CHAPTER 5

Bristol-Myers Squibb: Preparation of Chiral Intermediates for the Development of Drugs and APIs

RAMESH N. PATEL

SLRP Associates,LLC, Consultation in Biotechnology, 572 Cabot Hill Road, Bridgewater, NJ 08807 USA

*E-mail: rameshpatelphd@yahoo.com

5.1 Introduction

For the development of pharmaceuticals, agrochemicals, and nutraceuticals the synthesis of single enantiomers of products has become increasingly important, since the undesired enantiomer is either biologically inactive or even toxic in nature. There is an enormous potential of microorganisms and biocatalysts derived from microbial cultures for carrying out the conversion of various synthetic chemicals into desired products in highly chemo-selective, enantioselective and regioselective manner and in excellent yields. Such high selectivity offers advantages in pharmaceutical synthesis, such as minimizing side-reactions and eliminating the need of protection–deprotection steps, leading to shorter syntheses. Biocatalysis provides a technology that uses milder and environmentally safer processes, operated under ambient temperature, atmospheric pressure and often in aqueous media. Biocatalysis generally enables more sustainable routes to key intermediates and active pharmaceutical ingredients, effectively reducing the waste production.

During the last decade, progress in high level expression of enzymes in Escherichia coli, Pichia pastoris, and other microbial systems, and improvement in fermentation technology, has led to increased cell yields in shorter times. Advances in protein purification technology, determination of the structure of a protein along with molecular cloning, random and directed evolution biocatalysts has opened up unlimited access to a variety of enzymes and microbial cultures as tools in organic synthesis. In the last ten years, the development of directed evolution of biocatalysts under process conditions has led to increased enzyme activities, and improved selectivity and stability of biocatalysts. Several comprehensive review articles and book chapters have been published in this area.^1–161-¹6 Moreover, the development of efficient immobilization techniques for biocatalysts and their reusability have provided highly economical and energy efficient processes for the synthesis of key intermediates and drug products in pharmaceutical, agrochemicals, flavors, cosmetics, and pesticides industries. We at Bristol-Myers Squibb started a Multidisciplinary Biocatalysis Group in 1987 to support our process research and development department to access key chiral intermediates for the development of drugs by alternative green technology. This chapter provides an overview of our activity, and discusses selected examples for the synthesis of key intermediates and drug products via biocatalytic routes.

5.2 Anti-Alzheimer's Drug. Enzymatic Preparation of (R)-5,5,5-Trifluoronorvaline

(R)-5,5,5-Trifluoronorvaline (1, Figure 5.1) is an intermediate for a γ-secretase inhibitor 2 under development by Bristol-Myers Squibb for treatment of Alzheimer’s disease. Amyloid-β peptides (Aβ) are a major component of the plaques that are found in the brains of Alzheimer’s disease patients and have been proposed to play a causative role in the disease.^17–19 These peptides are produced from amyloid precursor protein by initial cleavage at the β-site by β-secretase followed by further cleavage at sites near the C-terminus by γ-secretase to generate Aβ forms, with Aβ42 most closely associated with Alzheimer’s disease. Inhibition of γ-secretase has been pursued as a pharmaceutical target to decrease the formation of Aβ peptides. In this field, BMS-708163, a γ-secretase inhibitor, causes a significant decrease of Aβ40 levels.^17–19

Figure 5.1 Anti-Alzheimer's drug: enzymatic preparation of (R)-5,5,5-trifluoronorvaline (1).

(R)-5,5,5-Trifluoronorvaline (1) was prepared from the corresponding keto acid 3 by reductive amination reaction using a commercially available (R)-amino acid dehydrogenase (provided by Biocatalytics, Inc.), coupled with a glucose dehydrogenase used for cofactor NADPH regeneration and recycling.^20,21 Amino acid 1 could also be prepared using a (R)-amino acid transaminase with alanine as the amino donor, but the transamination reaction also required lactate dehydrogenase, NAD⁺, formate, and formate dehydrogenase to remove pyruvate generated as by-product (to bring the reaction to completion).²¹ Thus, the reductive amination process was used for further development.

A proprietary (R)-amino acid dehydrogenase was prepared at Bristol-Myers Squibb by modification of the (R)-diaminopimelic acid dehydrogenase gene from Bacillus sphaericus and cloned in Escherichia coli. A glucose dehydrogenase gene was also cloned from Gluconobacter oxidans to regenerate and recycle cofactor NADH. Both genes were expressed in the same strain of Escherichia coli. In addition, to improve the enantiomeric excess (ee) of the product to 100%, the glutamate dehydrogenase gene was inactivated in the expression strain, to eliminate the background production of the (S)-amino acid from keto acid 3.^20,21

In the enzymatic reductive amination process, the reaction mixture in 1 L of water contained 26.7 g NH₄Cl, 62.5 g glucose, and 50 g keto acid 3. The pH of the reaction was adjusted to 9.0 followed by addition of 382 mg NADP⁺ and the extract of recombinant Escherichia coli containing 1250 units of recombinant (R)-amino acid dehydrogenase and 7500 units of glucose dehydrogenase. After 22 h reaction, the amino acid 1 was produced in 88.5% yield and 98.9% ee. The process was further scaled-up for production of intermediate 1 on a 50 kg level.²¹ The amino acid could be either isolated, or converted without isolation into the p-chlorophenylsulfonamide carboxamide 4 intermediate needed for the synthetic route to the γ-secretase inhibitor 2.

5.3 Cholesterol Lowering Agents

5.3.1 Enantioselective Enzymatic Acylation of Racemic Alcohol

Nowadays, hydroxymethyl glutarate coenzyme A (HMG-CoA) reductase inhibitors (statins) have worldwide sales of approximately $20 billion, led by Atorvastatin (5) and Crestor (6) (rosuvastatin) (Figure 5.2). All the synthetic statins share the chiral 3,5-dihydroxy acid side chain, which is essential for activity and represents the main synthetic challenge for the preparation of these drugs.²²

Figure 5.2 Cholesterol lowering agents: enantioselective enzymatic acylation to prepare (5R,3R)-alcohol, [4-[4α,6β(E)]]-6-[4,4-bis[4-fluorophenyl]-3-(1methyl-1H-tetrazol-5-yl)-1,3-butadienyl] tetrahydro-4-hydroxy-2H-pyren-2-one (7).

The (5R,3R)-alcohol 7, [4-[4α,6β(E)]]-6-[4,4-bis[4-fluorophenyl]-3-(1methyl-1H-tetrazol-5-yl)-1,3-butadienyl]tetrahydro-4-hydroxy-2H-pyren-2-one is a potential new anti-cholesterol drug which acts by inhibition of HMG CoA reductase.²³ An enzymatic diastereoselective acetylation process was developed for the resolution and synthesis of (5R,3R)-alcohol 7 from racemic alcohol (±)-8, using lipase PS-30 (lipase from Pseudomonas cepacia commercially available from Amano Enzyme company, Japan).²⁴ Lipase PS-30 efficiently catalyzed the enantioselective acetylation of racemic 8 (4 g L⁻¹) to yield the (5R,3S)-acetate 9 and the unreacted, desired (5R,3R)-7. Yields of 49% with enantiomeric excess of 98.5% were obtained for (5R,3R)-7 when the reaction was conducted in toluene in the presence of isopropenyl acetate as acyl donor. In methyl ethyl ketone at 50 g L⁻¹ substrate concentration, yields of 46% and enantiomeric excess of 98% were obtained for the (5R,3R)-alcohol 7.²⁴

The enzymatic process was scaled-up to a 640 L preparative batch using immobilized lipase PS-30 (lipase PS-30 immobilized on Accurel polypropylene). The substrate input was 4 g L⁻¹, with 0.05% water, isopropenyl acetate (as the acyl donor), and toluene as solvent. The reaction was carried out at 37 °C for 24 h. The enzyme was then filtered off with Niagara plate filter and reused for the next cycle. The toluene filtrate was extracted with methanol : water (50 : 50 v/v). The methanol content of the recovered extract was reduced to 20% in volume and the entire contents were extracted with ethyl acetate. The ethyl acetate layer was collected and concentrated under reduced pressure and the product was crystallized from heptane to afford the (5R,3R)-7 in 45% overall yield with 98.5% ee and 99.5% chemical purity. The (5R,3S)-acetate 9 obtained by this process was enzymatically hydrolyzed by lipase PS-30 in a biphasic system to prepare the corresponding (5R,3S)-alcohol 10 in 40% yield and 98% ee.²⁴

5.3.2 Enzymatic Synthesis of (3S,5R)-Dihydroxy-6-(benzyloxy)hexanoic Acid, Ethyl Ester

In the previous section the preparation of cholesterol lowering agent 7 by a kinetic resolution process was described, providing access to both diastereomers of alcohols 7 and 10 in 45% and 40% yields, respectively. To improve the process and obtain higher yields, the enantioselective enzymatic reduction of the diketone 3,5-dioxo-6-(benzyloxy)hexanoic acid, ethyl ester (11) to (3S,5R)-dihydroxy-6-(benzyloxy)hexanoic acid, ethyl ester (12, Figure 5.3) was demonstrated by cell suspensions of Acinetobacter sp. SC 13876. Compound 12 is a key chiral intermediate required for the chemical synthesis of 7, Lipitor (5) and Crestor (6), all of which are cholesterol lowering agents that act by inhibition of HMG CoA reductase.^25–27

Figure 5.3 Cholesterol lowering agents: Enzymatic synthesis of (3S,5R)-dihydroxy-6-(benzyloxy)hexanoic acid, ethyl ester (12).

Cell suspensions of Acinetobacter sp. SC 13876 reduced the ethyl diketoester 11 to a mixture of desired syn and undesired anti diastereomers. The desired syn-(3R,5S)-dihydroxy ester 12 was obtained in 99% ee and 63% diastereomeric excess (de).^26,27 Likewise, the cell suspensions reduced the t-butyl diketoester 13 to a mixture of mono- and dihydroxy esters with desired syn (3R,5S)-dihydroxy ester 14 in 87% ee and 51% de.^26,27 Cell extracts of A. calcoaceticus SC 13876 in the presence of NAD⁺, glucose, and glucose dehydrogenase reduced 11 to the corresponding monohydroxy compounds [5-hydroxy-3-oxo-6-(benzyloxy)hexanoic acid ethyl ester (15) and 3-hydroxy-5-oxo-6-(benzyloxy)hexanoic acid ethyl ester (16)]. Both 15 and 16 were further reduced to the (3S,5R)-dihydroxy compound 12 in 92% yield and 99% ee. Subsequently, (3S,5R)-dihydroxy compound 12 was converted into 17, a key chiral intermediate for the synthesis of 7, Lipitor (5) and Crestor (6).^26,27

Three different ketoreductases were identified and purified to homogeneity from cell extracts of Acinetobacter sp. SC 13876, and their biochemical properties were compared. Reductase I only catalyzes the reduction of ethyl diketoester 11 to its monohydroxy products 15 and 16, whereas reductase II catalyzes the formation of dihydroxy product 12 from monohydroxy substrates 15 and 16. A third reductase (III) was identified, catalyzing the reduction of diketoester 11 directly to the desired syn-(3R,5S)-dihydroxy ester 12.²⁷ Ketoreductase III was cloned and expressed in E. coli. A significant novel protein of the expected molecular weight was visible in soluble fractions showing superior expression, and the ketoreductase III represented nearly half of the total protein as judged by SDS-PAGE.²⁶ Cells and soluble extracts of recombinant culture expressing ketoreductase III gave complete reduction of the diketoester 11 directly to the desired syn-(3R,5S)-dihydroxy ester 12 with 99.3% yield, 99.9% ee and 99.8% de in 9 h reaction with both cell suspensions and soluble extracts at 50 g L⁻¹ substrate input.²⁸

5.4 Calcitonin Gene-related Peptide Receptors Antagonists (Migraine Treatment): Enzymatic Preparation of (R)-2-Amino-3-(7-methyl-1H-indazol-5-yl)propanoic Acid

(R)-2-Amino-3-(7-methyl-1H-indazol-5-yl)propanoic acid ((R)-18, Figure 5.4) is a key intermediate needed for the synthesis of antagonists of calcitonin gene-related peptide receptors (CGRP receptors) 19.²⁹ Such antagonists are potentially useful for the treatment of episodic and chronic migraine and other pain maladies.^29–31 We initially developed an enzymatic process for the conversion of the racemic amino acid 20 into (R)-amino acid 18 using (S)-amino acid deaminase (also known as (S)-amino acid oxidase) from Proteus mirabilis cloned and expressed in E. coli in combination with commercially available (R)-transaminase (Biocatalytics Inc.) and alanine as the amino donor.³² The process was optimized at 500 mL scale using racemic 20 (20 g), racemic alanine (40 g), pyridoxal phosphate monohydrate (2.65 mg), 100 g of E. coli frozen cell paste containing the cloned (S)-amino acid deaminase (27 U g⁻¹ cells) from P. mirabilis, and (R)-transaminase (Biocatalytics Inc., 200 mg). Yields of 85% with 96.5% ee were obtained for (R)-amino acid 18. The enzymatic process was further scaled-up to pilot plant for the conversion of 2.9 kg 20 into (R)-amino acid 18 in two batches. The average yield of the isolated product 18 was 1.79 kg with 61% overall yield and 98.6% ee.³²

Figure 5.4 Calcitonin gene-related peptide receptor antagonists (migraine treatment): enzymatic preparation of (R)-2-amino-3-(7-methyl-1H-indazol-5-yl)propanoic acid (18).

Upon availability of the more soluble ketoacid 21, (R)-amino acid 18 was prepared directly from ketoacid using the commercially available (R)-transaminase (Biocatalytics Inc.) without the necessity of (S)-amino acid deaminase required for the oxidation step. Starting the process from the ketoacid instead of from the racemic amino acid resulted in a more efficient process, but the reaction was still slow to reach completion, apparently due to inhibition by co-product pyruvate generated from alanine used as an amino donor in the transamination process. The average conversion after 70 h was 89%, and the average isolated yield was 76% with >99% ee for (R)-amino acid 18. As supposed above, pyruvate was confirmed to be a strong product inhibitor of the reaction. Addition of L-lactate dehydrogenase from rabbit muscle in the presence of L-lactate, formate dehydrogenase, NAD⁺, and sodium formate (required for cofactor regeneration) brought the reaction to near completion in as little as 4 h.³² However, this process required three enzymes as well as cofactor NAD⁺.

To improve the process further, the isolation, purification, cloning and overexpression of a (R)-transaminase – from a soil isolate identified as Bacillus thuringiensis – was carried out for the conversion of keto acid 21 into (R)-amino acid 18. In the enzymatic process using crude extract, our cloned and expressed (R)-transaminase in Escherichia coli SC 16557 was highly effective, outperforming the commercially available (R)-transaminase (Biocatalytics Inc.). Reaction was completed without any removal of pyruvate as the transaminase expressed in E. coli SC16557 was less sensitive to inhibition by pyruvate than the commercially available enzyme. Thus, the new process gave (R)-amino acid 18 in a reaction yield of 92.4% and ee of 99.6%. The process was scaled-up to prepare multi-kilos of (R)-amino acid 18 for synthesis CGRP antagonists.³²

5.5 Antidiabetic Drugs

5.5.1 Saxagliptin: Enzymatic Synthesis of (S)-N-Boc-3-hydroxyadamantylglycine

The synthesis of saxagliptin (22), a dipeptide peptidase-4 inhibitor (DPP-4) developed by Bristol-Myers Squibb, required (S)-N-Boc-3-hydroxyadamantylglycine (23) as an intermediate (Figure 5.5a). DPP-4 is a ubiquitous proline-specific serine protease responsible for the rapid inactivation of incretins, including glucagon-like peptide 1 (GLP-1), and glucose-dependent insulinotropic peptide. To alleviate the inactivation of GLP-1, inhibitors of DPP-4 are being evaluated for their ability to provide improved control of blood glucose for diabetics.^33–37

Figure 5.5 Antidiabetic drug, saxagliptin: (a) enzymatic synthesis of (S)-N-boc-3-hydroxyadamantylglycine (23); (b) enzymatic ammonolysis of (5S)-4,5-dihydro-1H-pyrrole-1,5-dicarboxylic acid, 1-(1,1-dimethylethyl)-5-ethyl ester (34).

An enzymatic process for the conversion of the keto acid 24 into the corresponding amino acid 25 was developed using (S)-amino acid dehydrogenases. Among various amino acid dehydrogenases tested for reductive amination of 24, a modified form of a recombinant phenylalanine dehydrogenase was found to be the best. The modified phenylalanine dehydrogenase contains two amino acid changes at the C-terminus and a 12 amino acid extension of the C-terminus.³⁸ Phenylalanine dehydrogenase from Thermoactinomyces intermedius was cloned in the yeast Pichia pastoris as well as in bacterial host Escherichia coli. Both enzymes were used for this process. Cofactor NAD⁺ was consumed in this process, producing NADH during the reaction that was recycled using formate dehydrogenase. The production of multi-kg batches was initially carried out with extracts of Pichia pastoris expressing the modified phenylalanine dehydrogenase from Thermoactinomyces intermedius, and endogenous formate dehydrogenase produced during growth of Pichia pastoris on methanol as carbon source. Methanol also served as an inducer for production of recombinant phenylalanine dehydrogenase.

The reductive amination process was further improved and scaled up using a preparation of the two enzymes (phenylalanine dehydrogenase and formate dehydrogenase) expressed in a single recombinant Escherichia coli. Thus single fermentation process was required for generation of both enzymes in shorter time with high activity. The amino acid 25 was directly protected as its Boc derivative without isolation to afford intermediate 23. Yields before isolation were close to 98% with 100% ee.³⁸ Likewise, cell extracts from E. coli strain SC16496 expressing modified phenylalanine dehydrogenase and cloned formate dehydrogenase were used – after clarification and concentration – to complete the reaction in 30 h with >96% yield and >99.9% ee of product 25. This process has now been used to prepare several hundred kg of Boc-protected amino acid 23 for the development of saxagliptin.³⁸

In the enzymatic process, cells of recombinant E. coli (25% w/v) in 50 mM ammonium formate pH 7 were suspended using a homogenizer, and disrupted by two passages through a microfluidizer at 12 000 psi at 4 °C. Cell debris was removed by centrifugation. The supernatant solution (266 mL) containing 2456 Units phenylalanine dehydrogenase and 8801 Units formate dehydrogenase was added to a 1 L flask. Another solution (266 mL) was prepared, containing ammonium formate (16.74 g) and keto acid 24 (29.76 g), and adjusted to pH 8.0 using concentrated ammonium hydroxide, and then NAD⁺ (372 mg) and dithiothreitol (81.8 mg) were added as well. Subsequently the prepared solution was added to the flask containing the E. coli extract. The reaction was carried out at 40 °C on a shaker at 40 rpm and pH 8.0 for 30 h. A yield of 100% and an ee of 99.9% were obtained for amino acid 25. This product was converted directly into N-Boc-amino acid 23 and isolated in overall yield of 88% and 99.9% ee.³⁸

5.5.2 Saxagliptin: Enzymatic Synthesis of N-Cbz-4,5-dehydro-L-prolineamide and N-Boc-4,5-dehydro-L-prolineamide

As stated in the previous section, saxagliptin (22) is a dipeptide peptidase-4 inhibitor (DPP-4) developed at Bristol-Myers Squibb. In another synthetic strategy, the intermediate N-Cbz-4,5-dehydro-L-prolineamide (26) or N-Boc-4,5-dehydro-L-prolineamide (27, Figure 5.6) were required for the synthesis of proline fragment 28. To this end, we developed an enzymatic process for the conversion of N-α-Cbz-L-ornithine (29) and N-α-Boc-L-ornithine (30) into N-Cbz-5-hydroxy-L-proline (31) and N-Boc-5-hydroxy-L-proline (32), respectively (Figure 5.6a). The reaction required α-ketoglutarate. The glutamate generated during this reaction was regenerated back to α-ketoglutarate by glutamate oxidase of Streptomyces sp. (cloned and expressed in Escherichia coli). The reaction mixture contained 1.33 g of N-α-Cbz-L-ornithine, 1.9 g of α-ketoglutarate disodium salt and 50 mL 0.1 M potassium phosphate buffer, pH 8, in a total volume of 250 mL. After the solids were dissolved, 0.5g of frozen E. coli cells BL21-DE3 (pBMS2000-LAT) expressing lysine-ɛ-aminotransferase from Sphingomonas paucimobilis was added and the reaction was carried out at 28 °C, 200 rpm at pH 8. After 39 h of biotransformation process, the product N-Cbz-5-hydroxy-L-proline (31) was isolated in 92% yield.³⁹ Subsequently 31 was converted chemically into N-Cbz-4,5-dehydro-L-prolineamide 26 (Figure 5.6b). Similarly, N-α-Boc-L-ornithine (30) was converted into N-Boc-5-hydroxy-L-proline (32) using the recombinant lysine-ɛ-amino transferase in 91% yield. Compound 32 was chemically converted into N-Boc-4,5-dehydro-L-prolineamide (27). To further improve the process, an amine oxidase from Pichia sp. SC 16539 was cloned and expressed in Pichia pastoris. These recombinant cells were used to convert L-ornithine (33), N-α-Cbz-L-ornithine (28) and N-α-Boc-L-ornithine (29) into N-5-hydroxy-L-proline, N-Cbz-5-hydroxy-L-proline (31) and N-Boc-5-hydroxy-L-proline (32, Figure 5.6a), respectively, in >98% yield. Moreover, α-ketoglutarate required for transamination reaction by lysine-ɛ-aminotransferase was not required for the oxidation reaction carried out by L-ornithine oxidase.³⁹

Figure 5.6 Saxagliptin: (a) enzymatic synthesis of N-Cbz-4,5-dehydro-L-prolineamide (26) and N-Boc-4,5-dehydro-L-prolineamide (27); (b) synthesis of N-Cbz-4,5-dehydro-L-prolineamide and N-Boc-4,5-dehydro-L-prolineamide from N-Cbz-5-hydroxy-L-proline (31) and N-α-Boc-5-hydroxy-L-proline (32), respectively.

5.5.3 Saxagliptin: Enzymatic Ammonolysis of (5S)-4,5-Dihydro-1H-pyrrole-1,5-dicarboxylic Acid, 1-(1,1-Dimethylethyl)-5-ethyl Ester

The synthesis of saxagliptin by a different approach required (5S)-aminocarbonyl-4,5-dihydro-1H-pyrrole-1-carboxylic acid 1-(1,1-dimethylethyl) ester (34, Figure 5.5b).^33–37 Direct chemical ammonolyses were hindered by the requirement of aggressive reaction conditions, which resulted in unacceptable levels of amide racemization and side-product formation. On the other hand, milder two-step hydrolysis–condensation protocols using coupling agents such as 4-(4,6-dimethoxy-1,3,5-triazin-2-yl)-4-methylmorpholinium chloride (DMTMM) were compromised by reduced overall yields.⁴⁰ To address this issue, a biocatalytic process was developed, based on Candida antarctica lipase B (CALB)-mediated ammonolysis of (5S)-4,5-dihydro-1H-pyrrole-1,5-dicarboxylic acid-1-(1,1-dimethylethyl)-5-ethyl ester (35) with ammonium carbamate to furnish 34 without racemization and with low levels of side-product formation.⁴¹ Experiments utilized process stream ester feed, which consisted of ca. 22% w/v (0.91 M) of the ester in toluene. Since the use of toluene precluded the use of free ammonia due to its low solubility in toluene, solid ammonium carbamate was employed. Reactions were performed using a mixture of neat process feed, ammonium carbamate (71 g L⁻¹) and biocatalyst (25 g L⁻¹), shaken at 400 rpm, 50 °C. Under these conditions, CALB provided racemization-free amide with yields of 69%, together with 21% of side-products. The inclusion of various additives was investigated to solve potential inhibitory phenomena, shifting the equilibrium towards amide synthesis and reducing side-product formation. Thus, drying agents such as calcium chloride led to significant improvements (79% amide and 13% side-products). Calcium chloride is known to complex alcohols as well as act as a desiccant, and it is presumed to act on the binding of ethanol released during the course of amide formation, thus mitigating deleterious effects of this alcohol on CALB catalysis. Likewise, a dramatic increase in amide yield to 84% and 95% was achieved by including soda lime and Ascarite, respectively, at 200 g L⁻¹ in the reaction headspace. This effect was presumably due to the adsorption of carbon dioxide liberated from the decomposition of ammonium carbamate. A further increase in yield to 98% was attained via the combined use of 100 g L⁻¹ of calcium chloride and 200 g L⁻¹ of Ascarite. A preparative–scale reaction with the process ester feed was then conducted. Ester (220 g L⁻¹) was reacted with 90 g L⁻¹ of ammonium carbamate, 33 g L⁻¹ of CALB, 110 g L⁻¹ calcium chloride and 216 g L⁻¹ of Ascarite (in the headspace), run at 50 °C for 3 days. Complete conversion of ester was achieved, with the formation of 96% (182 g L⁻¹) of amide and 4% of side-products. After work-up 98% potency amide of >99.9% ee was isolated in 81% yield.⁴¹

5.5.4 GLP-1 Receptor Agonists: Enzymatic Preparation of (S)-Amino-3-[3-{6-(2-methylphenyl)}pyridyl]-propionic Acid

Glucagon-like peptide 1 (GLP-1a) is a gastrointestinal hormone that exists predominantly as a 30 amino acid, C-terminally amidated peptide (GLP-1(7-36)-NH₂), or as the equally active, glycine-extended form, GLP-1(7-37). Both peptides act as a functional agonist of the GLP-1 receptor. The importance of GLP-1 and its receptor in regulating glucose homeostasis and food intake supports the view that GLP-1 receptor agonists have the potential to improve on the current standard of care in treating type 2 diabetes.⁴² Mapelli et al.^43,44 discovered a novel class of eleven amino acid GLP-1 receptor agonists. These peptides consist of a structurally optimized nine-mer, which is closely related to the N-terminal nine amino acids of GLP-1, linked to a substituted C-terminal biphenylalanine (BIP) dipeptide. Further studies demonstrated that peptide reduced plasma glucose excursions and increased plasma insulin in a mouse model of diabetes.^42–44 The (S)-amino-3-[3-{6-(2-methylphenyl)}pyridyl]propionic acid (36, Figure 5.7a) is a key intermediate required for synthesis of GLP-1 mimics 11-mer peptide or GLP-1 receptor modulators useful for the treatment of type II diabetes treatment.^42–44 (S)-Amino-3-[3-{6-(2-methylphenyl)}pyridyl]propionic acid was prepared by an enzymatic deracemization process in 72% isolated yield with >99.4% ee from racemic amino acid 37, using a combination of two enzymes, namely an (R)-amino acid oxidase from Trigonopsis variabilis, and an (S)-aminotransferase from Sporosarcina ureae, both enzymes cloned and expressed in Escherichia coli. (S)-Aspartate was used as amino donor.⁴⁵ Another (S)-aminotransferase was also purified from a soil organism identified as Burkholderia sp. and cloned and expressed in Escherichia coli and used in this process as well. In the enzymatic process, racemic amino acid 37 was first treated with (R)-amino acid oxidase for 4 h to convert racemic amino acid 37 into a mixture of (S)-amino acid 36 and keto acid 38. Subsequently in the same reaction mixture (S)-aminotransferase was charged to convert keto acid 38 into (S)-amino acid 36, to render 85% final yield. The process was scaled up to 70 L volume at a substrate input of 1.5 kg. From the reaction mixture product 36 was isolated in 70% yield and 99.9% ee.⁴⁵ In an alternative improved process (Figure 5.7b), the enzymatic dynamic kinetic resolution of racemic amino acid 37 was also demonstrated. (R)-Selective oxidation with Celite-immobilized (R)-amino acid oxidase from Trigonopsis variabilis expressed in E. coli in combination with chemical imine 39 reduction (with borane-ammonia) gave a 68% isolated yield and >99.99% ee of (S)-amino acid 36.⁴⁵

Figure 5.7 GLP-1 receptor agonists: enzymatic preparation of (S)-amino-3-[3-{6-(2-methylphenyl)}pyridyl]-propionic acid (36).

5.6 Antihypertensive Drugs

5.6.1 Enzymatic Synthesis of (S)-6-Hydroxynorleucine

Vanlev (40, Figure 5.8a) is an antihypertensive drug which acts by inhibiting both angiotensin converting enzyme (ACE) and neutral endopeptidase enzyme (NEP).⁴⁶ (S)-6-Hydroxynorleucine 41 is a key intermediate in the synthesis of Vanlev. The synthesis and complete conversion of 2-keto-6-hydroxyhexanoic acid (42) into (S)-41 was developed by enzymatic reductive amination using beef liver glutamate dehydrogenase.⁴⁷ As depicted (Figure 5.8a), compound 42 – in equilibrium with 2-hydroxytetrahydropyran-2-carboxylic acid sodium salt (43) – was converted biocatalytically into (S)-41. The reaction requires ammonia and NADH. The NAD⁺ produced during the reaction was recycled to NADH by the oxidation of glucose to gluconic acid using glucose dehydrogenase from Bacillus megaterium. The reaction was completed in about 3 h at 100 g L⁻¹ substrate input with reaction yields of 92% and ee of 99.8% for (S)-6-hydroxynorleucine.

Figure 5.8 Antihypertensive drug, Vanlev: enzymatic synthesis of (S)-6-hydroxynorleucine ((S)-41).

In a further approach leading to a more convenient process, the keto acid was prepared by treatment of racemic 6-hydroxynorleucine (44) (produced by hydrolysis of 5-(4-hydroxybutyl)hydantoin 45) with (R)-amino acid oxidase (Figure 5.8b). Once the % ee of the unreacted (S)-6-hydroxy norleucine had reached 99.8%, the reductive amination procedure was used to produce (S)-6-hydroxynorleucine (41) in 97% yield with 99.8% ee from racemic 6-hydroxynorleucine at 100 g L⁻¹ substrate concentration.⁴⁷ The (S)-6-hydroxynorleucine prepared by the enzymatic process was converted chemically into Vanlev.⁴⁸

5.6.2 Vanlev: Enzymatic Synthesis of Allysine Ethylene Acetal

To improve the process for the synthesis of Vanlev described in the previous section, a subsequent strategy used better intermediates, namely (S)-2-amino-5-(1,3-dioxolan-2-yl)pentanoic acid (46) [(S)-allysine ethylene acetal] (Figure 5.9a). An enzymatic process for the synthesis of (S)-46 was developed by reductive amination of keto acid acetal 47 using phenylalanine dehydrogenase (PDH) from Thermoactinomyces intermedius.⁴⁹ The reaction required ammonia and NADH, and produced NAD⁺ was regenerated by the oxidation of formate to CO₂ using formate dehydrogenase (FDH). T. intermedius PDH was cloned and expressed in E. coli and recombinant culture was used as a source of PDH. P. pastoris grown on methanol was used as sources of FDH. Expression of T. intermedius PDH in P. pastoris (inducible by methanol) allowed the generation of both enzymes (PDH and FDH) in a single fermentation process. This process was scaled-up to prepare 197 kg of (S)-46 with 91% yield and >98% ee.⁴⁴ (S)-Allysine ethylene acetal prepared by enzymatic process was subsequently converted into Vanlev.^48,49

Figure 5.9 Antihypertensive drug, Vanlev: (a) enzymatic synthesis of allysine ethylene acetal 46; (b) enzymatic synthesis of thiazepine 48.

5.6.3 Vanlev: Enzymatic Synthesis of Thiazepine

To optimize and shorten the synthesis of Vanlev, the intermediate [4S-(4a,7a,10ab)]-1-octahydro-5-oxo-4-[[(phenylmethoxy)carbonyl]amino]-7H-pyrido-[2,1-β][1,3]thiazepine-7-carboxylic acid (48, Figure 5.9b) was prepared by an enzymatic process. The oxidation of the ε-amino group of (S)-lysine in the thiol 49 generated in situ from disulfide N²-[N-(phenylmethoxy)carbonyl]-L-homocysteinyl] L-lysine-1,1′-disulphide (50) was developed to produce compound 48 using L-lysine ε-aminotransferase [LAT] from S. paucimobilis SC 16113 cloned and overexpressed in E. coli.⁵⁰ The aminotransferase reaction required α-ketoglutarate as the amine acceptor. The glutamate formed during this reaction was recycled back to α-ketoglutarate by means of glutamate oxidase [GOX] from Streptomyces noursei SC 6007.

The biotransformation of compound 50 to afford 48 was carried out using LAT from E. coli GI724 [pal781-LAT] in the presence of α-ketoglutarate and dithiothreitol or tributylphosphine and glutamate oxidase. Reaction yields of 65–67% were obtained for 48. To reduce the cost of producing two enzymes, the transamination reactions were carried out in the absence of GOX and with higher levels of α-ketoglutarate. The reaction yields in the absence of GOX averaged only about 33–35%. However, the reaction yield increased to 70%, by increasing the α-ketoglutarate level to 40 mg mL⁻¹ (a ten-fold increase in concentration), and conducting the reaction at 40 °C, equivalent to that in the presence of GOX.

5.6.4 Captopril: Enzymatic Preparation of (S)-3-Benzoylthio-2-methylpropanoic Acid

Captopril, designated chemically as 1-[(2S)-3-mercapto-2-methylpropionyl]-L-proline (51), and zofenopril (52) are angiotensin converting enzyme inhibitors (ACE) (Figure 5.10). They are used as antihypertensive agents through suppression of the renin–angiotensin–aldosterone system.^51–53 Both prevent the conversion of angiotensin I into angiotensin II by inhibition of ACE. The potency of captopril and zofenopril as ACE inhibitors depends critically on the configuration of the mercapto-alkanoyl moiety; the compound with the (S)-configuration is about 100 times more active than its corresponding (R)-isomer.^51–53

Figure 5.10 Antihypertensive drug, Captopril: enzymatic preparation of (S)-3-benzoylthio-2-methylpropanoic acid (55).

The required 3-mercapto-(2S)-methyl-propionic acid moiety has been prepared from the microbially-derived chiral 3-hydroxy-(2R)-methylpropionic acid, which is obtained by the hydroxylation of isobutyric acid.⁵⁴ In an alternative approach to prepare the chiral side-chain of captopril and zofenopril (Figure 5.10), the lipase-catalyzed enantioselective esterification of racemic 3-benzoylthio-2-methylpropanoic acid (±)-53 in an organic solvent was demonstrated to yield (R)-(+)-methyl ester 54 and unreacted acid enriched in the desired (S)-55.⁵⁵ Using lipase PS-30 with toluene as solvent and methanol as nucleophile, the desired (S)-55 was obtained in 37% yield (maximum theoretical yield is 50% for a kinetic resolution) with 97% ee. The amount of water and the concentration of methanol supplied in the reaction mixture were very critical. Water was used at 0.1% v/v concentration in the reaction mixture. Greater than 1% v/v water led to the aggregation of enzyme in the organic solvent with a decrease in the rate of reaction due to mass transfer limitation.

The rate of esterification decreased as the methanol-to-substrate ratio was increased from 1 : 1 to 4 : 1. Higher methanol concentrations inhibited the esterification reaction by stripping the essential water from the enzyme. Crude lipase PS-30 was immobilized on Accurel polypropylene (PP) with absorption efficiencies of 98.5%. The immobilized lipase efficiently catalyzed the esterification reaction at substrate input of 224 g L⁻¹ with 0.4 M methanol, 0.1% v/v water, and 120 g L⁻¹ immobilized enzyme giving a 45% reaction yield of (S)-55 with 97.7% ee. The immobilized enzyme under identical process conditions gave a similar ee and yield of product in 23 additional reaction cycles without any loss of activity and productivity.⁵⁵ (S)-55 is a key chiral intermediate in the synthesis of zofenopril⁵⁶ or captopril.⁵⁷

5.7 Antiviral Drugs. Case Study: Atazanavir

5.7.1 Atazanavir: Enzymatic Synthesis of (S)-Tertiary-leucine

Atazanavir (56) is an acyclic aza-peptidomimetic, a potent HIV protease inhibitor^58,59 approved by the Food and Drug Administration for treatment of auto immune diseases (AIDS). The synthesis of atazanavir required (S)-tertiary leucine (57, Figure 5.11a). Furthermore, due to its bulky and hydrophobic side-chain, (S)-tertiary leucine (57) is also a key chiral amino acid required for the synthesis of a number of other drugs containing peptides such as boceprevir and telaprevir.^60,61

Figure 5.11 Atazanavir: (a) enzymatic synthesis of (S)-tertiary-leucine 57; (b) enzymatic preparation of (1S,2R)-[3-chloro-2-hydroxy-1-(phenylmethyl)propyl]-carbamic acid, 1,1-dimethyl-ethyl ester (59).

Using a leucine dehydrogenase and a formate dehydrogenase from Candida boidinii to enzymatically reduce 2-ketocarboxylic acids, with in situ cofactor regeneration, constitutes an industrially established method for preparing optically active L-α-amino acids. In particular, this route is suitable for synthesizing the non-proteinogenic amino acid L-tert-leucine, which is produced on the ton scale via this biocatalytic method using a membrane reactor. This successful strategy has been described in detail in the open literature.^62,63 However, a disadvantage of this method is the requirement of isolated enzymes in purified form.

Therefore, an enzymatic reductive amination of ketoacid 58 to amino acid 57 by recombinant E. coli expressing leucine dehydrogenase from Thermoactinomyces intermedius has been demonstrated. The reaction required ammonia and NADH as a cofactor, which was recycled using recombinant E. coli expressing formate dehydrogenase from Pichia pastoris. Reaction yields of >95% with an ee of >99.5% was obtained for (S)-tertiary leucine at 100 g L⁻¹ substrate input (R. Hanson, S. Goldberg, and R. Patel, unpublished results).

Soda et al.⁶⁴ describe the use of a whole-cell catalyst for the enzymatic synthesis of L- and D-amino acids from α-keto acids with E. coli cells expressing heterologous genes. L-Amino acids were produced with thermostable (S)-amino acid dehydrogenase and formate dehydrogenase (FDH) from α-keto acids and ammonium formate with only an intracellular pool of NAD⁺ for the regeneration of NADH. They constructed plasmids containing, in addition to the FDH gene, the genes for amino acid dehydrogenases, including leucine dehydrogenase, alanine dehydrogenase and phenylalanine dehydrogenase. L-Leucine, L-valine, L-norvaline, L-methionine, L-phenylalanine and L-tyrosine were synthesized in the presence of the recombinant E. coli cells with high chemical yields (>80%) and high optical yields (up to >99.9% ee). If preference were to be given to taking advantage of the intracellular pool of NAD⁺ in E. coli, the final concentration of product would be restricted to about 0.35–0.38 M.

Gröger et al.⁶⁵ have developed a process for preparing enantiomerically enriched (S)-α-amino acids or their salts by reacting the corresponding 2-ketocarboxylic acid with an ammonium ion donor in the presence of a whole-cell catalyst, which consists of a cloned gene encoding a cofactor-dependent amino acid dehydrogenase and a cloned gene encoding an enzyme that regenerates the cofactor. A total input of substrate per reaction volume was >500 mM, with the addition of the substrate being metered such that the stationary concentration of 2-ketocarboxylic acid remains <500 mM, and that the external addition of cofactor – based on the total input of substrate – corresponds to <0.0001 equivalents. Reaction yields of 84% and ee of 99% was obtained for (S)-tertiary leucine at 130 g L⁻¹ substrate input.

5.7.2 Atazanavir: Enzymatic Preparation of (1S,2R)-[3-Chloro-2-hydroxy-1-(phenylmethyl)propyl]carbamic Acid, 1,1-Dimethyl-ethyl Ester

Focusing again on atazanavir (56, Figure 5.11b), an enzymatic process was developed for the preparation of (1S,2R)-[3-chloro-2-hydroxy-1-(phenylmethyl)propyl]carbamic acid 1,1-dimethylethyl ester 59, a key chiral intermediate. The diastereoselective reduction of (1S)-[3-chloro-2-oxo-1-(phenylmethyl)propyl] carbamic acid, 1,1-dimethylethyl ester (60) was carried out using Rhodococcus, Brevibacterium and Hansenula strains to provide 59. Three strains of Rhodococcus gave >90% yield with a diastereomeric purity of >98% and an ee of 99.4%.⁶⁶ An efficient single-stage fermentation–biotransformation process was developed for the reduction of ketone 60 with cells of Rhodococcus erythropolis SC 13845 to yield 59 in 95% yield with a diastereomeric purity of 98.2% and an ee of 99.4% at a substrate input of 10 g L⁻¹. The reduction process was further improved by generating mutants and selection of desired mutant for conversion of 60 into (1S,2R)-59 at substrate inputs of 60 g L⁻¹. Compound (1S,2R)-59 was converted into epoxide 61 and used in the synthesis of atazanavir.⁶⁷ Chemical reduction of chloroketone 60 using NaBH₄ produces the undesired chlorohydrin diastereomer.⁶⁸

In this area, other companies have been active. Codexis has developed a ketoreductase that catalyzed the reduction of 60 to (1S,2R)-59. Wild-type recombinant ketoreductase gave only 5% conversion and 80% enantiopurity at a substrate input of 20 g L⁻¹ and enzyme input of 5 g L⁻¹ in 24 h. However, the enzyme was evolved based on ProSAR, generated library and screening of entire library of mutants. They developed a highly stereospecific and diastereoselective reduction process with isopropanol in water as solvent and NADP as cofactor by a custom-evolved ketoreductase. This enzyme catalyzed the reaction at 200 g L⁻¹ substrate input with 1 g L⁻¹ enzyme input and 0.01 g L⁻¹ NADP input with 99.9% conversion and >99% enantiomeric purity in 24 h reaction time.⁶⁹

5.8 Antianxiety Drug. Buspirones: Enzymatic Preparation of 6-Hydroxybuspirone

Buspirone (Buspar® 62, Figure 5.12) is used for treatment of anxiety and depression. Its therapeutic effects are thought to occur by binding to the serotonin 5HT1A receptor.^70,71 As a result of physiological hydroxylation reactions, buspirone is extensively converted into various metabolites^70–73 and blood concentrations return to low levels a few hours after dosing. A major metabolite, 6-hydroxybuspirone, produced by the action of liver cytochrome P450 CYP3A4, is present at much higher concentrations in human blood than buspirone itself. Remarkably, this metabolite has anxiolytic effects in an anxiety model using rat pups and binds to the human 5-HT1A receptor. Although the metabolite has only about a third of the affinity for the human 5HT1A receptor as buspirone, it is present in human blood at 30–40 times higher concentration than buspirone following a dose of buspirone, and therefore may be responsible for much of the effectiveness of the drug. For the development of 6-hydroxybuspirone as a potential antianxiety drug, preparation and testing of the two enantiomers as well as the racemate was of interest. Both the (R)- and (S)-enantiomers, isolated by chiral HPLC, were effective in tests using a rat model of anxiety. Whereas the (R)-enantiomer showed somewhat tighter binding and specificity for the 5HT1A receptor, the (S)-enantiomer had the advantage of being cleared more slowly from the blood.

Figure 5.12 Buspirones: enzymatic preparation of 6-hydroxybuspirone.

An enzymatic process was developed for resolution of 6-acetoxybuspirone (63, Figure 5.12a). L-Amino acid acylase from Aspergillus melleus (Amano acylase 30 000) was used to hydrolyze racemic 6-acetoxybuspirone to (S)-6-hydroxybuspirone (64) in 96% ee after 46% conversion. The remaining (R)-6-acetoxybuspirone (65) with 84% ee was converted into (R)-6-hydroxybuspirone (66) by acid hydrolysis.⁷⁴ The ee of both enantiomers could be improved to >99% by subsequent crystallization. Direct hydroxylation of buspirone to (S)-6-hydroxbuspirone by Streptomyces antibioticus ATCC 14980 has also been described.⁷⁴

In an alternative process to obtain >50% yield, an enantioselective microbial reduction of 6-oxobuspirone (67, Figure 5.12b) to either (S)-6-hydroxybuspirone (64) or (R)-6-hydroxybuspirone (66) was described.⁷⁵ About 150 microorganisms were screened for the enantioselective reduction process. From them, Rhizopus stolonifer SC 13898, Rhizopus stolonifer SC 16199, Neurospora crassa SC 13816, Mucor racemosus SC 16198, and Pseudomonas putida SC 13817 gave >50% reaction yields and >95% ee of (S)-6-hydroxybuspirone. The yeast strains Hansenula polymorpha SC 13845 and Candida maltosa SC 16112 gave (R)-6-hydroxybuspirone in >60% reaction yield and >97% ee.

In a subsequent step of the research, the NADP-dependent (R)-reductase (RHBR) which catalyzes the reduction of 6-oxobuspirone to (R)-6-hydroxybuspirone was purified to homogeneity from cell extracts of Hansenula polymorpha SC 13845, cloned and expressed in E. coli. To regenerate the required NADPH cofactor, the glucose-6-phosphate dehydrogenase gene from Saccharomyces cerevisiae was also cloned in the same E. coli.^75,76

The NAD-dependent (S)-reductase (SHBR), which catalyzes the reduction of 6-ketobuspirone to (S)-6-hydroxybuspirone, was also purified to homogeneity from cell extracts of Pseudomonas putida SC 16269, and again cloned and expressed in E. coli. In this particular case, for NADH regeneration required for the reduction, the formate dehydrogenase gene from Pichia pastoris was cloned and expressed in E. coli as well. Recombinants E. coli expressing (S)-reductase or (R)-reductase catalyzed the reduction of 6-ketobuspirone to (S)-6-hydroxybuspirone and (R)-6-hydroxybuspirone, respectively, in >98% yield and >99.9% ee.⁷⁶

5.9 Antiviral Drugs. Hepatitis B Viral (HBV) Inhibitor: Enzymatic Asymmetric Hydrolysis and Acetylation

Chiral monoacetate esters 68 and 69 (Figure 5.13) are key intermediates for total chemical synthesis of Baraclude (70), a potential drug for hepatitis B virus infection.^77–79 Baraclude is a carboxylic analogue of 2′-deoxyguanosine in which the furanose oxygen is replaced with an exocyclic double bond. It has recently been approved by FDA for treatment of HBV infection.

Figure 5.13 Hepatitis B viral (HBV) inhibitor: enzymatic asymmetric hydrolysis and acetylation.

The enzymatic hydrolysis of (1α,2β,3α)]-2-[(benzyloxy)methyl]-4-cyclopenten-1,3-diol diacetate has been demonstrated by Griffith and Danishefsky to afford the corresponding monoester using acetylcholine esterase from electric eel.^80,81 They have used a very expensive enzyme acetylcholine esterase in asymmetric hydrolytic reaction to obtain 98% yield of product with 95% ee. In this area, we have developed a process for the enantioselective asymmetric hydrolysis of (1α,2β,3α)-2-[(benzyloxy)methyl]-4-cyclopenten-1,3-diol diacetate (71, Figure 5.13) to the corresponding (+)-monoacetate 68 by cheap and readily available enzymes lipase PS-30 from Pseudomonas cepacia and also by pancreatin. Reaction yields of 85% and an ee of 98% were obtained using lipase PS-30. Using pancreatin, reaction yields of 75% and an ee of 98.5% were obtained. We have also developed an enzymatic process for the asymmetric acetylation of (1α,2β,3α)-2-[(benzyloxy)methyl]-4-cyclopenten-1,3-diol (72) to the corresponding (−)-monoacetate 69 in 80% yield and 98% ee using lipase PS-30.⁸² Thus, methods for the preparation of both (R)- and (S)-monoacetate were developed.

5.10 Chemokine Receptor Modulators: Enzymatic Desymmetrization of Dimethyl Ester

The chiral monoester, (1S,2R)-2-(methoxycarbonyl)cyclohex-4-ene-1-carboxylic acid (73, Figure 5.14) is a key intermediate for the synthesis of a potential drug candidate 74 for the modulation of chemokine receptor 2 (CCR2) activity useful in treatment of rheumatoid arthritis.^83–85 Both the (1S,2R)-monoester 73 and its enantiomer (1R,2S)-monoester 75 were obtained by the resolution of the racemic acid 73 with alkaloids.^86,87 The meso-desymmetrization process has been developed affording either the (1S,2R)-monoester 73 or its enantiomer (1R,2S)-monoester 75 by desymmetrization of the meso-anhydride, cis-1,2,3,6-tetrahydrophthalic anhydride (76) by alcoholysis using cinchona alkaloids. The (1R,2S)-monoester 75 has been synthesized by porcine liver enzyme-catalyzed desymmetrization of the diethyl ester 77.^88,89

Figure 5.14 Chemokine receptor modulators: enzymatic desymmetrization of a dimethyl ester.

At Bristol-Myers Squibb we have initially developed the quinine catalyzed alcoholysis of the anhydride 76 to prepare kilogram quantities of the (1S,2R)-monoester 73 with 90.8% ee. To improve the enantioselectivity of desired 73, an alternative enzymatic process was evaluated.⁹⁰ Thus, a screening of various enzymes was carried out to prepare 73. After evaluating yields and ee values of the desired product, reaction rates, and the cost of the enzyme, the immobilized lipase from Candida antarctica (Novozym 435) was chosen for further development of the desymmetrization of 77 to the desired monoester 73. After optimizing the reaction parameters such as pH, temperature, substrate and enzyme input, a small scale reaction at 50 mL (57.2 g) of diester 77 was conducted. A yield of 96% and >99.9% ee was obtained for the desired (1S,2R)-monoester 73 after 24 h reaction. Preparative, kilogram-scale batches were performed to prepare 3.4 kg of (1S,2R)-monoester 73 in two batches. The two batches were performed on a 1.7 kg scale with an observed conversion profile similar to that of the smaller-scale batch. The desired (1S,2R)-monoester was obtained in >99.9% ee, 98.5% yield and 99% purity.⁹⁰

5.11 Anticancer Drugs

5.11.1 Paclitaxel Semisynthetic Process

Antimitotic agents such as paclitaxel (Taxol®) (78, Figure 5.15a), a complex, polycyclic diterpene, exhibit a unique mode of action on microtubule proteins responsible for the formation of the spindle during cell division. Various types of cancers have been treated with paclitaxel and it was approved for use by the FDA for treatment of ovarian cancer and metastatic breast cancer. Paclitaxel was originally isolated from the bark of the yew Taxus brevifolia, and it has also been found in other Taxus species. Paclitaxel was obtained from T. brevifolia bark in very low (0.07%) yield, and cumbersome purification of paclitaxel from other related taxanes was required. It is estimated that about 20 000 pounds of yew bark (equivalent to about 3000 trees) are needed to produce 1 kg of purified paclitaxel.^91,92 Therefore, the development of a semi-synthetic process for the production of paclitaxel from baccatin III (79) or 10-deacetylbaccatin III (10-DAB) (80) and C-13 paclitaxel side-chain (2R,3S)-81, or acetate (R)-82 was a very promising approach. Taxanes, baccatin III and 10-DAB can be derived from renewable resources such as the needles, shoots and young Taxus cultivars.⁹³ Thus, the preparation of paclitaxel by a semi-synthetic process would eliminate the harvesting of yew trees.

Figure 5.15 Paclitaxel (78) semi-synthetic process: (a) C-13 taxolase, (b) C-10 deacetylase and (c) C-7 xylosidase.

Using selective enrichment techniques, two strains of Nocardioides were isolated from soil samples that contained the novel enzymes C-13 taxolase and C-10 deacetylase.^93–95 The extracellular C-13 taxolase derived from the filtrate of the fermentation broth of Nocardioides albus SC 13911 catalyzed the cleavage of the C-13 side-chain from paclitaxel and related taxanes such as taxol C (see side-chain 83), cephalomannine (see side-chain 84), 7-β-xylosyltaxol, 7-β-xylosyl-10-deacetyltaxol and 10-deacetyltaxol (Figure 5.15a). The intracellular C-10 deacetylase derived from fermentation of Nocardioides luteus SC 13912 catalyzed the cleavage of the C-10 acetate from paclitaxel, related taxanes and baccatin III to yield 10-DAB (Figure 5.15b). The C-7 xylosidase derived from fermentation of Moraxella sp. (Figure 5.15c) catalyzed the cleavage of the C-7 xylosyl group from various taxanes such as xylosylpaclitaxel (85), xylosyltaxol C (86), xylosylcephalomannine (87) to paclitaxel, taxol C, and cephalomannine, respectively.⁹⁶

Fermentation processes were developed for growth of N. albus SC 13911 and N. luteus SC 13912 to produce C-13 taxolase and C-10 deacetylase, respectively, in 5000 L batches. A bioprocess was demonstrated for the conversion of paclitaxel and related taxanes in extracts of Taxus cultivars into the single compound 10-DAB using both enzymes. The concentration of 10-DAB was increased by 5.5 to 24 by treatment with the two enzymes. The bioconversion process was also applied to extracts of the bark of T. brevifolia to give a 12-fold increase in 10-DAB concentration. The enhancement of the 10-DAB concentration in yew extracts was useful in increasing the amount and ease of purification of this key precursor for the paclitaxel semi-synthetic process using renewable resources.⁹⁴

Another key precursor for the paclitaxel semi-synthetic process is the chiral C-13 paclitaxel side-chain. The enantioselective microbial reduction of 2-keto-3-(N-benzoylamino)-3-phenylpropionic acid ethyl ester (88, Figure 5.16a) to yield (2R,3S)-N-benzoyl-3-phenylisoserine ethyl ester (81) was demonstrated using two strains of Hansenula.⁹⁷ Preparative-scale bioreduction of 88 was demonstrated using cell suspensions of Hansenula polymorpha SC 13865 and Hansenula fabianii SC 13894 in independent experiments. In both batches, a reaction yield of >80% and ee’s of >94% were obtained for (2R,3S)-81. In a single-step bioreduction process, cells of H. fabianii were grown in a 15 L fermentor for 48 h, and then the bioreduction process was initiated by the addition of 30 g of substrate and 250 g of glucose, reacting for 72 h. A reaction yield of 88% with an ee of 95% was obtained for (2R,3S)-81.⁹⁷

Figure 5.16 Paclitaxel side-chain synthesis: (a) enzymatic reduction process and (b) resolution process.

In an alternate process for the preparation of the C-13 paclitaxel side-chain, the enantioselective enzymatic hydrolysis of racemic acetate cis-3-(acetoxy)-4-phenyl-2-azetidinone (89, Figure 5.16b) to the corresponding (S)-alcohol 90 and the unreacted desired (R)-acetate 91 was demonstrated using lipase PS-30 from Pseudomonas cepacia (Amano International Enzyme Company), and BMS lipase (extracellular lipase derived from the fermentation of Pseudomonas sp. SC 13856).⁹⁸ Reaction yields of 48% (theoretical maximum yield 50%) with ee >99.5% were obtained for the (R)-91. BMS lipase and lipase PS-30 were immobilized on Accurel polypropylene (PP), and the immobilized lipases were reused (ten cycles) without loss of enzyme activity, productivity or the ee of the product (R)-91. The enzymatic process was scaled up to 250 L (2.5 kg substrate input) using immobilized BMS lipase and lipase PS-30, respectively. From each reaction batch, (R)-acetate 91 was isolated in 45% yield (theoretical maximum yield 50%) and 99.5% ee. The (R)-acetate was chemically converted into (R)-alcohol 82. The C-13 paclitaxel side-chain synthon, (2R,3S)-81 or (R)-82 produced either by the reductive process or resolution process, respectively, could be coupled to baccatin III (79) or 10-deacetylbaccatin III (80) after protection and deprotection steps to prepare paclitaxel by a semi-synthetic process.^91–93

5.11.2 Water-soluble Taxane Derivatives

Owing to the poor solubility of paclitaxel, various research groups have been involved in the development of water-soluble taxane analogs.^99–101 Taxane (92, Figure 5.17) is a water-soluble derivative which, when given orally, was as effective as paclitaxel in five tumor models [murine M109 lung and C3H mammary 16/C cancer, human A2780 ovarian cancer cells (grown in mice and rats) and HCT/pk colon cancer].^102,103

Figure 5.17 Water-soluble taxane derivatives: (a) side-chain synthesis and (b) semi-synthetic process.

The chiral intermediate (3R-cis)-3-acetyloxy-4-(1,1-dimethylethyl)-2-azetidinone (93, Figure 5.17a) was prepared for the semi-synthesis of the new orally active taxane 92. The enantioselective enzymatic hydrolysis of racemic cis-3-acetyloxy-4-(1,1-dimethylethyl)-2-azetidinone (94) was carried out to afford the corresponding undesired (S)-alcohol 95 and unreacted desired (R)-acetate 93 using immobilized lipase PS-30 or BMS lipase. Reaction yields higher than 48% (theoretical maximum yield 50%) with ee’s of >99% were obtained for the (R)-acetate 93. Acetoxy β-lactam 93 was converted into (R)-hydroxy-β-lactam 96 for its use in the semi-synthesis of 92.¹⁰⁴

The synthesis of oral taxane 92 also required 4,10-dideacetylbaccatin III (97, Figure 5.17b) as starting material for the chemical synthesis of the C-4 methyl-carbonate derivative of 10-deacetyl baccatin III (98). A microbial process was developed for deacetylation of 10-deacetylbaccatin III (80) to 4,10-dideacetylbaccatin III (97) using a Rhodococcus sp. SC 162949 isolated from soil using culture enrichment techniques.¹⁰⁵

5.11.3 Epothilones: Epothilone B and Epothilone F

The clinical success of paclitaxel has stimulated research into compounds with similar modes of activity, in an effort to emulate its antineoplastic efficacy while minimizing its less desirable aspects, which include non-water solubility, difficult syntheses, and emerging resistances. In this respect, the epothilones are a novel class of natural product cytotoxic compounds derived from the fermentation of the myxobacterium Sorangium cellulosum. Epothilones are non-taxane microtubule-stabilizing compounds that trigger apoptosis.^106,107 It is now known that epothilones exert microtubule-stabilizing effects similar to paclitaxel (Taxol®), and cytotoxic activity against rapidly proliferating cells such as tumor cells or other hyper-proliferative cellular diseases. The natural product epothilone B (99, Figure 5.18) has demonstrated broad spectrum antitumor activity in vitro and in vivo, including tumors with paclitaxel resistance. As stated above, the epothilone analogs were synthesized in an effort to optimize the water solubility, in vivo metabolic stability and antitumor efficacy of this class of antineoplastic agents.^108–111 Ixabepilone 100, a semi-synthetic intravenous epothilone derived from natural product epothilone B, was the first epothilone to be tested in humans, and recently received US Food and Drug Administration (FDA) approval as an anticancer agent. The drug is indicated as monotherapy for the treatment of metastatic or locally advanced breast cancer following the failure of an anthracycline, a taxane, and capecitabine (Xeloda), and as combination therapy with capecitabine following the failure of an anthracycline and a taxane.^111,112

Figure 5.18 Epothilones: epothilones B, epothilone F and hydroxylation of epothilone B to epothilone F.

A fermentation process was developed for the production of epothilone B, and the titers of epothilone B were optimized and increased by a continuous feed of sodium propionate during fermentation. The inclusion of XAD-16 resin during fermentation to adsorb epothilone B and to carry out volume reduction made the recovery of product very simple.¹⁰⁷ In addition, high levels of free epothilone B – which were inhibitory to the growth of producing culture – were avoided by supplying XAD-16 resin during the fermentation process.

To develop another epothilone derivative as an anticancer agent, 102, a microbial hydroxylation process was developed for the conversion of epothilone B (99) into epothilone F (101) by Amycolatopsis orientalis SC 15847. Bioconversion yields of 37–47% were obtained when the process was scaled up to 100–250 L. The epothilone B hydroxylase along with the ferredoxin gene from A. orientalis SC 15847 have been cloned and expressed in Streptomyces rimosus at Bristol-Myers Squibb. Mutants and variants thereof of this cloned enzyme have been used in the hydroxylation of epothilone B to epothilone F to obtain >87% yields of products.^113,114

5.11.4 IGF-1 Receptor Inhibitor: Enzymatic Preparation of (S)-2-Chloro-1-(3-chlorophenyl)ethanol

The synthesis of the leading candidate compound in an anticancer program^115,116 required (S)-2-chloro-1-(3-chlorophenyl)ethanol (103, Figure 5.19) as an intermediate. Other possible candidate compounds used analogs of the (S)-alcohol. From a microbial screening of the reduction of ketone 104 to the (S)-alcohol 103, two cultures displaying the highest enantioselectivity were identified, namely Hansenula polymorpha SC13824 (73.8% ee), and Rhodococcus globerulus SC SC16305 (71.8% ee). A ketoreductase from Hansenula polymorpha (after purification to homogeneity), rendered (S)-alcohol 103 with >99.99% ee. Amino acid sequences from the purified enzyme were used to design PCR primers for cloning the ketoreductase gene. The ketoreductase was cloned and expressed in E. coli together with a glucose-6-phosphate dehydrogenase from Saccharomyces cerevisiae to allow regeneration of the NADPH required by the reduction process. An extract of E. coli containing the two recombinant enzymes was used to reduce 2-chloro-1-(3-chloro-4-fluorophenyl)ethanone and two related ketones to the corresponding (S)-alcohols. Moreover, intact E. coli cells supplemented with glucose were used to prepare (S)-2-chloro-1-(3-chloro-4-fluorophenyl)ethanol 103 in 89% yield with >99.99% ee.¹¹⁷

Figure 5.19 IGF-1 receptor inhibitor: enzymatic preparation of (S)-2-chloro-1-(3-chlorophenyl)ethanol (103).

5.11.5 Retinoic Acid Receptor Agonist: Enzymatic Preparation of 2-(R)-Hydroxy-2-(1′,2′,3′,4′-tetrahydro-1′,1′,4′,4′-tetramethyl-6′-naphthalenyl)acetate

Retinoic acid and its natural and synthetic analogs (retinoids) exert a wide variety of biological effects by binding to or activating a specific receptor or sets of receptors.¹¹⁸ They have been shown to effect cellular growth and differentiation and are promising drugs for the treatment of cancers.¹¹⁹ A few retinoids are already in clinical use for the treatment of dermatological diseases, such as acne and psoriasis.¹²⁰ (R)-3-Fluoro-4-[[hydroxy-(5,6,7,8-tetrahydro-5,5,8,8-tetramethyl-2-naphthalenyl)-acetyl]amino]benzoic acid (105, Figure 5.20) is a retinoic acid receptor gamma-specific agonist potentially useful as a dermatological and anticancer drug.¹²¹ Ethyl 2-(R)-hydroxy-2-(1,2,3,4-tetrahydro-1,1,4,4-tetramethyl-6-naphthalenyl)acetate (106), and the corresponding acid 107, were prepared as intermediates in the synthesis of the retinoic acid receptor gamma-specific agonist.¹²¹ Enantioselective microbial reduction of ethyl 2-oxo-2-(1,2,3,4-tetrahydro-1,1,4,4-tetramethyl-6-naphthalenyl)acetate 108 to alcohol 106 was carried out using Aureobasidium pullulans SC 13849 in 98% yield and with an ee of 96%. At the end of the reaction, hydroxyester 106 was adsorbed onto XAD-16 resin and, after filtration, recovered in 94% yield from the resin with acetonitrile extraction. The recovered (R)-hydroxyester 106 was treated with Chirazyme L-2 or pig liver esterase to convert it into the corresponding (R)-hydroxyacid 107 in quantitative yield. The enantioselective microbial reduction of ketoamide 109 to the corresponding (R)-hydroxyamide 105 by A. pullulans SC 13849 has also been demonstrated.¹²²

Figure 5.20 Retinoic acid receptor agonist: enzymatic preparation of 2-(R)-hydroxy-2-(1,2,3,4-tetrahydro-1,1,4,4-tetramethyl-6-naphthalenyl)acetate (106).

5.12 Microbial Hydroxylation of Mutilin and Pleuromutilin

Pleuromutilin (110, Figure 5.21) is an antibiotic from Pleurotus or Clitopilus basidiomycetes strains, which kills mainly Gram-positive bacteria and mycoplasms. A more active semisynthetic analogue, tiamulin, has been developed for the treatment of animals and poultry infection, and has been shown to bind to prokaryotic ribosomes and inhibit protein synthesis.¹²³ The metabolism of pleuromutilin derivatives results in hydroxylation by microsomal cytochrome P-450 at the 2- or 8-position and inactivates the antibiotics.¹²⁴ Microbial hydroxylation of pleuromutilin (110) or mutilin (111) would provide a functional group at this position to allow further modifications. The target analogues would maintain the biological activity of the parent compounds but would not be susceptible to metabolic inactivation. Biotransformation of mutilin and pleuromutilin by microbial cultures was carried out to provide a source of 8-hydroxymutilin or 8-hydroxypleuromutilin.¹²⁵Streptomyces griseus strains SC 1754 hydroxylated mutilin to (8S)-hydroxymutilin (112), (7S)-hydroxymutilin (113), and (2S)-hydroxymutilin (114) while Cunninghamella echinulata SC 16162 gave (2S)-hydroxymutilin or (2R)-hydroxypleuromutilin (115) from biotransformation of mutilin or pleuromutilin, respectively. The biohydroxylation of mutilin by the S. griseus strain SC 1754 was scaled up to a 100 L fermentor to produce a total of 49 g of (8S)-hydroxymutilin, 17 g of (7S)-hydroxymutilin, and 13 g of (2S)-hydroxymutilin from 162 g of mutilin.¹²⁵ A C-8 ketopleuromutilin 116 derivative has been synthesized from the biotransformation product 8-hydroxymutilin which upon acid catalyzed conversion provided novel pleuromutilin 117.¹²⁶

Figure 5.21 Microbial hydroxylation of pleuromutilin (110) and mutilin (111).

5.13 Conclusions

The demand for chiral compounds continues to rise in general, and in particular a very high demand from the pharmaceutical industry is fueled by the need for chiral active pharmaceutical ingredients (APIs), in recognition of the fact that enantiomers of a chiral compound could have dramatically different biological activities and toxicity. Chiral APIs were previously usually formulated as racemates, and the preference nowadays is for single enantiomers. Furthermore, the switch from a racemic to a single-enantiomer API is key to the life cycle management. To meet these challenges, pharmaceutical industries are evaluating a variety of chiral technologies, including biocatalysis, to deliver single APIs enantiomers. In addition, apart from those APIs, many other nutraceuticals and fine chemicals used in flavors, fragrance and agrochemicals are chiral molecules.

Microbial cultures and enzymes derived therefrom are highly enantio-, chemo-, and regioselective across a diverse range of reactions under mild conditions of pH, temperature, and pressure. Enzymes (biocatalysts) can be overexpressed and immobilized and reused for many cycles to make biocatalytic processes economically attractive. By using techniques like “directed evolution” and “random mutagenesis”, enzymes can be tailored for certain process conditions, increasing activity towards specific substrate, enhance (or invert) selectivity, increase tolerance to solvents, attenuate inhibition by products or substrates, change the thermostability, or change the pH tolerance, among other options. Given that importance, many industrial and academic laboratories are now using enzyme engineering and high-throughput screening methods during process development.

Microbial P-450 systems are quite useful to generate various drug metabolites – via enantioselective and regioselective hydroxylations, epoxidations, n-oxidations, or de-alkylation reactions – for biological activity testing as well as generating novel compounds. For instance, mutants of bacterial cytochrome P450 from Bacillus megaterium have been developed and are now being commercialized. The availability of “off the shelf” enzymes has been significantly increased. In addition to hydrolytic enzyme such as lipases, proteases and esterases, more enzyme collections such as ketoreductases (conversion of ketones into chiral alcohols), amino acid dehydrogenases (conversion of keto acid into chiral amino acids), transaminases (conversion of keto acid and ketone into chiral amino acids and amines), enoate reductases (hydrogenation type reactions), nitrilases, acylases, and epoxide hydrolases are now available, and can be used for rapid screenings towards various substrates. In many cases the biocatalytic step can provide a direct approach to the desired intermediate without requiring any protection/de-protection steps typically needed in chemical synthesis.

Dehydrogenases, aminotransferases and amino acid dehydrogenases have been successfully used along with cofactors and cofactor regenerating enzymes for the synthesis of chiral alcohols, amino-alcohols, amino acids and amines. Aldolases, decarboxylases and lyases are used effectively in asymmetric synthesis by aldol condensation, acyloin condensation reactions.

Monooxygenases have been used in enantioselective and regioselective hydroxylation, epoxidation and Baeyer–Villiger reactions. Dioxygenases have been used in the chemo-enzymatic synthesis of chiral diols. A current challenge in asymmetric synthesis is the preparation of chiral compounds in 100% chemical and optical yields starting from (inexpensive) racemates. Several approaches have been described so far, such as re-racemization and repeated resolution, dynamic kinetic resolutions, or the transformation of enantiomers via enantioconvergent pathways, usually achieved by combination of chemo- and/or biocatalysts in sequential reactions or by a single biocatalyst.

During the last decade, progress in fermentation technologies, protein chemistry, molecular cloning, random and site-directed mutagenesis, bio-recovery processes and directed evolution of biocatalysts has opened up unlimited access to various enzymes and microbial cultures as tools in organic synthesis.

Acknowledgements

The author would like to acknowledge Drs Ronald Hanson, Animesh Goswami, Amit Banerjee, Venkata Nanduri, Jeffrey Howell, Steven Goldeberg, Robert Johnston, Mary-Jo Donovan, Dana Cazzulino, Thomas Tully, Thomas LaPorte, Lawrence Parker, John Wasylyk, Michael Montana, Ronald Eiring, Rapheal Ko, Linda Chu, Clyde McNamee, Paul Cino, Laszlo Szarka, John Scott, Richard Mueller, Robert Waltermire and many chemists and chemical engineers at Bristol-Myers Squibb for research collaboration in various projects.

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