NAD+-Dependent Enzymatic Route for the Epimerization of Hydroxysteroids

Article abstract of DOI:10.1002/cssc.201801862

Epimerization of cholic and chenodeoxycholic acid (CA and CDCA, respectively) is a notable conversion for the production of ursodeoxycholic acid (UDCA). Two enantiocomplementary hydroxysteroid dehydrogenases (7α- and 7β-HSDHs) can carry out this transformation fully selectively by specific oxidation of the 7α-OH group of the substrate and subsequent reduction of the keto intermediate to the final product (7β-OH). With a view to developing robust and active biocatalysts, novel NADH-active 7β-HSDH species are necessary to enable a solely NAD⁺-dependent redox-neutral cascade for UDCA production. A wild-type NADH-dependent 7β-HSDH from Lactobacillus spicheri (Ls7β-HSDH) was identified, recombinantly expressed, purified, and biochemically characterized. Using this novel NAD⁺-dependent 7β-HSDH enzyme in combination with 7α-HSDH from Stenotrophomonas maltophilia permitted the biotransformations of CA and CDCA in the presence of catalytic amounts of NAD⁺, resulting in high yields (>90 %) of UCA and UDCA.

Full text of DOI:10.1002/cssc.201801862

Accepted Article
Title: NAD+-dependent enzymatic route for the epimerization of
hydroxysteroids
Authors: Fabio Tonin, Linda G. Otten, and Isabel Arends
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10.1002/cssc.201801862
ChemSusChem
NAD+-dependent enzymatic route for the epimerization of
hydroxysteroids
Fabio Tonin[a], Linda G. Otten[a] and Isabel W.C.E. Arends[a,b],
*
[a] Department of Biotechnology, Delft University of Technology, Van der Maasweg 9, 2629 HZ
Delft, The Netherlands.
[b] Present address: Faculty of Science, Utrecht University, Budapestlaan 6, 3584 CD Utrecht,
The Netherlands.
Email: I.W.C.E.Arends@tudelft.nl
* Corresponding author
1 Abstract:
The epimerization of cholic and chenodeoxycholic acid (CA and CDCA, respectively) is a
notable conversion for the production of ursodeoxycholic acid (UDCA). Two
enantiocomplementary hydroxysteroid dehydrogenases (7α- and 7β-HSDHs) can carry out this
transformation fully selectively by specific oxidation of the 7α-OH group of the substrate and
subsequent reduction of the keto intermediate to the final product (7β-OH). In our quest to
develop robust and active biocatalysts, we sought novel NADH-active 7β-HSDH’s, allowing us
to carry out a solely NAD+-dependent redox-neutral cascade for UDCA production. In this
research a wild-type NADH dependent 7β-HSDH from Lactobacillus spicheri (Ls7β-HSDH) was
identified, recombinantly expressed, purified and biochemically characterized. Using this novel
1
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NAD+ dependent 7β-HSDH enzyme in combination with 7α-HSDH from Stenotrophomonas
maltophilia, permitted the biotransformations of CA and CDCA in the presence of catalytic
amounts of NAD+, resulting in high yields (>90%) of UCA and UDCA.
2 Keywords:
Bile acids; Biotransformation; Hydroxysteroid dehydrogenases; UDCA; nicotinamide cofactor
switching.
2
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3 Introduction
Ursodeoxycholic acid (UDCA) is an active pharmaceutical ingredient (API) used in clinical
therapy for the treatment of cholestatic diseases[1]. It is a secondary bile acid that, as reported in
several papers and reviews, shows several pharmacological effects[2, 3]. Particularly, UDCA
dissolves cholesterol gallstones, acting as a surfactant[4, 5]. Additionally, other functions and
influences on human physiology have been discovered: for example, UDCA improves the liver
function in cholestatic diseases through secretion of glutathione[6, 7] and interaction with several
receptors[8, 9], and significantly decreases cholesterol saturation in the bile[10, 11]. Recently, other
studies demonstrated anti-tumorigenic effects and a lower risk for some cancers[12]. In
comparison to chenodeoxycholic acid (CDCA) which is also used in therapies, UDCA provides
total absence of side effects maintaining a high efficacy, in terms of pharmacological effects[13,
14]
.
This API has been known in Chinese traditional medicine for many years, but natural UDCA can
only be obtained by isolation from bear bile, making it very expensive[15]
.
The current viable route for this API consists of chemical transformation of cholic acid (CA) or
CDCA. These bile acids are obtained from the cheaper and more available bovine bile, that
represents, in terms of industrial feasibility, the only source of hydroxysteroids. Currently, CA is
chemically transformed into UDCA by a seven step chemical synthesis[16, 17]. Unfortunately this
route, first proposed by Hofmann[16], requires the employment of toxic and dangerous reagents
(eg. hydrazine, Cr2O, pyridine) and results in large amounts of waste. In addition, it only has an
overall yield of about 30%. In order to improve the efficiency of UDCA synthesis, alternative
chemical routes have been proposed. Dangate[18] for example achieved the transformation of CA
into UDCA (53% yield using o-iodobenzoic acid, hydrazine and metallic sodium in n-propanol)
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without the protection and de-protection steps used in the original route. However, this process is
still not optimized in terms of costs and environmental impact. UDCA can also be obtained by
epimerization of CDCA (intermediate in the synthesis of CA). In the latter case a viable
production route would need to be complemented by a novel synthetic route for CDCA, because
of the low natural availability of this compound.[19]
An emerging technology for the modification of hydroxysteroids is the use of enzymatic
transformations[20, 21]. The regiospecificity of enzymes offers the possibility to avoid protection
steps of the hydroxyl groups not employed in the transformation. This approach was followed by
several research groups: attention was focused on the identification of enzymes able to play a role
in this synthesis. In an enzymatic process for the production of UDCA from its precursors,
several steps have to be performed. These are hydrolysis/deconjugation of glycine and taurine
CA derivatives (used as raw starting material), the stereoinversion of the hydroxyl functions (by
oxidation and subsequent reduction) and the specific hydroxylation and dehydroxylation of
desirable positions in the steroid rings[20]
.
The epimerization reaction (stereoinversion of 7α-OH group), can be performed by two enzymes
(7α- and 7β- hydroxysteroid dehydrogenase (HSDH)): the first enzyme specifically oxidizes the
7α-OH group to the ketone, that is further reduced by the second enzyme to the corresponding
alcohol group with inverted stereochemistry (β-OH). HSDHs are enzymes of the class of alcohol
dehydrogenases (ADH) that use NAD(P)+ and NAD(P)H as electron acceptor and donor,
respectively. In literature, several cascades that use these enzymes are reported[22-27]: however,
since known enzymes have different cofactor specificities, i.e. 7α-HSDHs (NAD+ dependent) and
7β-HSDHs (NADPH dependent) the oxidative and reductive reaction steps are decoupled in
currently reported procedures (Figure 1 – state of art). The cofactors, in this case, are regenerated
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by the addition of two other enzymes and two sacrificial substrates[21-23]. In this specific case, the
reaction is driven towards completion by the addition of sacrificial substrate (in great surplus):
Although good yields were achieved (>80%), large amounts of side-products and waste were
formed that have to be removed during the downstream processing steps, increasing the overall
cost and the environmental burden of UDCA production.
Pedrini et al., already in 2006,[28] reported the successful epimerization of CDCA to UDCA using
a catalytic NAD+-dependent cascade reaction, with two NAD+ dependent dehydrogenases
isolated from Xanthomonas maltophilia CBS 897.97 (recently reclassified as Stenotrophomonas
maltophilia). In this way the requirement of external systems for cofactor regeneration was
circumvented and UDCA was obtained with a final yield of 75%. However, the cultivation of this
class 2 safety level microorganism is complex[29] (microaerophilic conditions have to be used)
and the 7α-HSDH and 7β-HSDH enzymes were only produced in low amounts (4 and 0.2
mg/Lculture, respectively). In addition, the amino acid or DNA sequences of these enzymes were
not reported, making the recombinant expression of these proteins not possible. Since the activity
and cofactor specificity is not clear from sequence alone, the univocal identification of the target
[23, 24, 30, 31]
corresponding gene in genomes databases is not possible. In addition, other works
have been reported on the employment of a NADP+ dependent redox neutral cascade for CDCA
epimerization: however, in comparison to NAD+, the use of NADP+ is non-optimal because of
the higher price, lower stability[32] and lower natural availability. Generally, by the employment
of a redox neutral cascade, 80% of UDCA is produced.
In this paper we report the development of an efficient enzymatic cascade, using well defined and
easy to produce enzymes, for the production of UDCA and UCA from CDCA and CA,
respectively. A full NAD+ mediated cascade, that circumvents the need for cofactor regeneration
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was set-up resulting in conversion above 90%. The key discovery in this paper is the protein
engineering of the NADP+ dependent 7β-HSDH from Clostridium sardiniense that led to the
identification of key residues responsible for NAD/NADP cofactor recognition. With this new
sequence at hand, the natural NAD+ dependent enzyme from Lactobacillus spicheri, was
disclosed for the first time as being highly active in the reduction of the 7-oxo-derivatives of CA
or CDCA to either UCA or UDCA.
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4 Results and discussion
4.1 Protein expression and purification
The genes coding for the Sm7α-HSDH, Cs7β-HSDH and Ls7β-HSDH were cloned into
pET24d(+) plasmid, yielding enzymes containing a C-terminal 6x His-tag. The recombinant
enzymes were produced in E. coli BL21(DE3) host cells grown at 37 °C in LB medium. IPTG
was added at the late exponential phase of growth and the cells were collected after another 18 h
of incubation at 25 °C while shaking. The expression level under these conditions of the different
proteins is reported in Table 1 and Supplementary Table 2. The His-tagged enzymes were
purified by HiTrap chelating chromatography: all of the enzymes were isolated with ≥ 95%
purity, as judged by SDS-PAGE analysis (Supplementary Figure 2).
4.2 Mutagenesis of Cs7β-HSDH
Since the three-dimensional structure of 7β-HSDH from C. sardiniense is unknown, a model of
the protein was built by using the comparative homology-modeling server SWISS-MODEL[33]
.
7β-HSDH from Collinsella aerofaciens (PDB code 5GT9) was chosen as template because of the
high sequence identity (42%) with Cs7β-HSDH[34]. The binding mode of the co-substrate NADP+
in the model of the Cs7β-HSDH active site was analyzed: the ribose bound phosphate group,
interacts with two arginine residues (R40 and R41) (Figure 2a).
On the basis of the in silico analysis, site-saturation mutagenesis (SSM)[35] was performed at
positions 39, 40 and 41 using the QuikChange kit and the wild-type Cs7β-HSDH cDNA as
template. In the first round of mutagenesis an aspartate was specifically introduced in position 39
while positions 40 and 41 were more randomly changed. The use of a DHK codon in these
positions allows the introduction of 13 different amino acids (A, D, E, F, I, K, L, M, N, S, T, V
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and Y) with low repetition, representing different chemical functionalities and steric
encumbrances[36]. The use of this codon drastically reduces the number of clones that have to be
screened. The activity of Cs7β-HSDH variants on NAD+ as co-substrate was screened on a
microtiter plate using a spectrophotometric method (production of NADH, measured at 340 nm)
and an automated liquid-handling system. For the first round of mutagenesis, (G39D, R40X and
R41X), 769 clones were screened, giving a probability of 91% that every combination of amino
acids is measured. The clones most active on NAD+ as identified through the screening procedure
were isolated and the substitutions were identified by DNA sequencing.
After the first round of mutagenesis and screening most of the enzyme variants obtained are
inactive or have an activity lower than the control activity of wild-type Cs7β-HSDH on NAD+(~1
mU mL-1 under these conditions). However, four Cs7β-HSDH variants were isolated that showed
a clear activity on NAD+: G39D, R40L, R41N (DLN variant); G39D, R40F, R41F (DFF variant);
G39D, R40K, R41S (DKS variant) and G39D, R40K, R41V (DKV) variant. From the molecular
modelling of these variants it could be observed that the hydroxyl group of threonine in position
17 may interfere with aspartate 39 in the hydrogen binding of the free OH-group of ribose
(Figure 2b). A second pair of degenerated primers were designed in order to mutate that
threonine (T17X). For this second round of mutagenesis, all four variants were used as template
for the mutagenesis and 380 clones (96 clones for each variant) were screened (99% coverage).
Out of all screened mutants, one variant (T17A, G39D, R40L, R41N – ADLN variant) showed a
very high activity towards NAD+ as cosubstrate in the microtiter plate assay employing cell
extract (Figure 2c).
The ADLN variant was expressed in E. coli BL21(DE3) cells and purified by HiTrap chelating
chromatography (>90 % purity). This variant shows an expression yield similar to that of the
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wild-type Cs7β-HSDH (in terms of purified protein/liter of fermentation broth, see
Supplementary Table 2). The Michaelis-Menten kinetics of all selected enzyme variants were
measured using NAD+ or NADP+ as cofactor.
As shown in Table 2, among the Cs7β-HSDH variants obtained by SSM, the ADLN was
identified because of a thirty-fold increase in specific activity compared to wild-type enzyme.
In order to further increase the activity on NAD+, the T17A, E18D, G39D, R40L, R41N
(ADDLN) variant was designed and recombinantly expressed: the substitution of the glutamate
with the smaller aspartate in position 18 may result in the formation of a second hydrogen bond
between the cofactor and the protein (Figure 2d). The comparison of the kinetic parameters of the
ADLN and ADDLN variants of Cs7β-HSDH indicates that the addition of a second hydrogen
bond increases the affinity of the ADDLN variant for the NAD+ (8-fold decrease in Km), see
Table 2. However, the specific activity on NAD+ of this variant decreases (10-fold lower in
comparison to previous isolated one). In order to restore the specific activity on NAD+ a fourth
round of SSM was carried out employing the same primers used in the first round and the
ADDLN variant as template. The T17A, E18D, G39D, R40A, R41A variant (ADDAA) was
isolated. The specific activity under standard conditions of this variant is 0.3 U mg-1 (Table 2).
Interestingly the affinity for the cofactor is lower than the one observed in the previous variants.
However, the expression level of this protein is fifteen-fold higher than the wild-type Cs7β-
HSDH (486 vs. 20 mg/Lculture), resulting in the higher activity under the screening conditions.
Overall, we conclude that ADLN is the best obtained variant for our purpose due to its highest
catalytic efficiency.
4.3 New 7β-HSDH from Lactobacillus spicheri
With the new sequence of NADH dependent 7β-HSDH (ADLN and ADDLN variants) in hand, a
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new alignment search was carried out in order to find potentially better, previously non-identified
enzymes, able to convert CA and its derivatives. The Ls7β-HSDH sequence was identified using
the Basic Local Alignment Search Tool (BLASTp): the predicted sequence analysis showed a
792 bp ORF corresponding to a protein of 264 amino acids residues.
This enzyme was identified as a putative NADH dependent 7β-HSDH based on the fact that the
amino acids relevant to the binding and recognition of NADH are present: specifically, the
alanine and aspartate in position 18 and 19 and the stretch DYS in position 40-42 are
conserved[37], analogous to the situation in analogue Cs7β-HSDH (Figure 3b). The predicted Mw
of 29 kDa and the predicted homodimeric quaternary structure (Figure 3a) put Ls7β-HSDH in the
short chain dehydrogenase/reductase superfamily.
4.4 Biochemical characterization of HSDHs
Sm7α-HSDH showed a strict NAD+ activity on both CDCA and CA, although the activity on CA
is considerably lower (halved). No activity was detected when NADP+ was used as electron
acceptor. Sm7α-HSDH displayed a Km of 0.22 and 0.96 mM for CDCA and CA, respectively.
The Sm7α-HSDH did not show any substrate inhibition on CA, but a Ki of 11 mM on CDCA was
measured. The Km value for NAD+ is 0.55 mM (Table 3) which is similar to earlier reported
enzymes.
The pH and temperature dependence of Sm7α-HSDH activity was investigated: both the
maximum activity and stability occurred at slightly alkaline pH values (Figure 4a). The enzyme is
quite thermophilic, showing an optimum at around 70 °C (Supplementary Figure 3a), and is quite
stable: after 24 h incubation at 25 and 37 °C, the enzyme maintained ca. 100 and 70% of its
initial activity, respectively. Incubations at higher temperatures (60 °C) resulted in a complete
loss of enzymatic activity. The enzymatic activity of Sm7α-HSDH was also investigated in the
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presence of different concentrations of methanol, that could be used as a potential co-solvent in
order to increase the solubility of hydroxysteroids in water. The enzyme shows no loss of activity
in 10% methanol and it conserves 90% of its activity in 20% methanol (Figure 5a).
On the other hand, wild-type Cs7β-HSDH showed a strict NADP+ activity on UDCA (0.74 U mg-
under standard conditions). The activity on NAD+ is roughly 100-fold lower showing a Km of
1
2.6 mM and a specific activity of 0.023 U mg-1. Cs7β-HSDH displayed a Km of 0.16 mM for
UDCA (Table 4). The pH and temperature dependence of Cs7β-HSDH activity was investigated:
both the maximum activity and stability occurred at slightly alkaline pH values (Figure 4b).
Notably, the pH optimum for the reduction reactions was detected at pH 6-7. In the presence of
NADPH, this enzyme is able to reduce both 7-oxo-DCA and 7-oxo-LCA (see Table 4).
In general, the enzyme is less thermophilic than the Sm7α-HSDH, showing an optimum at around
60 °C (Supplementary Figure 3b), and less stable: after 24 h incubation at 25 and 37 °C, Cs7β-
HSDH maintained ca. 85 and 62% of its initial activity, respectively. Cs7β-HSDH shows a good
tolerance to concentrations of methanol above 10% and 70% of its activity is conserved in
presence of 20% methanol.
The ADLN, ADDLN and ADDAA variants showed an increase in activity towards NAD+ and
NADH as cosubstrate, with little change in specificity for the different substrates (Table 4).
Interestingly, the isolated variants showed higher stability than the wild-type Cs7β-HSDH. From
this comparison, it can be observed that the ADLN variant maintains 95% of its activity after
incubation for 24h at 25 °C (compared to the 85% for the wild-type under the same conditions).
As predicted by in silico analysis, Ls7β-HSDH showed a strict NAD+ activity (3.10 U mg-1 in
standard condition). A weak activity was detected when NADP+ was used as electron acceptor
(Table 2). Ls7β-HSDH showed a 0.15, 0.04 and 0.13 mM Km for UDCA, 7-oxo-LCA and 7-oxo-
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DCA, respectively. Ls7β-HSDH is inhibited by UDCA showing a Ki of 0.8 mM for this substrate.
The Km value for NAD+ is 0.08 mM. From all the 7β-HSDH-variants in this paper, this enzyme
shows the highest NAD+/NADH activities for both the reduction and the oxidation of 7β-OH
hydroxysteroids and its derivatives. All kinetic data are presented in Tables 2 and 4.
From the biochemical characterization it can be observed that this enzyme showed similar
features to the other isolated enzymes, with the exception of the pH dependence: the maximum
activity for both oxidation and reduction reactions was observed at lower pH (6-7) (Figure 4f).
Ls7β-HSDH is quite thermophilic, showing an optimum at around 70 °C (Supplementary Fig.
4C), and it is stable at 25 and 37 °C, maintaining, after 24 h of incubation, ca. 98 and 72% of its
initial activity, respectively. Ls7β-HSDH conserves 60% of its activity in presence of 20%
methanol: thus for follow-up bioconversion studies 10% methanol was used, which limits the
amount of substrate that can be loaded in a biotransformation (Figure 5f).
4.5 Bioconversion of CDCA and CA
In order to assay the applicability of the enzymes described above as biocatalyst for the
production of UDCA and UCA in a batch bioreactor, lab-scale bioconversions were carried out:
10 μmol of CDCA in the presence of 1 μmol of NAD+ and 10% MeOH was converted into
UDCA (92% yield) in 200 min by 1 U of Sm7α-HSDH (0.23 μg) and 0.6 U of Ls7β-HSDH (190
μg) (Figure 6a). HPLC analysis showed no formation of undesired products, except for the 7-
oxo-LCA (0.5 μmol), intermediate of the enzymatic cascade (Supplementary Figure 4a).
Under the same conditions, 10 μmol of CA was converted into UCA (91%) in 120 min (Figure
6b).
Surprisingly the addition of MeOH (10-20%) resulted in an increase (10%) of conversion (Table
5): without cosolvent only 80% of UDCA can be obtained by this biotransformation.
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The effect of cosolvents on the equilibrium of the epimerization reaction was already discussed in
literature[38] but never observed for this specific biotransformation.
Additionally, in order to optimize the reaction conditions, a set of bioconversions were carried
out: as reported in Table 5, >90% of UDCA was observed employing 1 or 0.5 μmol of NAD+.
Biocatalytic conversions were also carried out in the presence of 0.2 μmol of NAD+ using CDCA
as substrate: in this case the conversion is slower, yielding 24% of UDCA in 20 h (Figure 6c).
Notably, here the detected amount of keto intermediate was lower than 1% (0.1 μmol).
Bioconversions were also tested at different pHs (6 and 7) but no improvements were observed
(after 150 min in presence of 1 mM NAD+, 77%, 81% and 87% conversion was observed at pH
6, 7 and 8, respectively). Unfortunately, the increase of substrate loading to 20 mM negatively
effects the bioconversion, yielding 20% of product after 12 h of incubation.
The ADLN variant of Cs7β-HSDH was also tested in the cascade reaction in presence of 1 mM
NAD+ for the epimerization of CDCA and CA: although no conversion was observed when
CDCA was used as substrate, 10 μmol of CA was converted into UCA (91% yield) by 790 μg of
enzyme after 30 min (Figure 6d and Supplementary Figure 4b). This behavior can partially be
explained by the kinetic parameters of this variant: Ki for the intermediate 7-oxo-LCA is 4 times
lower than the one for 7-oxo-DCA.
Notably, in the different bioconversions the amount of keto intermediate is limited by the amount
of cofactor used. Thus in a typical reaction where 10% cofactor was used, 5% keto intermediate
is formed as byproduct.
5 Conclusion
Epimerization of CA and its derivatives is a pivotal step in the synthesis of UDCA (see Figure 1).
Enzymatic epimerization circumvents the needs for protection/deprotection steps and can be
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carried out with high yield. A solely NAD+ dependent cascade for CDCA and CA epimerization,
in which NAD+ is present in catalytic amounts, represents a promising way towards a more
efficient process in terms of atom economy and number of reagents required.
In order to achieve that goal it is fundamental to employ robust, active and selective enzymes:
Sm7α-HSDH shows kinetic parameters better or comparable to its E. coli or Bacteroides
homologues, widely used by researchers in these kind of processes[22, 27, 39]. This enzyme
selectively oxidizes the 7α-OH group of CA and CDCA. In terms of stability it shows promising
tolerance to the presence of methanol and elevated temperatures.
Until now, all sequences of 7β-HSDHs reported in literature are NADP+ dependent enzymes[20]
.
However, protein engineering provides a straightforward method to change cofactor specificity:
this has been the subject of many researchers in both academia and industry for the past
decades[40-42]. Unfortunately, the resulting enzymes usually have a perfect cofactor switch, but in
several cases, the enzyme activity and/or substrate specificity decreases tremendously[43]. A
software for the prediction of the specificity-determining residues (CSR-SALAD[44, 45]) has been
developed and is now widely used[46], but there is still no comprehensive solution for cofactor
switching.
Following this approach, an iterative mutagenesis of Cs7β-HSDH was performed. A semi-
rational approach was pursued, based on the identification of the residues responsible for the
cofactor recognition discovered in the homology model of the enzyme.
In this way, residues G39, R40 and R41 were identified to be responsible for binding of the
phosphate group at the 2’-position of the ribose. Interestingly, the importance of arginine in the
binding of NAD(P)+ was shown before on the enantiocomplementary 7α-HSDHs[47].The amino
acid in position 39 was mutated to an aspartate. Notably, a similar attempt of cofactor switching,
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performed by Bakonyi and Hummel on the Clostridium difficile 7α-HSDH[48] has led to the
identification of similar variants (A37D). In our case, the free hydroxyl group at 2’-of ribose
forms a hydrogen bond with the aspartate (length 2.2 Å). R40 and R41 residues were randomly
mutagenized in order to identify a couple of amino acids that I) do not interfere with aspartate in
NAD+ binding and II) create a binding pocket for NAD+. SSM with limited codons helped to
introduce in the protein a certain degree of variability while generating smart libraries containing
<1000 clones for each round.
In the same way, T17 was subjected to SSM. From the 3D structure of the model it was observed
that the OH-group of the side chain is 2.5 Å from the aspartate. The obtained ADLN variant
shows a reasonable activity on NAD+ (in comparison with the wild-type enzyme on NADP+).
The Km of this variant for NAD+ is 2 mM, which is actually close to the value reported by Pedrini
et al., for their 7β-HSDH from S. maltophilia[28]. Additionally the mutation E18D improved the
affinity for NAD+ about 10-fold but unfortunately, the specific activity of the protein decreases.
We hypothesize that the E18 residue in the Cs7β-HSDH also has a structural role. From this
optimization round we thus concluded that the ADLN variant was the best mutant.
These data suggest that the efficiency and the selectivity in cofactor binding depends for a large
part, on the molecular dynamics of the Cs7β-HSDH. Although this feature can be reasonably
simulated, the engineering of the target enzyme is still complex[49]. For this reason, the attention
was focused on the identification of a wild-type enzyme that conserved both I) the structure of
Cs7β-HSDH and II) the residues identified in Cs7β-HSDH to be responsible for cofactor
recognition. The new analysis lead us to identify a sequence of a putative NAD+ dependent 7β-
HSDH from Lactobacillus spicheri. Ls7β-HSDH shows indeed a NAD+ dependence activity (Km
80 μM) and an activity higher than the Cs7β-HSDH on the respective cofactor. As shown, both
the ADLN variant of Cs7β-HSDH and the Ls7β-HSDH are suitable for the epimerization of CA
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(to UCA) but, only the Ls7β-HSDH is able to epimerize CDCA (to UDCA): this can partially be
explained by the comparison of the kinetic parameters of the two enzymes: firstly, the ADLN
variant, although it was engineered, shows a Km for the NADH 35-fold higher than the
Lactobacillus one; secondly the ADLN variant is inhibited by 7-oxo-LCA (Ki=2.0 mM). In
general, Ls7β-HSDH is a more interesting biocatalyst because of the high activity on both 7-oxo
derivatives.
Bioconversions were carried out at the substrate concentration used in previous studies (10 mM).
This choice was done in order to I) have a straight comparison between the reported systems and
the current redox neutral cascade and II) ensure solubility, because solubility of CDCA in a water
environment (pH 8.0) is low (around 10 mM as specify by Zheng[23]).
The developed cascade shows an epimerization yield >90% for both CA and CDCA under non-
optimized conditions. This shows that the currently developed set of enzymes and the
epimerization cascade belonging to it, provides a highly promising alternative for both chemical
and enzymatic synthetic procedures applied currently[20, 21]. This could provide a basis for a much
more elegant and atom efficient synthesis of UDCA from CDCA.
Since the epimerization of CA and CDCA is carried out with catalytic amounts of NAD+, the
shift of the equilibrium depends on the difference in physio-chemical properties between reagents
and products. Particularly, in a preliminary study using a molecular mechanics approach, we have
identified a difference in the ΔG0 between the two epimers of -1.23 kJ mol-1 (calculated at
standard conditions). In addition the differences in solubility and in critical micelles
concentration (CMC) of the two isomers can play a role in the shift of the equilibrium to the
products. Similar results were shown by Dangate[18] and Giovannini[50] in chemical and chemo-
enzymatic routes, respectively: Dangate et al. reported that 7-oxo-DCA is preferably reduced to
the 7β epimer (UCA:CA – 80%:10% yield) in presence of sodium in anhydrous n-propanol.
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Under the same conditions, Giovannini et al. showed similar results for the reduction of 7-oxo-
LCA (UDCA:CDCA – 82%:15% yield). Also in our case we observed that the reduction towards
the 7β epimer is thermodynamically favored, but differently from the chemical cascade, the
reversibility of the enzymatic catalyzed reactions allows to reach the thermodynamic equilibrium
between the two epimers. In comparison to the previously reported NADP+-dependent redox
neutral systems we observed an higher conversion yield (>90% vs. 80%. reported by Zheng[23]).
This increase can be explained by the use of the cosolvent (10% MeOH) that influences the
reaction environment (Table 5) without affecting the activity of the employed enzymes. Another
advantage of our system is given by the use of NAD+ instead of NADP+, that lowers the cost of
the required cofactor and shows higher stability in solution.
As previously observed by Zheng[23], in all the epimerization reactions, 5% of keto intermediate
(7-oxo-DCA or 7-oxo-LCA) was obtained: For future studies in order to decrease this byproduct,
since the amount of intermediate is related to the amount of cofactor in solution, lower NAD+
concentrations should be used at the expense of a longer reaction time (e.g. if 0.2 mM of NAD+ is
used, the amount of intermediate cannot exceed the 2%). In order to evaluate the performance of
our system, process metrics were calculated: under the condition described in this study (1 and
0.6 U mL-1 of Sm7α-HSDH and Ls7β-HSDH, respectively) the system shows a TTN of 2.9
million and a space-time yield of 26 g L-1 d-1. Despite a good TTN, the space time yield is not yet
high enough for industrial application. Further studies, including the use of flow reactors, will be
carried out to address the down-stream processing aspects of this reaction. This, in combination
with solvent engineering and the employment of biphasic systems, both targeted to the increase
of substrate loading (and decrease the cofactor loading), could optimize the condition for the
industrial preparation of UDCA[51]
.
In conclusion, we have shown that epimerization reaction of CA and CDCA can be achieved by a
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two-enzyme-one-cofactor cascade which benefits from favorable themodynamics for the 7β-OH
isomer. In particular Sm7α-HSDH and Ls7β-HSDH are a compatible, stable set of enzymes and
promising candidates as biocatalyst in the synthesis of hydroxysteroid derivatives. Further
investigations will be carried out in order to develop a flow system or a membrane reactor similar
to the ones described in literature[22, 52] for our newly developed biocatalytic UDCA synthesis
procedure and to increase the substrate loading in the process.
6 Experimental Section
6.1 Bacterial strains and materials
Escherichia coli TOP 10 [F- mcrA Δ(mrr-hsdRMS-mcrBC) φ80lacZΔM15 ΔlacX74 nupG recA1
araD139 Δ(ara-leu)7697 galE15 galK16 rpsL(StrR) endA1 λ-] and BL21(DE3) [F– ompT gal dcm
–
–
lon hsdSB(rB mB ) λ(DE3 [lacI lacUV5-T7p07 ind1 sam7 nin5]) [malB+]K-12(λS)] were purchased
from Invitrogen (Darmstadt, Germany). Tryptone, yeast extract, cholic acid, ursodeoxycholic
acid and nicotinamide adenine dinucleotide cofactors (NAD(P)+/H) were from Sigma-Aldrich
(St. Louis, US). Ursodeoxycholic acid was from Tokio Chemical Industry (TCI) (Tokyo, Japan).
Restriction enzymes and Phusion Q-5 DNA polymerase mastermix were from New England
Biolabs (NEB, Ipswich, US). All other reagents were of analytical grade and commercially
available.
6.2 Identifying and homology models of 7α-hydroxysteroid dehydrogenases
The 7α-hydroxysteroid dehydrogenase from Stenotrophomonas maltophilia (Sm7α-HSDH) and
7β-hydroxysteroid dehydrogenase from Lactobacillus spicheri (Ls7β-HSDH) gene sequences
were
identified
by
multiple
sequence
analysis
using
BLASTp
(NCBI,
https://blast.ncbi.nlm.nih.gov): for Sm7α-HSDH, protein sequences with known activity from
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Clostridium sardiniense (GenBank: AET80685.1), Clostridium difficile (GenBank: CAJ66880.1,
putative), Clostridium sordelli (GenBank: L12058.1), Eubacterium sp. VPI 12708 (GenBank:
M58473.1), Bacteroides fragilis (GenBank: AF173833.2) and Escherichia coli (Gene ID:
ACI83195.1) were used as query, restricting the organisms list to Stenotrophomonas maltophilia
strains (taxid:40324). A multiple sequence analysis of all 7α-HSDH is shown in Supplementary
Figure 1a.
A
3D structure model of this enzyme was obtained using SWISS-MODEL
(https://swissmodel.expasy.org/interactive), employing the crystal structure of the 7α-HSDH
from Escherichia coli (PDB ID: 1AHI.1) as template.
For Ls7β-HSDH, protein sequences from Clostridium sardiniense (GenBank: AET80684.1),
Ruminococcus gnavus (Genbank: WP004843516.1), Ruminococcus torques (GenBank:
WP015528793.1) and Collinsella aerofaciens (GenBank: WP006236005.1) together with the
variant of the Cs7β-HSDH obtained (see below) were used as query. A multiple sequence
analysis of all 7β-HSDH is shown in Supplementary Figure 1b.
The 3D models of Cs7β-HSDH (GenBank: AET80684.1) and Ls7β-HSDH were built using the
same server employing the entire structure of 7β-HSDH from Collinsella aerofaciens (PDB code
5GT9) as template.
6.3 Cloning and expression of Sm7α-HSDH, Cs7β-HSDH and Ls7β-HSDH
The synthetic cDNAs encoding the Sm7α-HSDH, Cs7β-HSDH and Ls7β-HSDH were designed
by in silico back translation of the amino acid sequences (GenBank: KRG42928.1, AET80684.1
and WP045806907, respectively). In order to subclone into the pET24d(+) plasmid (Merck
Millipore, Burlington, US), sequences corresponding to NcoI (CCATGG) and XhoI (CTCGAG)
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restriction sites were added at the 5’- and 3’-ends of the cDNAs, respectively. The codon usage
of the synthetic genes was optimized for expression in E. coli and produced by BaseClear B.V.
(Leiden, The Netherlands). Sm7α-HSDH, Cs7β-HSDH and Ls7β-HSDH cDNAs were cloned into
the pET24d(+) vector using the NcoI and XhoI sites, resulting in 6.0-, 6.0- and 6.1-kb construct
(pET24-Sm7α-HSDH, pET24-Cs7β-HSDH and pET24-Ls7β-HSDH). The genes were cloned in
frame with the C-terminal 6-Histidines tag of the vector.
The obtained expression plasmids were then used to transform BL21(DE3) E. coli cells.
Starting cultures (100 mL) were prepared from a single recombinant BL21(DE3) E. coli colony
grown in LB medium containing kanamycin (30 μg mL-1), under vigorous shaking (200 rpm) at
37 °C. These cultures were diluted to a starting OD600nm of 0.1 in 1 L of LB medium (LB, 10 g L-
bacto-tryptone, 10 g L-1 NaCl and 5 g L-1 yeast extract) and then incubated at 37 °C on a
1
rotatory shaker at 200 rpm until an OD600nm of 1.0 was reached. Protein expression was induced
by adding 0.25 mM IPTG: cultures were grown for another 12 h at 25 °C with shaking (200 rpm).
Cells were harvested by centrifugation at 10,000 x g for 10 min at 4 °C, washed with 50 mM KPi
buffer pH 8.0 and stored at −20 °C for at least 1 day before purification.
6.4 Protein purification
E. coli cell pellets were resuspended in lysis buffer (50 mM KPi buffer, 1 M NaCl, 5% glycerol
(v/v) and 10 μg mL-1 DNAse, pH 8.0) and disrupted by French press (Constant Systems Limited,
Low March, UK) (2 cycles, 180 psi). The insoluble fraction of the lysates were removed by
centrifugation at 39,000 x g for 30 min at 4 °C. Crude extract was loaded onto a HiTrap chelating
affinity columns (GE Healthcare, Little Chalfont, UK), previously loaded with Ni2+ metal ions
and equilibrated with 50 mM KPi buffer, 1 M NaCl and 5% glycerol (v/v) pH 8.0. The columns
were washed with this buffer until the absorbance value at 280 nm was that of the buffer and the
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bound proteins were eluted with 50 mM KPi buffer, 250 mM imidazole and 5% glycerol (v/v),
pH 8.0. The fractions containing the desired activity were dialyzed overnight against 50 mM KPi
buffer and 5% glycerol (v/v), pH 8.0, using a 3-kDa dialysis tube.
During the purification procedure, 7α-HSDH and 7β-HSDH activities were assayed using the
standard activity assay (see below).
6.5 Activity and kinetic measurements
7α-HSDH’s enzymatic activity in the crude extract and of the purified enzyme was determined at
25 °C using 1.0 mM CDCA, 2.0 mM NAD+, in 50 mM KPi buffer and 10% methanol (v/v), pH
8.0. 7β-HSDH’s enzymatic activity was determined at room temperature (25 °C) using 1.0 mM
UDCA, 2.0 mM NAD(P)+, in 50 mM KPi buffer and 10% methanol (v/v), pH 8.0. The
production or the consumption of NAD(P)H was followed at 340 nm (extinction coefficient of
NAD(P)H is 6,220 M−1·cm−1). One unit (U) was defined as the amount of enzyme producing 1
µmol of product per minute at 25 °C and at pH 8.0. Blank measurements were performed in
absence of CDCA or UDCA, NAD+ and enzyme.
The kinetic parameters of the purified samples were determined at room temperature in the
presence of: different concentrations of substrates (5–10000 μM), 2.0 mM NAD(P)+ in 50 mM
KPi buffer and 10% methanol (v/v), pH 8.0, at 25 °C; different concentrations of NAD(P)+ (1–
5000 μM), 2.0 mM CDCA (for 7α-HSDH) or UDCA (for 7β-HSDH) in 50 mM KPi buffer and
10% methanol (v/v), pH 8.0, at 25 °C. The specific activity was expressed as unit per mg of
protein (determined by spectrophotometric analysis at 280 nm). The kinetic data were fitted to the
Michaelis–Menten equation, or to the one modified to account for substrate inhibition when
necessary.
The effect of pH on the enzymatic activities was determined using 1.0 mM CDCA (for 7α-
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HSDH) or UDCA (for 7β-HSDH), 2.0 mM NAD(P)+, in 100 mM citrate-phosphate buffer (66
mM citrate, 34 mM Na2HPO4) and 10% methanol (v/v), in the 3.0–10.0 pH range.
The effect of methanol concentration on the enzymes activity toward CDCA and UDCA was
determined using 1.0 mM CDCA (for 7α-HSDH) or UDCA (for 7β-HSDH), 2.0 mM NAD(P)+ in
50 mM KPi buffer and different concentration of methanol (0-50% (v/v)), pH 8.0, at 25 °C.
Temperature dependence of the enzymatic activities was determined using 1.0 mM CDCA (for
7α-HSDH) or UDCA (for 7β-HSDH), 2.0 mM NAD+ in 50 KPi buffer and 10.0% methanol (v/v),
pH 8.0 in the 18–95 °C temperature range.
Enzymatic stability was measured by incubating the enzyme solution in 100 mM citrate-
phosphate buffer (66 mM citrate, 34 mM Na2HPO4) in the 3.0–9.0 pH range at 25 °C[53], in 50
mM KPi buffer with different concentration of methanol (0-50% (v/v)) at pH 8.0 at 25 °C and in
50 mM KPi buffer, at pH 8.0 at different temperatures: samples were withdrawn at different
times and residual activity was determined using the enzymatic activity assay.
6.6 SDS-PAGE
Proteins from crude extract and the purified enzyme fractions were separated by SDS-PAGE on a
12% polyacrylamide resolving gel (BioRAD, Hercules, US): samples were resuspended in an
appropriate volume of Laemmli sample buffer and boiled. Proteins were visualized by staining
with SimplyBlue safe stain (Novex, Carlsbed, US).
6.7 Site-saturation mutagenesis (SSM) and screening NAD+ dependent enzyme variants
SSM was carried out at different amino acid positions of Cs7β-HSDH by using whole plasmid
PCR[54]; pET24d-Cs7β-HSDH vector as template and a set of degenerated synthetic
oligonucleotides was employed to prepare the mutant libraries (Supplementary Table 1). PCRs
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were carried out in a final volume of 20 μL: in all cases, 1 μL of template DNA (~25 ng μL-1),
0.5 μL of each primer (final concentration 0.25 μM), 10 μL of Phusion Q-5 DNA polymerase
mastermix and 8 μL of MilliQ water were added in sterile PCR tube. After an initial denaturation
step (98 °C for 1 min), reaction was carried out for 30 cycles (denaturation at 98 °C for 30 sec.,
annealing at 58 °C for 30 sec. and elongation at 72 °C for 7 min). A final elongation step (72 °C
for 7 min.) was added. The template DNA was eliminated by enzymatic digestion with 1 μL
DpnI restriction enzyme at 37 °C for 2 hours; the PCR products were used to transform E. coli
TOP10 cells. Subsequently, the recombinant plasmids were transferred to E. coli BL21(DE3)
cells, and these clones were used for the screening procedure. The introduction of the mutations
was confirmed by automated DNA sequencing. The mutant libraries obtained from SSM were
screened by means of a rapid colorimetric assay based on the reduction on NAD+ (as described
before) and by means of an automated JANUS Liquid Handler Workstations (Perking Elmer,
Waltham, US). To a saturated E. coli culture (1 mL, growth in 2mL DeepWell plate) 0.250 mM
IPTG were added and the culture was then incubated at 25 °C for 18 h. The culture was
centrifuged at 5,000×g for 2 min, and the cell pellet was resuspended in 200 μL of 50 mM KPi
buffer, pH 8.0 containing of 1 mg mL-1 lysozyme. Cell lysis was performed incubating the plate
for 30 minutes at 37 °C, 200 rpm. The crude extracts were centrifuged at 5,000×g for 30 min and
then 50 μL of the supernatant was transferred to a 96-well flat-bottom plate. The activity was
assayed on the crude extract by adding 150 μL of 1.33X substrate solution (1.33 mM UDCA,
2.66 mM NAD+, 13.3% MeOH in 50 mM KPi buffer, pH 8.0).
The initial activity was determined by measuring the increase of the absorbance at 340 nm for 5
min at 25 °C in a microtiter plate reader (Synergy2, Biotek, Winoosky, US) and compared with
cultures expressing the wild-type Cs7β-HSDH and untransformed cells as controls. The selected
variants were sequenced and biochemically characterized.
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6.8 Preparation of 7-oxo-DCA, 7-oxo-LCA, UCA and UDCA
In a 2 L round-bottom flask 2.1 g of CA, 133 mg of NAD+ and 1.98 g of oxalacetate were
dissolved in 50 mM of KPi buffer, pH 8.0 with 10% MeOH (final volume 1 L). The reaction was
initiated adding 15 mg of Sm7α-HSDH (5100 Utot) and 200 Utot of malate dehydrogenase (Sigma-
Aldrich, St. Louis, US).
The reaction was gently stirred at 25 °C. Conversion was checked with HPLC. At complete
reaction (~1 h) the solution was acidified with HCl till pH 2.0, leading to the formation of white
suspension. In order to increase the precipitation 50 g of NaCl was added and dissolved. The
product was filtered with a porous glass filter and washed with 20 mL of 0.01 M HCl solution.
The powder was then dried and crystallized in MeOH giving 7-oxo-DCA. Yield: 1.9 g (95%);
Purity: 98% (assayed by HPLC analysis); 13C NMR (100 MHz, DMSO): δ 211.75 (C7=O),
175.15 (COOH), 70.26 (C12α-OH), 69.33 (C3α-OH); full 1H and 13C NMR spectra are reported in
supplementary material.
The same procedure was used for the preparation of 7-oxo-LCA from 1.9 g of CDCA. Yield: 1.6
13
g (84%); Purity: 98% (assayed by HPLC analysis); C NMR (100 MHz, DMSO): δ 211.40
1
(C7=O), 174.86 (COOH), 69.09 (C3α-OH); full H and 13C NMR spectra are reported in
supplementary material.
UCA and UDCA were obtained by using the enzymatic cascade reported in this paper starting
from CA and CDCA, respectively: 0.55 g of CA or 0.51 g of CDCA and 86 mg of NAD+ were
dissolved in 50 mM of KPi buffer, pH 8.0 with 10% MeOH (final volume 130 mL). The reaction
was initiated by adding 0.18 mg of Sm7α-HSDH (63 Utot) and 12.54 mg of Ls7β-HSDH (39 Utot).
The reaction was gently stirred at 25 °C. After 24 h the reaction was complete (92% and 91%
conversion of CA into UCA and of CDCA into UDCA, respectively). The reactions was acidified
with HCl (final pH 3.0) and extracted with diethyl ether. UDCA and UCA were purified by
24
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Reveleris X2 apparatus (GRACE, Columbia, US) equipped with C18 cartridge (40μm, 12 g)
employing Water (TFA 0.1%) : acetonitrile gradient. UCA: Yield: 478 mg (87%); Purity: >99%
(assayed by HPLC analysis); 13C NMR (100 MHz, DMSO): δ 174.98 (COOH), 70.66 (C7β-OH),
69.83 (C12α-OH), 69.56 (C3α-OH). UDCA: Yield: 438 mg (86%); Purity: >99% (assayed by
HPLC analysis); 13C NMR (100 MHz, DMSO): δ 174.90 (COOH), 69.74 (C7β-OH), 69.46 (C3α-
OH). Full 1H and 13C NMR spectra are reported in supplementary material.
13
1H and C NMR spectra, in agreement with previous results[18, 29, 50] were acquired with Agilent
ProPulse 400 MHz (Santa Clara, US) and are reported in supplementary materials.
Molecular mass of 7-oxo-DCA, 7-oxo-LCA, UCA and UDCA were confirmed using an RP-ESI-
Q-TOF-MS system (M-Class and Q-TOF Premier, Waters, UK) which was operated in positive
ionization mode (ES+). Mass calibration was performed using [Glu 1 ]-Fibrinopeptide B. Data
were analyzed using the MassLynx 4.1 tool box. The MS analyses are reported in supplementary
material.
6.9 Bioconversion of CDCA to UDCA.
All bioconversions were carried out employing 1 Utot of purified Sm7α-HSDS and 0.6 Utot of
purified 7β-HSDS on 10 mM (if not differently specified) of CDCA and CA, NAD+ (0.2, 0.5 or
1.0 mM). As general procedure, 1 mL of reaction mixture containing 10% MeOH and 50 mM of
KPi buffer, pH 8.0, was incubated at 25 °C: at fixed times 50 μL of reaction were withdrawn,
diluted with 250 μL of MeOH and centrifuged at 14000 xg for 2 min. 10 μL of the obtained
samples were analyzed by HPLC. HPLC analyses were performed on a Shimadzu (Kyoto, Japan)
apparatus equipped with a LC20AT pump and an ELSD-LTII detector and fitted with a XTerra
RP C18 column (Waters, Milford, US) (length/internal diameter 150/4.6 mm, pore size 5 μm)
under the following conditions: for analyses of CA, 7-oxo-DCA and UCA, eluent
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H2O/CH3CN/TFA (70/30/0.1), flow 1.0 mL min-1. Retention times CA=6.75 min, 7-oxo-
DCA=4.82 min, UCA= 4.03 min. For analyses of CDCA, 7-oxo-LCA and UDCA, eluent
H2O/CH3CN/TFA (65/35/0.1), flow 1.0 mL min-1. Retention times CDCA=4.11 min, 7-OXO-
LCA=3.49 min, UDCA=3.21 min.
7 Acknowledgments:
This work was done as part of the ONE-FLOW project (https://one-flow.org) that has received
funding from the European Union's FET-Open research and innovation actions (proposal 737266
- ONE-FLOW). We thank Dr. M. Pabst and P. van Dam for LC-MS measurement and Prof. U.
Hanefeld, Dr. F. Hollmann, Dr. D. McMillan and all the members of the biocatalysis section of
TU Delft for the scientific discussions.
8 Abbreviations:
API, active pharmaceutical ingredient; CA, Cholic acid; CDCA, Chenodeoxycholic acid; DCA,
Deoxycholic acid; DHCA, Dehydrocholic acid; LCA, Lithocholic acid; UCA, Ursocholic acid;
UDCA, Ursodeoxycholic acid; HSDH, Hydroxysteroid dehydrogenase; LDH, Lactate
dehydrogenase; GDH, Glucose dehydrogenase; SSM, site-saturation mutagenesis.
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