Development of a Chemoenzymatic Process for Dehydroepiandrosterone Acetate Synthesis

Article abstract of DOI:10.1021/acs.oprd.6b00215

Dehydroepiandrosterone (DHEA, 2) is an important endogenous steroid hormone in mammals used in the treatment of a variety of dysfunctions in female and male health,1 as well as an intermediate in the synthesis of steroidal drugs, such as abiraterone acetate which is used for the treatment of prostate cancer.2-4 In this manuscript we describe a novel, concise, and cost-efficient route toward DHEA (2) and DHEA acetate (3) from 4-androstene-3,17-dione (4-AD, 1). Crucial to success was the identification of a ketoreductase from Sphingomonas wittichii for the highly regio- and stereoselective reduction of the C3-carbonyl group of 5-androstene-3,17-dione (5) to the required 3β-alcohol (2, >99% de). The enzyme displayed excellent robustness and solvent stability under high substrate concentrations (up to 150 g/L).

Full text of DOI:10.1021/acs.oprd.6b00215

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Full Paper
Development of a Chemoenzymatic Process
for Dehydroepiandrosterone Acetate Synthesis
Anna Fryszkowska, Justine A Peterson, Nichola L. Davies, Colin Dewar, George Evans, Matthew Bycroft,
Neil Triggs, Toni Fleming, Srikanth Gorantla, Garrett Hoge, Michael Quirmbach, Upadhya Timmanna,
Srinivas Reddy Poreddy, D Naresh Kumar Reddy, Vilas H. Dahanukar, and Karen Elizabeth Holt-Tiffin
Org. Process Res. Dev., Just Accepted Manuscript • DOI: 10.1021/acs.oprd.6b00215 • Publication Date (Web): 05 Jul 2016
Downloaded from http://pubs.acs.org on July 9, 2016
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Development of a Chemoenzymatic Process for
Dehydroepiandrosterone Acetate Synthesis
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Anna Fryszkowska,†, ¶ Justine Peterson,† Nichola L. Davies†, ǂ Colin Dewar,† George Evans,†
Matthew Bycroft,† Neil Triggs,† Toni Fleming,† Srikanth Sarat Chandra Gorantla,‡,‡‡ Garrett
Hoge,†,†† Michael Quirmbach,§,§§ Upadhya Timmanna,‡, ¶¶ Srinivas Reddy Poreddy,‡ D Naresh
Kumar Reddy,‡ Vilas Dahanukar,‡ Karen E. HoltꢀTiffin†*
† Dr. Reddy’s Laboratories EU Ltd, Chirotech Technology Centre, 410 Cambridge Science Park,
‡
Cambridge CB4 0PE, United Kingdom; Dr. Reddy’s Laboratories Ltd, Custom Pharmaceutical
§
Services, Bollaram Road, Miyapur, Hyderabad, 500049, India; Dr. Reddy's Laboratories SA
Elisabethenanlage, 11CHꢀ4051 Basel, Switzerland.
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Table of content
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We report a novel chemoenzymatic process to prepare dehydroepiandrosterone acetate (3), a key
building block in the synthesis of antiꢀprostate cancer drug abiraterone acetate (4) from 4ꢀ
androsteneꢀ3,17ꢀdione (1). The key step of our route is a robust and stereoselective enzymatic
reduction of 5–androsteneꢀ3,17ꢀdione (5) to dehydroepiandrosterone (2, >99% de) at high
substrate concentration (100ꢀ150 g/L), highlighting the potential of ketoreductases in industrial
steroid drug manufacture.
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Abstract
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Dehydroepiandrosterone (DHEA, 2) is an important endogenous steroid hormone in mammals
used in the treatment of a variety of dysfunctions in female and male health,1 as well as an
intermediate in the synthesis of steroidal drugs, such as abiraterone acetate which is used for the
treatment of prostate cancer.2–4 In this manuscript we describe a novel, concise and costꢀefficient
route towards DHEA (2) and DHEA acetate (3) from 4–androsteneꢀ3,17ꢀdione (4ꢀAD, 1).
Crucial to success was the identification of a ketoreductase from Sphingomonas wittichii for the
highly regioꢀ and stereoselective reduction of the C3ꢀcarbonyl group of 5ꢀandrosteneꢀ3,17ꢀdione
(5) to the required 3βꢀalcohol (2, >99% de). The enzyme displayed excellent robustness and
solvent stability under high substrate concentrations (up to 150 g/L).
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KEYWORDS: Steroids, stereoselective synthesis, reduction, biocatalysis, ketoreductases,
screening.
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1. Introduction
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3βꢀHydroxyandrostꢀ5ꢀenꢀ17ꢀone (2), also known as dehydroepiandrosterone (DHEA) or
prasterone is an important endogenous steroid hormone1 and a precursor in the synthesis of other
steroidal drugs such as abiraterone acetate (Zytiga®), a drug developed for the treatment of
castrationꢀresistant prostate cancer.2–4 Current routes towards DHEA (2) and DHEA acetate (3)
are typically based on the selective degradation of yamꢀderived diosgenin to 16ꢀ
dehydropregnenolone acetate (29),5,6 followed by oxime formation and rearrangement (Scheme
1).5–8
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Scheme 1. Diosegenin (28) and stigmasterol (31) as precursors for DHEA acetate (3) synthesis.
Dependence on rawꢀmaterials with variable availability often drives the development of
alternative processes for steroid drug manufacture.5 Here, we aimed to develop a novel and
scalable route to DHEA acetate (3) from 4ꢀandrosteneꢀ3,17ꢀdione (4ꢀAD, 1), which is derived
from phytosterols obtained from soybeans (e.g. stigmasterol, 31; Scheme 1).5,9–11
Nature offers a vast array of enzymes that can be used to access the structural and functional
diversity of steroids10,11and they often display high substrate specificity, as well as remarkable
regioꢀ and stereoselectivity. During the initial phase of route scouting for possible ways to make
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dehydroepiandrosterone acetate (3) from 4ꢀAD (1) we considered several possible
chemoenzymatic approaches, including routes A and B depicted in Scheme 2.
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Scheme 2. Proposed synthetic strategies to convert 4ꢀAD (1) to DHEA acetate (3).
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Although hydrolases can be used for the regioselective esterification of polyhydroxylated
steroid derivatives,12,13 our initial trials with 5ꢀandrosteneꢀ3,17ꢀdiol (5ꢀADꢀdiol, 6, Scheme 2,
Route A) indicated that with Novozyme 435 (Candida antarctica Lipase B) the reaction
progressed slowly and required high enzyme loading (60h, 100% w/w enzyme, complete
conversion to 28). Inspired by natural metabolic pathways for androgens, we turned our attention
to oxidoreductases (Route B, Scheme 2) .14–20 In mammals 4ꢀAD (4) is synthesised from DHEA
(2) by the action of two enzymes: 3βꢀhydroxysteroid dehydrogenase (3βꢀHSD) catalysing its
oxidation to 5ꢀandrosteneꢀ3,17ꢀdione (5ꢀAD, 5)14,21–26 and ketosteroid isomerase (KSI), which
catalyses the thermodynamically favoured isomerisation of the double bond to the conjugated
α,βꢀunsaturated ketone 1 (Keq = 2400).20,27–30 Thus, we hoped to take advantage of the
reversibility of biooxidation and use an enzyme for the stereoꢀ and regioselective reduction of the
C3ꢀcarbonyl group – a reverse sequence to the natural mammalian metabolic pathway.
Ultimately this proved successful and the results are described hereafter.
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2. Results and Discussion
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In order to successfully convert 4ꢀAD (1) into DHEA acetate (3), we envisaged a three step
process. Firstly, we planned to convert 4ꢀAD to 5ꢀAD (5) through a selective ꢀ4 – ꢀ5 –
isomerisation.31–33 Secondly, we needed to identify a highly stereoselective ketoreductase with
appropriate industrial properties to selectively reduce 5ꢀAD (5) to DHEA (2), which was the key
to success of this project. As ketoreductases and hydroxysteroid dehydrogenases may display
vastlyꢀdifferent degrees of regioselectivity (3ꢀ vs. 17ꢀpositions) and stereoselectivity (α vs.
β),14,21–26 finding the enzyme with exclusive 3βꢀstereoselectivity was crucial. Finally, we planned
to convert DHEA (2) into DHEA acetate (3) and to develop an efficient acetylation and
crystallisation to provide the material with the required purity (≥99.5% by HPLC area).
2.1 Isomerisation
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Several approaches to prepare 5ꢀAD (5) are described in the literature,19,34,35 however these
were considered subꢀoptimal for our process requirements, as they required multiꢀstep syntheses.
Our attention was drawn by short communications on deconjugation of α,βꢀunsaturated
ketones,31–33 including an example of a low yielding potassium hydroxideꢀcatalysed
isomerisation of 4ꢀAD (1).32 After an initial screen, we were able to identify a promising initial
set of reaction conditions that afforded 5ꢀAD (5) in >80% yield (Scheme 3). Treatment of 4ꢀAD
(1) with 10 eq. of potassium tertꢀbutoxide in tertꢀbutanol for 90 min, followed by the kinetic
quench of the extended enolate (32) into 10% v/v aqueous acetic acid allowed precipitation of
the product 5, which was then isolated by filtration.
Scheme 3. Isomerisation of 4ꢀAD (1) to 5ꢀAD (5)
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Encouraged by these results, we proceeded to develop the process further. Initially, we
observed great variability in the reaction outcome, with the relative amount of the desired 5ꢀAD
(5) varying from 50 to 95% (by HPLC area%). From consideration of the isomerisation
mechanism (Scheme 3), the root cause of this problem became evident. The initial formation of
the dianion 32 occurs readily (<30 min) using 2.0ꢀ2.5 equivalents of potassium tertꢀbutoxide.
(Note: while we have drawn this as the dianion, this is purely based on mechanism proposed in
the literature31 and the observation that we required > 2 eq of base and we did not obtain any
additional confirmation of the structure). The subsequent kinetic protonation results in the
desired 5ꢀAD (5) isomer, which is sensitive to even mildly basic conditions and readily
isomerises back to 4ꢀAD (1). Fortunately, the exposure of the 5ꢀAD (5) to basic conditions could
be minimised by quenching the enolate into an excess of acetic acid, rather than by adding the
acid to the reaction mass. The addition rate, efficient mixing and temperature (<30 °C) have to
be carefully controlled to achieve high yields and to prevent localised high pH ‘hotspots’ during
the quench. Failure to control these physical parameters leads to significant isomerisation and
apparent poor conversion. Moreover, 5ꢀAD (5) quenched this way precipitated from the solution
and could be readily isolated by filtration, which is a significant process advantage.
While the controlled quench addressed one of the issues with the reaction, we observed
formation of several other impurities, as evidenced by additional peaks in the HPLC
chromatogram and NMR spectra. The levels of these impurities varied from batch to batch and
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their content changed upon storage of the 5ꢀAD (5). Moreover, the formation of the same
impurities was then also observed during the bioreduction step (vide infra).
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Scheme 4. Impurities generated during isomerisation of 4ꢀAD (1) to 5ꢀAD (5)
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The main impurities were isolated chromatographically and characterised by NMR and MS
(see supplemental info for details). The spectroscopic data obtained for one of the main sideꢀ
products was in agreement with the structure of androstꢀ4ꢀeneꢀ3,6,17ꢀtrione (19, m/z 301
[M+1]+). This oxygenated impurity was reported previously by Pollack et al. and its formation
ascribed to autooxidation of the steroid backbone by molecular oxygen present in the reaction.19
We identified two further impurities, 6βꢀ and 6αꢀhydroxyandrostꢀ4ꢀeneꢀ3,17ꢀdione (22 and 23,
respectively, m/z 303 [M+1]+).36 In addition we were able to isolate small quantities of two
further compounds, which proved unstable and we observed their conversion to trione 19 over
1
the course of the NMR experiments. Based on the similarity of their H NMR spectra to the
isomeric 6ꢀhydroxyandrostꢀ4ꢀeneꢀ3,17ꢀdiones (22 and 23) and mass spectrometry data (both had
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m/z of 319), these compounds were tentatively assigned as 6βꢀ and 6αꢀperhydroxy–androstꢀ4ꢀ
eneꢀ3,17ꢀdiones (20 and 21), respectively. Their presence would be in agreement with the
involvement of oxygen in the sideꢀproduct formation; however no further elucidation of the
mechanism of autoxidation and possible interconversion between the compounds 19-23 was
performed. Nevertheless, this highlighted the need to carry out the isomerisation under oxygenꢀ
free conditions. Thus, the introduction of a rigorous nitrogen sparging protocol during the
reactions reduced the levels of these impurities sufficiently; on 300g scale we were able to
prepare crude 5ꢀAD (5) of acceptable purity: typically 89ꢀ93% by HPLC area, containing 3ꢀ5%
of 4ꢀAD (1) and 3ꢀ4% oxygenated impurities 19-23.
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In summary, our efforts resulted in an efficient 4ꢀAD (1) isomerisation procedure, giving 5ꢀAD
(5) in approx. 80% yield and acceptable purity. We reduced the base requirement from 10 to 2
equivalents of potassium tertꢀbutoxide, providing significant cost reduction for this step. The
material generated under the optimised conditions was used directly in the subsequent
biocatalytic step, as it was not suitable for prolonged storage.
2.2 Development of enzymatic step
Ketoreductase screening. The key step of our proposed route relied on identifying a highly
regioꢀ and stereoselective ketoreductase enzyme to convert 5ꢀAD (5) to the desired 3βꢀ
hydroxyandrostꢀ5ꢀeneꢀ17ꢀone (DHEA, 2) exclusively. The bioreduction of diketone 5 could
theoretically result in up to 16 possible products: 4 possible diastereoisomers of
monohydroxysteroids (C3ꢀOH or C17ꢀOH) and further 4 possible diols (6). An additional 8
isomeric products could be expected, if the double bond moves back to C4. The complexity of
the chromatographic analysis of such a mixture and lack of commercially available reference
materials added to the challenging nature of the enzymatic screen. Therefore to simplify and
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accelerate the identification of regioselective enzymes, we took advantage of the reversibility of
the ketoreductaseꢀcatalysed reactions. Specifically, we screened the enzyme libraries towards the
oxidation of hydroxysteroids as depicted in Figure 1. The screen was performed using a routine
colourimetric formazanꢀlinked assay, which detects reduced NAD(P)H cofactor, formed during
the biooxidation.37 In parallel, we tested ketoreductases for the oxidation of commercially
available DHEA (2), testosterone (9) and androstꢀ5ꢀeneꢀ3β,17βꢀdiol (5ꢀADꢀdiol, 6), observing
redꢀcolouration in the wells with biocatalyst active towards individual compounds after 60 min at
30 °C (the low substrate solubility in aqueous media did not allow monitoring of the reactions
spectrophotometrically and the plates were inspected visually).
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Figure 1. The principle of screening for regioselective ketoreductases in the oxidative direction
using a NAD(P)Hꢀlinked colourimetric assay.
O
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The pictures show snapshots of typical screening plates tested with three different
hydroxysteroids: top panel – DHEA (2); middle panel – 5ꢀADꢀdiol (6); bottom panel –
testosterone (9). NAD(P)H cofactor formed in the oxidation in the presence of tetrazolium (2ꢀ(4ꢀ
iodophenyl)ꢀ3ꢀ(4ꢀnitrophenyl)ꢀ5ꢀphenylꢀ2Hꢀtetrazolium chloride ꢀ INT) and diaphorase causes
redꢀcolouration of the wells with an active enzyme. Highlighted in blue circles – an enzyme with
exclusive activity towards C3βꢀhydroxyl group; in red circles – an enzyme with exclusive
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activity towards C17βꢀhydroxyl group; in grey circles – enzymes active towards both C17ꢀ and
C3ꢀhydroxyl groups (nonꢀselective).
Using this screening strategy we quickly discriminated between the enzymes displaying 3βꢀ vs.
17βꢀregioselectivity (see Figure 1, top vs. bottom panel). Androstꢀ5ꢀeneꢀ3β,17βꢀdiol (6, Figure 1,
middle panel) was added to the screening program as a control to confirm the regioselectivity
towards C3 and C17 hydroxyl groups.
The colourimetric screening strategy simplified and accelerated the screening for
regioselective enzymes, without the need to put effort into complex and lengthy chromatographic
analysis. We efficiently tested a collection of almost 400 proprietary and commercial
ketoreductases to identify a shortꢀchain dehydrogenase/reductase from Sphingomonas wittichii38
as the sole enzyme with the desired 3βꢀstereoselectivity. Subsequent studies on the enzymatic
reduction of 5ꢀAD (5) demonstrated that the Sphingomonas wittichii ketoreductase catalysed
formation of DHEA with high diastereoselectivity (2, >99% de) − spiking experiments showed
no detectable formation of 3αꢀepimer 13 by HPLC. Moreover, the enzyme showed almost no
activity towards the isomer 4ꢀAD (1).
Reaction optimisation.
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We optimised process parameters, such as substrate and ketoreductase loading, coꢀsolvent,
phase ratio, pH, temperature in a series of experiments (details are shown in Table 1). We aimed
to maximise the substrate concentration and to minimise the enzyme loading in order to improve
the volumetric efficiency and the cost of the process. In parallel, we also improved the gene and
the protein expression of the ketoreductase to minimise the fermentation requirement (vide
infra).
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Table 1. Optimisation studies of 5ꢀAD (5) bioreduction.
Parameter
Evaluated range
NAD+/NADP+
6.0ꢀ8.0
Set value
NAD+
6.3
Cofactor
pH
Enzyme loading & form
Enzyme addition
0.25ꢀ10% (w/w or v/w)
1st generation ketoreductase:
3% w/w (powder, 1 batch)a
One batch vs. multiple
additions
2nd generation ketoreductase:
5% v/w (liquid, 4 batches)a
3rd generation ketoreductase:
0.25% w/w (powder, 1 batch)b
Temperature
Cosolventc
20ꢀ50 °C
33 °C
2ꢀMeTHF, EtOAc, MTBE, 2ꢀMeTHF, EtOAc
DIIPE, BuOAc, toluene,
nꢀBuOH, THF, DMSO, DMF
Cosolvent volume
Buffer
0–66% v/v
50ꢀ60% v/v
phosphate
100 g/L
TRIS, phosphate
3ꢀ150 g/L
Substrate conc.
d
MgCl2
0ꢀ5 mM
0 mM
Base
2.0 M NaOH, 1.7 M Na2CO3, 1.5 M K2CO3
1.5 M K2CO3
b
c
a used for the >200gꢀscale campaign; loading for the optimised enzyme powder; optimised
water immiscible coꢀsolvents were initially tested at 30% v/v while water miscible ones at 6%
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v/v; magnesium chloride was added to see if it would improve the protein stability/activity
under the reaction conditions, however no difference was observed
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All the bioreductions were conducted in a Multimaxꢀsystem, using 50 mL reactors with
overhead stirring, temperature control and pHꢀcontrol. We used glucose and glucose
dehydrogenase to recycle the NAD(P)Hꢀcofactors, adjusting the pH with a base (2.0 M NaOH or
1.7 M Na2CO3). The best conversions were obtained at 25ꢀ35 °C and pH 6.0ꢀ7.0. Very early into
the process development, we concluded that biotransformations proceeded more efficiently
under biꢀphasic reaction conditions, using 2ꢀMeTHF or EtOAc – this increased the substrate
solubility in the reaction media and allowed much higher conversions and substrate loadings than
the addition of water miscible coꢀsolvents. Consequently we were able to run >200 gꢀscale
batches at concentrations up to 150 g/L, achieving full conversion. EtOAc was selected as the
best solvent overall, since it was significantly cheaper, although it required 10ꢀ20% higher
enzyme loadings. Moreover, EtOAc offered a process safety advantage, as 2ꢀMeTHF may form
explosive peroxides. We replaced 2 M NaOH with 1.7 M Na2CO3 to avoid localised high pH
‘hotspots’ leading to sideꢀproduct formation. Ultimately, we preferred 1.4 M K2CO3 solution as a
base, as it possesses higher solubility in water in comparison to Na2CO3 (112 g/100 mL vs.
21.5 g/100 mL at 20 °C), which improved process reproducibility and eliminated the risk of line
blockage in the pH control apparatus seen occasionally with saturated sodium carbonate
solutions.
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Under the optimised conditions >200 gꢀscale enzymatic reductions proceeded efficiently using
our 2nd generation biocatalyst (liquid cellꢀfree extract of the gene and expression optimised
ketoreductase). Such biotransformations reached full conversions in 24ꢀ28h, giving crude DHEA
(2) in 88ꢀ90% purity by HPLC. Further improvements of biocatalyst form (powder, 3rd
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generation enzyme) further reduced the time required to reach >95% conversion to 6ꢀ8h at 0.25ꢀ
0.30% w/w enzyme loading.
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During these bioreductions, we observed the formation of similar impurities to the
isomerisation step. Unsurprisingly, their content increased significantly at elevated temperatures
(>40 °C). Under such conditions, at pH 7.5 and in the presence of air, 5ꢀAD (5) decomposed
almost completely over 3 days. The most prominent impurities were 4ꢀAD (1), trione (19) as
well as hydroxyꢀ and perhydroxysteroids 20-23. Nevertheless, maintaining the pH at 6ꢀ7 and
temperature <35 °C ensured an efficient biotransformation and minimised the reverse
isomerisation. The content of 4ꢀAD (1) present in starting 5ꢀAD (5) material did not increase by
more than 2ꢀ3%. Sparging of the bioreduction solutions with nitrogen was necessary to keep the
level of the oxygenated impurities below 8% from the initial 4ꢀ6% present in the input crude 5ꢀ
AD (5).
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In addition, we observed the formation of some impurities specific to the bioreduction itself.
We detected low levels of 5ꢀADꢀdiol (6, <0.5% by HPLC area), possibly arising from reduction
at C17 catalysed by the Sphingomonas wittichii ketoreductase and/or other endogenous, nonꢀ
selective E. coli derived ketoreductase enzymes present in either the ketoreductase or glucose
dehydrogenase preparations. This impurity could be minimised by monitoring conversion and
stopping the reaction as soon as it reached completion.
Molecular biology and fermentation. The 1st generation ketoreductase preparation used in
the screen and the initial process development had relatively low activity, 17ꢀ35 U/g of enzyme
powder, as it contained a large amount of insolubly expressed protein. Consequently, high
enzyme loadings (3ꢀ5% w/w, 2U per g of substrate) were required to achieve full conversion of
5ꢀAD (5). Therefore, we decided to improve the protein expression, fermentation and
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downstream processing of the enzyme. This resulted in 10ꢀ15 fold improved enzyme
productivity, lowering enzyme loadings, which also resulted in better process parameters
(elimination of the emulsions we observed with the first generation enzyme preparations). It is
worth noting that such improvements in biocatalyst production can be very effective in
improving the efficiency of a biotransformation and are rapidly implemented.39
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Firstly, we optimised the gene encoding the ketoreductase for heterologous expression in a
phageꢀresistant E. coli BL21(DE3) strain containing pET26b vector. This afforded an isopropyl
βꢀDꢀ1ꢀthiogalactopyranoside (IPTG)ꢀinducible E. coli strain, in which 90% of the ketoreductase
was solubly expressed. Secondly, we developed high optical density fermentation (20L, glucose
feed, 176 OD, see Supplemental Information for details), which after clarification and
ultrafiltration (10 kDa membrane) resulted in 2nd generation enzyme produced as liquid cell free
extract of 39 U/mL activity.
Unfortunately, upon storage for 1 month at 2ꢀ8 °C we observed that the enzyme preparation
lost 75% of its activity and the later batches of the bioreduction required higher than expected
enzyme loading. We discovered that lyophilisation improved its storage and stability over time.
Consequently, later batches of the enzyme were lyophilised to give 3rd generation ketoreductase
powder with 400ꢀ500 U/g activity. This represented an overall 10ꢀ15 fold improvement in
comparison to the 1st generation biocatalyst (17ꢀ35 U/g) and it allowed a decrease in the enzyme
loading from 3ꢀ5% to 0.25ꢀ0.30% w/w to achieve 95ꢀ99% conversion in 6h. Not only did the
reduction in biocatalyst loading lower the cost of the biocatalyst, but it improved the ease of the
workup, as quick phase separation was observed after the biotransformation.
Purification of DHEA. Initially we intended to develop an efficient crystallisation of crude
DHEA (2) after the biotransformation step (79ꢀ91% purity by HPLC). A literature evaluation
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indicated that five polymorphs of DHEA (2) are known (forms IꢀV), as well as three
monohydrate polymorphs and numerous solvates with solvents like alcohols and ethers.40,41
Crystallisation studies of the crude DHEA (2) isolated from the biotransformation gave product
of >98% purity using a variety of solvents, but the recoveries were low (10ꢀ55%, results not
shown). Therefore, we decided to telescope the crude DHEA to the subsequent acetylationꢀstep
and focus on developing an efficient crystallisation of DHEA acetate (3) instead.
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2.3 Acetylation and final crystallisation
The acetylation of DHEA (2) to its acetate (3) proved to be a facile reaction and proceeded
quantitatively under a variety of reaction conditions tested. Successful reaction conditions
included: (i) dichloromethane/acetic anhydride/pyridine/DMAP; (ii) dichloromethane/acetic
anhydride/boron trifluoride diethyl etherate; (iii) acetic acid/acetic anhydride/sodium acetate; (iv)
toluene/acetic anhydride/triethylamine/DMAP (details not shown). In the early stages of
development DHEA acetate (3) was prepared using acetic anhydride, triethylamine with catalytic
DMAP in toluene.38 These conditions gave a fast, robust reaction with low cost raw materials
and acetate (3) was obtained in >98% purity (by HPLC) as 2 crops in 85ꢀ87% yield.
During further process development and optimisation work we aimed to reduce the number of
unit operations and to develop a facile acetylation procedure that could be telescoped into the
final crystallisation process. After some further optimisation we achieved full conversion of
DHEA (2) using acetic anhydride in the presence of sodium acetate in acetic acid over 16h.
Quench of the reaction into a mixture of water and methanol (9:1, 1.5 vol) precipitated crude
DHEA acetate (3) in 88% yield and 98% purity. This material was subsequently recrystallised
from methanol (4 vol), affording final product 3 in 74% yield and >99.5% purity (by HPLC),
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meeting the product specifications. These modifications considerably simplified the process and
provided both cost and operational benefits.
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Scheme 5. Optimised conditions for the >200 gꢀscale process to covert 4ꢀAD (1) to DHEA
acetate (3).
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(i) tBuOK (2.1 eq), tBuOH (6 vol), 30ꢀ35 °C, N2, 20 min; (ii) 2.5% AcOH in water (20 vol),
20ꢀ30 °C, N2, 60 min, 94% yield, 93% pure; (iii) 100 g/L, EtOAc:phosphate buffer (40 mM, pH
6.5) 60:40 v/v, NAD+ (0.075% w/w), glucose (1.1 eq), glucose dehydrogenase (0.044% w/w),
ketoreductase from Sphingomonas wittichii (liquid preparation, 2 U per g of 5ꢀAD), temp. 33 °C,
29h, N2, >99% conversion, 91% purity (iv) 2.0 eq Ac2O, AcOH (3 vol), NaOAc (1.25 eq), 60 °C,
N2, 16h, quench with MeOH: water (1.5 vol); (v) crystallisation from hot MeOH (4 vol), 74%,
purity >99.5% (by HPLC). 64% overall yield.
3. Summary
We developed a novel, high yielding chemoenzymatic route to DHEA acetate (3) from 4ꢀAD
(1). The key enabling reaction of this process was a highly regioꢀ and stereoselective
bioreduction of 5ꢀAD (5) to DHEA (3). A fast colourimetric assay was crucial in the
identification of a shortꢀchain dehydrogenase from Sphingomonas wittichii, which possessed the
desired selectivity. The enzyme displayed good robustness and performance under the biꢀphasic
reaction conditions,38 which enabled the biotransformations to be conducted at high substrate
concentrations (up to 150 g/L). Optimisation of the gene expression as well as the fermentation
conditions of an E. coli host strain resulted in 10ꢀ15 fold improvement of the enzyme
productivity and gave us access to a costꢀeffective biocatalyst. Development of both the 4ꢀAD to
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5ꢀAD isomerisation and the final acetylation and purification provided a robust process, suitable
for further scaleꢀup. Identification of the impurities generated in the process enabled the
development of control strategies and was crucial to obtain high purity final product.
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4. Experimental
Materials and methods
4ꢀAD (1) was supplied by Dr. Reddy's de Mexico, CuernavacaꢀCuautla, 62578 Civac Jiutepec
Morelos, Mexico. Testosterone (9) was obtained from Sigma. Epitestosterone (10) was obtained
from LGC Standards, Queens Road, Teddington, Middlesex, TW11 0LY, UK. 3β,17βꢀ5ꢀADꢀ
diol (6) was provided by Dr Reddy’s Laboratories Ltd., Custom Pharmaceutical Services, TDCꢀ1
Bollaram Road, Miyapur, Hyderabad 500049, Telangana, India. 3αꢀDHEA acetate (14) was
obtained from Bujno Synthesis, Dorodna 16, Warsaw, Poland. 3αꢀDHEA (13) and 3αꢀDHEA
acetate (14) were also synthesised from 3βꢀDHEA (2) using a Mitsunobu reaction sequence
according to a previously reported method42 (See Supplemental Info for details). All enzymes
used in the screen were proprietary to Dr Reddy’s Laboratories or obtained from commercial
suppliers (Almac, Codexis, Daicel, Evocatal, Johnson Matthey, Prozomix and Syncore). Glucose
dehydrogenase GDH CDXꢀ901 (66 U/mg) was supplied by Codexis. Nicotinamide cofactors
NAD+ and NADP+ were obtained from Europa Bioproducts. Routine NMR spectra were
1
recorded on a Bruker DRX 400MHz spectrometer and chemical shifts for H spectra were
referenced relative to internal tetramethylsilane or residual solvent peaks. If additional resolution
and sensitivity was required, compounds were analysed on a 700 and 500 MHz NMR instrument
with a cryoprobe, School of Chemistry, University of Cambridge, Lensfield Road, Cambridge
CB2 1EW, UK.
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Biology and fermentation: For detailed information of the methods, please see the
Supplemental Information.
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Analytical methods
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The isomerisation and biotransformations were monitored by HPLC: Method A using ELSD
and UV (λ = 200 nm) detectors and Luna C18 column 150 × 4.6 mm, column temp. 50 °C, flow
1 mL/min, MP 50:50 A:B, A: water, B: MeOH:MeCN 30:70, inj. vol: 10 µL. The progress of the
biotransformation reactions was also monitored by the base consumption as well as by 1H NMR
spectroscopy. LCꢀMS data was collected using ESI(+) ion detector and HPLC method A, where
0.1% v/v was added to water.
The purity of the DHEA acetate (3) was determined by HPLC: Method B using UV detector
(λ = 205 nm) and Kinetex C8 column 2.6 µm 100Å 150 × 4.6mm, flow 1 mL/min, column temp.
40 °C, using mobile phase gradient (A = 80:20 water:MeCN, B = MeCN, see Table 2), inj. vol:
10 µL.
Table 2. Mobile phase gradient for HPLC analysis of DHEA acetate (3) using Kinetex C8
column (Method B).
Time (min)
0
5
20
35
65
25
10
90
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5
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5
55
65
35
60
65
35
Mobile phase A
Mobile phase B
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65
35
95
95
Enzyme activity assay. The activity of the ketoreductase from Sphingomonas wittichii was
determined in 5 gꢀuse tests or spectrophotometrically using 4ꢀheptanol as a substrate, by
following its oxidation at 340 nm, monitoring the conversion of the NAD(P)+ cofactor to
NAD(P)H. One unit of activity is defined as sufficient enzyme to catalyse the oxidation of 1
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ꢁmol/min of 4ꢀheptanol under the assay conditions (see the Supplemental Information for
details).
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Colourimetric enzyme screening using INT/diaphorase
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In a spectrophotometric 96ꢀwell plate ketoreductase (0.1ꢀ0.6 mg/mL), diaphorase (0.1ꢀ0.6
mg/mL), NAD+ (0.4 mg/mL) and NADP+ (0.4 mg/mL) were reꢀhydrated with a solution of INT
in deionised water (1 mg/mL, 190 µL). 10 µL of a DMSO solution of the respective
hydroxysteroid (20 mg/mL stock solution of 2, 6 or 9) was added to commence the reaction. The
plates were sealed and incubated in a Heidolph shaker at 30 °C, 750 rpm and they were visually
inspected after 1h, 5h and 24h.
Biotransformation reactions (optimisation studies) - General procedure:
In a Multimax reactor vessel 5ꢀAD (5, 0.3ꢀ3.0 g) was preꢀdissolved in water immiscible coꢀ
solvent (5ꢀ25 mL, MTBE, 2ꢀMeTHF, EtOAc). Buffer cocktail (10ꢀ25 mL) containing
ketoreductase (1ꢀ3% w/w), glucose (1.1ꢀ2.0 equivalents), NAD+ (7ꢀ10 mg) and NADP+ (7ꢀ10
mg) and glucose dehydrogenase (CDXꢀ901, 3ꢀ10 mg, 66 U/mg) in phosphate buffer (50 mM, 10ꢀ
25 mL) was added and the reaction mixtures were stirred for 21h at 20ꢀ50 °C. The reaction pH
(5.5ꢀ8.5) was maintained by autotitration with 2 M NaOH or 1.7 M sodium carbonate solution.
The products were analysed by HPLC and NMR spectroscopy.
Chemistry
The initial reaction conditions were performed as described previously.38 The optimised
process was performed as described below.
Preparation of 5-androsten-3,17-dione (5)
Tertꢀbutanol (2.00 L) was sparged with nitrogen for 90 min at 30ꢀ35°C. Potassium tertꢀ
butoxide (235 g, 2.09 mol) was added and the mixture was stirred with nitrogen sparging for a
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further 90 min at 30ꢀ35 °C. 4ꢀAD (1, 300 g, 1.00 mol) was added and the reaction was stirred for
a further 1h with nitrogen sparging. Separately, acetic acid (150 mL) was added to a 20 L vessel
containing water (6.00 L) that had been sparged with nitrogen for 24 h and the mixture was
stirred at 20 °C under nitrogen. The reaction mixture was then slowly added to the acetic acid
solution via cannula over 10ꢀ15 min. Once the addition was complete, the mixture was cooled to
12 °C and stirred for 1h. The precipitate obtained was filtered, washed with water (3 × 2.00 L)
and dried under vacuum at 40 °C. The product is obtained as a free flowing white powder (283 g,
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94% yield), purity by HPLC: 93%, 4ꢀAD (1) ꢀ 3%, oxygenated impurities (19-23) – 4%). H
NMR (CDCl3, 400 MHz)
δ 5.38 (d, 1H), 3.28ꢀ3.33 (m, 1H), 2.86 (dd, 1H), 2.45ꢀ2.50 (2H), 1.05ꢀ
2.60 (m, 15H), 1.22 (s, 3H), 0.92 (s, 3H).
Preparation of dehydroepiandrosterone (2)
5ꢀAD (5, 8.0 g, 27.9 mmol, 94% pure by HPLC, 2% 4ꢀAD, 4% impurities 19-23) was
dissolved in EtOAc (40 mL) at 33 °C under nitrogen atmosphere and stirred vigorously. To this
was added a solution of phosphate buffer (50 mM, 40 mL, pH 6.3) containing glucose (6.0 g,
33.3 mmol, 1.2 eq), NAD+ (15 mg), glucose dehydrogenase (CDXꢀ901, 8 mg, 528 U) and
ketoreductase powder (3rd generation, 20 mg, 0.25% w/w, 9 U) was added. The reaction
temperature was maintained at 32.5 °C (jacket temp 33 °C), while the pH 6.3ꢀ6.5 was kept
constant by addition of 1.5 M K2CO3 solution in water. The reaction conversion was monitored
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by base consumption as well as by HPLC and H NMR spectroscopy. After 6h the reaction
reached ~95% conversion. It was stirred for a total of 21 h to give >99% conversion. The
mixture was warmed up to 70 °C to fully dissolve partially precipitated DHEA and the phases
were separated. The separated organic layer was dried over magnesium sulfate and evaporated to
give crude DHEA as solid residue (7.2 g, 24.2 mmol, 90% yield). Purity by HPLC: 93% (5ꢀAD
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(5, 0.5%), 4ꢀAD (1, 1.9%), 5ꢀADꢀdiol (6, 0.5%)). H NMR (CDCl3, 400 mHz) δ 5.38 (d, 1 H),
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3.50ꢀ3.58 (m, 1 H), 2.47 (dd, 1 H), 2.22ꢀ2.39 (m, 2H), 2.04ꢀ2.12 (m, 2H), 1.92ꢀ1.98 (m, 1 H),
1.83ꢀ1 .89 (m, 3H), 1 .44ꢀ1 .71 (m, 6H), 1.22ꢀ1.29 (m, 2H), 0.97ꢀ1.14 (m, 2H), 1.04 (s, 3H), 0.89
(s, 3H).
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Preparation of DHEA acetate (3) from 5-AD (5) on 200 g-scale (telescoped process)
5ꢀAD (5, 235 g, 0.82 mol) was loaded into a 5 L reactor and sparged with nitrogen for 30 min
upon gentle stirring. EtOAc (1.15 L) was added and the mixture was stirred at 33 °C under
nitrogen until the substrate was fully dissolved. In a separate flask KH2PO4 (7.85 g) and glucose
(162 g, 0.90 mol) were dissolved in 1.65 L of deionised water at 20 °C and the pH was adjusted
to 6.3 using solid KOH (1.40 g, ≥85% assay). The solution was sparged with nitrogen for 2ꢀ5h.
NAD+ (0.20 g), glucose dehydrogenase (CDXꢀ901, 165 mg, 10.9 kU) and ketoreductase cell free
extract (2nd generation, 4.0 mL, 40 U) were added and stirred until fully dissolved. The buffer
solution of enzymes was added to the vigorously stirred solution of 5ꢀAD in EtOAc at 32.5 °C.
The reaction was stirred at 32.5 °C and pH 6.3ꢀ6.5 was maintained by autotitration with 1.7 M
Na2CO3 solution in water. The reaction conversion was monitored by base consumption as well
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as by H NMR spectroscopy. After 4 h the reaction reached 43% conversion and additional
portions of ketoreductase (3 mL, 30 U), NAD+ (300 mg) and GDH (160 mg, 10.9 kU) were
added; after 8 h more ketoreductase (4 mL, 40 U), NAD+ (300 mg) and GDH (160 mg, 10.9 kU)
were added. The mixture was stirred for 22 h and reached full conversion. The reaction mixture
was warmed up to 55 °C to fully dissolve the partially precipitated DHEA and the phases were
separated. The aqueous layer was extracted with EtOAc (1 L) at 55 °C. The combined organic
layers were washed with brine (0.4 L), water (0.4 L) and passed through a magnesium sulfate
plug, washed with additional EtOAc (0.4 L). The solvent was partially evaporated under vacuum
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(100 mbar/ 50 °C water baths) to give 643 g of DHEA (2) in EtOAc, which solidified. Sample
analysis indicated 36% potency by HPLC, equivalent to 0.80 mol, yield 97% (91% purity by
HPLC, 4ꢀAD – 7%, oxygenated impurities – 2%). This material was suspended in acetic acid
(400 mL) and the residual EtOAc was removed under reduced pressure. Additional 320 mL of
acetic acid were added (720 mL total, 3 vols), acetic anhydride (150 mL, 1.60 mol) and sodium
acetate (80.0 g, 0.98 mol). The reaction was stirred at 60 °C for 16 h under a nitrogen
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atmosphere. Upon completion (by H NMR) the reaction was cooled to 25 °C and a mixture of
water (1125 mL) and methanol (125 mL) was added to the reaction mixture over 10 min and it
was stirred for 1h. The precipitate was filtered and washed with further 3:1 water/methanol
solution (1000 mL). The wet cake (320 g) was dissolved in hot methanol (1000 mL) and this
solution was allowed to cool to 20 °C, then cooled further in an ice bath. The solid was collected
and washed with additional methanol:water mixture (1:2, 300 mL). The solid was placed in a
drying oven at 40 °C overnight to give DHEA acetate (3) as a white solid (171.4 g, 74% yield for
1
2 steps, and purity 99.5% by HPLC). H NMR (CDCl3, 400 MHz)
δ 5.41 (d, 1H), 4.57ꢀ3.65 (m,
1H), 2.46 (dd, 1H), 2.28ꢀ2.36 (m, 2H), 2.04 (s, 3H), 1.80ꢀ2.20 (m, 6H), 0.95ꢀ1.75 (m, 10), 1.05
(s, 3H), 0.89 (s, 3H).
ACKNOWLEDGMENT
We acknowledge Pieter de Koning for invaluable discussions and his help in preparation of this
manuscript.
ASSOCIATED CONTENT
Supporting Information. Synthesis and characterisation of the main process impurities;
structural assignment data for the oxygenated compounds 19-23; experimental for molecular
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biology and fermentation; enzyme activity assay and sample chromatograms. This material is
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available free of charge via the Internet at http://pubs.acs.org.
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AUTHOR INFORMATION
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Corresponding Author
* email: keholt.tiffin@drreddys.com
Notes
The authors declare no competing financial interest.
Present Addresses
¶
Department of Process Research, Merck Research Laboratories Merck & Co., Inc., P.O. Box
2000, Rahway, New Jersey 07065, USA
ǂ AstraZeneca, 310 Cambridge Science Park, Milton Road, Cambridge CB4 0FZ, UK
‡‡
Zydus Cadila Ltd, SarkhejꢀBavla N. H. No. 8A, Moraiya Tal: Sanand, Dist: Ahmedabadꢀ
382210, India.
§§ CordenPharma International GmbH, Otto Hahn Strasse 1, 68723 Plankstadt, Germany
¶¶ Inogent Laboratories Private Ltd, 28A, IDA Nacharam, Hyderabadꢀ 500 076, India
††Almac Group, 25 Fretz Road, Souderton, PA 18964, USA
Author Contributions
The manuscript was written by Anna Fryszkowska through equal contributions of all authors.
All authors have given approval to the final version of the manuscript.
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ABBREVIATIONS
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DHEA − dehydroepiandrosterone, 5ꢀAD − androstꢀ5ꢀeneꢀ3,17ꢀdione, 4ꢀAD − androstꢀ4ꢀeneꢀ
3,17ꢀdione, HSD – hydroxysteroid dehydrogenase, KSI – ketosteroid isomerase, NAD+ −
nicotinamide adenine dinucleotide (oxidised form), NADH − nicotinamide adenine dinucleotide
(reduced form), NADP+− nicotinamide adenine dinucleotide phosphate (oxidised form), NADPH
− nicotinamide adenine dinucleotide phosphate (reduced form), GDH − glucose dehydrogenase,
IPTG − isopropyl βꢀDꢀ1ꢀthiogalactopyranoside.
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