phenoxy-1[bis(4-methylphenyl)]methoxy 2-acetoxypropane

100,0 90,0 Starting Material 80,0 Product (2-R-Ac)

70,0 60,0 50,0 40,0

Concentration(%) 30,0 20,0 10,0 0,0 0 2000 4000 6000 8000 10000 12000 Time (min)

Figure 29 - PCL-catalyzed resolution of rac-3-chloro-1[bis(4-methylphenyl)]methoxypropane-2-ol

The isolated (R)-acetate of each compound was deprotected and yielded the corresponding free alcohol in near analytical yields; recoveries from the deprotection were greater than 97% in all cases.

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4 Discussion

4.1 Preparation of diazo compounds

The two diazo compounds used in this study were prepared in good yield in keeping with previous preparations of these materials with melting points slightly lower than those previously obtained. The lower yields are likely due to the loss of material upon filtration of the crystallized material One major shortcoming of the preparation of these diazo compounds is the generation of mercury waste.

4.2 Protection studies towards primary benzhydrylation of vicinal diols

The dimethyl and dimethoxy derivatives were chosen for this study due to their short reaction times and high regioselectivity observed in protection studies of other vicinal diol substrates such as partially protected monosaccharides ( Petursson, 1979) and their previous use in resolving 1,2-vicinal diols (Petursson & Jonsdottir, 2012; Petursson, 2009).

Comparison of tin(II) halide catalysts

Two tin(II) halide catalysts were selected in the hope that the regioselectivity of one of the catalysts would present a clear preference for the formation of the corresponding 1-benzhydryl ether from bis(4-dimethylphenyl)diazomethane. Indeed,

the use of SnBr2 showed higher 1-ether selectivity in the case of both 1,2-propanediol

and 3-phenoxy-1,2-propanediol as compared to SnCl2-catalyzed reactions. Previously

published studies using SnBr2-catalyzed reactions with diazo compounds to protect 1,2-propanediol have shown that regioselectivity was not complete necessitating the removal of the unwanted 2-ether (Petursson & Jonsdottir, 2012; Petursson, 2009).

However, the yields of SnBr2-catalyzed reactions were consistently lower than those

using SnCl2 and the evolution of a dark-brown component with a strong halogen-like smell was observed suggesting the presence of molecular bromine. The reaction mixture was checked on a TLC plate which showed a red-brown spot of a mobility of

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less than 0.1; co-spotting with authentic bromine resulted in a similar behavior. Tin halides, particularly tin bromides, are known to decompose into elemental tin and halogens in the presence of light and heat through a radical mechanism. It is

speculated that the SnBr2-catalyzed reactions liberated molecular bromine which may have resulted in lower yields. This complication has not before been observed with

other reactions catalyzed by SnBr2 on other diols, such as partially protected monosaccharides (Scully and Petursson, unpublished results). The author speculates

that the exclusion of molecular oxygen may prevent the degradation of SnBr2 and subsequent work has excluded oxygen and moisture by performing reactions under nitrogen.

Due to the low yields and the formation of a dark-brown color, SnBr2-catalyzed

reactions were discontinued in favor of using the SnCl2 despite the latter’s lower selectivity.

The SnBr2 catalyst seemed to drive the reaction to completion more quickly than the

SnCl2. The author suggests that a potential reason for the difference in behavior is due to the strength of catalyst binding to the more acidic hydroxyl group differing

between the two catalysts. SnCl2 may bind to a vicinal diol more strongly due to the higher electronegativity (and thus electron withdrawing potential) of the two chloro

groups (EN = 3.5) whereas the electronegativity of the bromides of SnBr2 is lower (EN = 3.0).

Recent work on tin(II) halide systems has revealed that the selectivity of the benzhydrylation may be dependent on the concentration of the catalyst during the protection of diol substrates (Petursson, Scully, & Jonsdottir, 2014). Further work in tuning the primary-directed monoetherification of diol systems, such as 1,2- propanediol and closely related derivatives, may benefit from an exploration of catalyst concentration although the use of lower concentrations of catalysts may result in longer reaction times.

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Comparison of Diazo-[bis(4-methylphenyl)]methane and Diazo-[bis(4- methoxyphenyl)]methane for selective mono-etherification

Diazo-[bis(4-methylphenyl)]methane, which has previously demonstrated a high degree of regioselectivity in other vicinal diols in good yields, demonstrated a preference for the primary hydroxyl group leaving the secondary hydroxyl group available for the lipase-catalyzed acetylation for resolving the racemic diols. However, the selectivity was not complete for the primary hydroxyl group of the 1,2- diols under study. The resultant 1- and 2-ethers were not easily separated by column chromatography. This necessitated the removal of the unwanted 2-monoether by a reaction with a protecting group with a high selectivity for primary hydroxyl groups. In this case, the trityl group was chosen to separate the target 1-ether from the 2-ether to produce the quantities of 1-ether diols needed for subsequent lipase studies.

While the use of a trityl protecting group is selective for primary hydroxyl groups and the procedure itself is a highly facile, the trityl group itself is highly acid liable being deprotected by even mild, dilute acids. Additionally, this step necessitates the need for an extra separation prior to lipase catalyzed resolution.

The use of diazo-[bis(4-methoxyphenyl)]methane, due to its high reactivity, was also investigated. The use of this particular diazo compound, however, has been problematic and proved to be problematic in the protection of 1,2-diols due to the quantity of side products formed and the large excess of the diazo compound needed.

The role of the diol substrate’s structure may also be relevant in the explaining the differences of regioselectivity due to both steric and electronic features. The 3- phenoxy-1,2-propanediol has a bulky phenyl group which could limit the accessibility of the adjacent hydroxyl group on C2 thus making the primary hydroxyl on C1. Indeed, the ratio of 1-ether to 2-ether formation for the 3-phenoxy-1,2-propanediol reactions was higher than reactions under the same conditions for 1,2-propanediol. The electron withdrawing ability of the chloro group on 3-chloro-1,2-propanediol also seems to enhance the regioselectivity as the ratio of 1-ether to 2-ether formation was 10.2 to 1. This can be explained as the electronegativity of the chloro group decreases

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the nucleophilicity of the adjacent hydroxyl group on C2; the reaction time of the 3- chloro-1,2-propanediol was also longer than the other two diol substrates.

Use of diazo[bis(4-methoxyphenyl)]methane

Despite high-regioselectivity for the primary hydroxyl group, the presence of side products greatly complicated the subsequent separation. For this reason, the use of bis(4-dimethoxyphenyl)diazomethane was discontinued. In the case protecting 3- phenoyx-1,2-propanediol, a large excess of the diazo compound (6 equivalence) was required to drive the reaction to near completion. HPLC analysis revealed that traces of the starting material were present in roughly the same quantity (based upon peak area) after 4 and 6 total equivalence had been added. As with other reactions previously performed with this particular diazo compound, at least two characteristic by-products of yellow color were produced. While this side products were not characterized, previously performed characterization (Petursson, 1979) indicates that it is likely the corresponding azine and tetra(methoxyphenyl)ethane that forms as a result of an attack of one molecule of the diazo compound on another. Both side products complicated chromatographic efforts to separate the desired ether. The yield for this reaction was low and the separation was complicated by an impurity that co- eluted with the 1- and 2-ethers. Further reactions with using diazo[bis(4- methoxyphenyl)]methane were not performed.

4.3 Kinetic resolution of partially protected diols using lipases

The resolution of 1,2-propanediol derivatives is particularly challenging due to the small size of the R group (in this case a methyl group). Thus the challenge is to find a sufficiently large protecting group for the primary hydroxyl group and a selective enough lipase to accommodate only one enantiomer of 1,2-PD in such a configuration within the active site as to allow the acylation of the secondary hydroxyl group. Kazlaskas’ rule states that the acylation of the secondary hydroxyl group with the absolute (R) configuration is preferred over the (S) enantiomer. Five commercially

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available lipases immobilized on ceramic beads were investigated for their ability to resolve 1-O-benzhydryl-propan-2-ol as a model compound.

4.3.1 Comparison of lipases for 2(R) acetylation of rac-1[bis(4- methylphenyl)]methoxypropane-2-ol

The criteria for lipase selection for subsequent preparative scale resolutions was that the reaction did not consume more than 50% of the substrate, indicating that only one enantiomer is acylated, and a relatively short reaction time. According to Kazlauskas’ rule, the faster reacting alcohol will be the (R) enantiomer.

The lipases showed a variety of capability to resolve 1-ether protected 1,2-PD. The lipase from Pseudomonas fluorescens (PFL) and P. cepacia (PCL) seem capable of resolving enantiomers of 1-O-ether-propan-2-ol as approximately 50% of the substrate was consumed and the reaction did not proceed further over a period of days. Of the two enzymes, the PCL seemed to reach completion more quickly although this could be due to differences in the quantity of enzyme loadings on the ceramic beads or due to some of the enzyme present being inactive. This is supported as the stated enzyme activity for each of these two lipases are 15000 U/mg and >40 U/mg, respectively. The chromatographs for the PFL catalyzed reaction did, however, show some shouldering perhaps indicating a side reaction taking place. For these reason, PCL was chosen for further study.

The attempted resolution using Candida cylindracea lipase (CCL) showed a very low rate of reaction with the substrate with some traces of product appearing after a reaction time of 8 days. The other two lipases isolated from fungi, C. antarctica (CAL) and M. mehei (MML), were not suitable for the resolution of the 1,2-PD derivative as evidenced by the consumption of the racemic starting material proceeding past a 1:1 ratio of starting material to product. This indicates that these lipases are not stereoselective towards the substrate. Perhaps this is not surprising as fungal hydrolases are often more promiscuous than their bacterial counterparts and that these two organisms are psychophiles which typically have accommodating

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active sites to counter the thermodynamic challenges of operating at lower temperatures (Georlette et al., 2003).

4.4.2 Chemoenzymatic resolution and deprotection of rac-1[bis(4- methylphenyl)]methoxy derivatives of 1,2-propanediol

The resolution of the racemic 1-ether derivatives of 1,2-propanediol, 3-phenoxy-1,2- propanediol, and 3-chloropropanediol and the subsequent deprotections of the (2R)- acetyl derivatives gave good yields. The 3-chloro and 3-phenoxy substrates reacted more slowly than the 1-ether of 1,2-propanediol likely for electronic and steric reasons, respectively; this could have been minimized by using more of the immobilized lipase which can be filtered off and reused. The PCL-catalyzed reactions of each of the racemic 1-ethers only proceeded to approximately 50% which is highly indicative that the reaction is selective for one enantiomer. This could be further confirmed by measuring the optical rotations of the resolved partially protected diols.

An attempt was made to measure the optical rotation of the resolved R and S ethers however the rotations were so small that they could not be accurately measured with a path length of 10 cm or 1 cm. The resolved compounds, were, however, of roughly equal magnitude but rotated light in the opposite directions. A preferable method to determine the rotation and subsequent enantiomeric excess would have involved using an automated polarimeter or examining 1H NMR after derivatization with a chiral reagent. Alternately, the use of a chiral HPLC column (such as α-cyclodextrin) could have determined the enantiomeric purity.

The enzymatic resolution of 1,2-propanediol and closely related compounds has been achieved in the literature with good enantiomeric excess although the reported yields are low. Oxidative and reductive resolutions using organisms such as Saccharomyces cerevisiae (Baker’s yeast) have also been reported as mentioned in Section 1.2.3. However, the resultant enantiomeric excess values are low as are yields; the low enantiomeric purity would thus necessitate multiple rounds of yeast mediated resolution. The resolution of racemic compounds in this manner may be acceptable for highly inexpensive commodity chemicals but may not be suitable for highly sensitive compounds or expensive substrates. Another potential pitfall of this method

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is that only one enantiomer can be isolated meaning that another route to the other enantiomer must be used.

This work demonstrates that the use of a benzhydryl protecting group to block the reactivity of the primary hydroxyl group followed by the protection of one enantiomer with an acyl group is possible. While the overall yields are somewhat lower due to the incomplete regioselectivity during the tin(II) halide catalyzed protection of the primary hydroxyl group, both enantiomers can be obtained in reasonable yield and enantiomeric purity.

4.4 Future directions

Recent experimental evidence has indicated that the concentration of the tin(II) halide catalyst may play a role in the regioselectivity of reactions with vicinal diol systems of partially protected hexoses (Petursson et al., 2014). The “tune-ability” of 1,2-diol systems requires exploration in the hopes of improving the regioselectivity in order to decrease or negate the need for the removal of undesirable 2-ethers.

An alternative method for preparing bis(4-methoxyphenyl)methyl ethers from the corresponding alcohol in good yields using copper(II) bromide was reported by (Mezaache et al., 2009) although the regioselectivity for diol systems was not explored. The use of copper(II) halides could present an alternative to tin(II) halide systems should they be found to catalyze the introduction of benzhydryl ethers.

The production of diazo compounds using heavy metal oxides, such as mercury(II) oxide, is a somewhat problematic procedure due to the disposal of stoichiometric quantities of mercury residue. New methods for the generation of diazo compounds, such as the use of chromium(II) oxide (Magtrieve™) allow for the rapid generation of diazo compounds using large excesses of the metal which can be magnetically retrieved(Lee & Donald, 1997). This method could also potentially be used for the one-pot protection of alcohols as the protection of carboxylic acids using diazo compounds using this method has already been reported (Wan & Peng, 2008).

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All of the commercial lipases explored were of low temperature or mesophilic origin. It has been noted that lipases of thermophilic origin have more structurally rigid active sites. As such, narrower active sites may be more selective for 1,2-diol systems using smaller protecting groups on the primary hydroxyl group.

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5 Conclusions

The use of tin(II) halide catalyzed alkylations for the protection of primary alcohols for three vicinal 1,2-diols (1,2-propanediol, 3-chloro-1,2-propaneidol, and 3-phenoxy- 1,2-propanediol) undergoing lipase-catalyzed kinetic resolutions presents a novel strategy with some advantages over techniques reported in the literature to data. Further optimization of the catalyst system is needed to improve selectivity for the primary alcohol position of the vicinal 1,2-diol system. Incomplete selectivity for the primary hydroxyl group, however, required the use of another reaction and separation to remove the unwanted 2-ether. The subsequent kinetic resolution of the racemic mixture to the acylated R enantiomer and free S enantiomer of the 1-ether using Pseudomonas cepacia lipase showed the benzhydryl ether derivative to be highly selective for the R enantiomer.

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Appendix A Kinetic Data

Figure 30 – PCL-catalyzed reaction of 1-bis(4-methylphenyl)methyl-propan-2-ol with vinyl acetate

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Figure 31 -PFL-catalyzed reaction of 1-bis(4-methylphenyl)methyl-propan-2-ol with vinyl acetate

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Figure 32 – MML-catalyzed reaction of 1-bis(4-methylphenyl)methyl-propan-2-ol with vinyl acetate

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Figure 33 – CCL-catalyzed reaction of 1-bis(4-methylphenyl)methyl-propan-2-ol with vinyl acetate

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Figure 34 – CAL-catalyzed reaction of 1-bis(4-methylphenyl)methyl-propan-2-ol with vinyl acetate

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Appendix B 1H and 13C NMR Spectra rac-1[bis(4-methylphenyl)]methoxypropane-2-ol

Chemical Formula: C18H22O2 Molecular Weight: 270,37

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(2R)-1[bis(4-methylphenyl)]methoxy-2-acetoxypropane

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1-O-bis(4-methoxyphenyl)methyl-propan-2-ol

OMe OH O PhO

OMe

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3-Phenoxy-1-O-bis(4-methoxyphenyl)methyl-propan-2-ol

PhO O OH

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(2R)-3-phenoxy-1[bis(4-methylphenyl)]methoxy 2-acetoxypropane

PhO O OAc

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3-phenoxy-1[bis(4-methylphenyl)]methoxy 2-acetoxypropane

PhO O OH

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1-O-bis(4-methylphenyl)-3-chloro-propan-2-ol