Catalytic hydroxylation in biphasic systems using CYP102A1 mutants

Article abstract of DOI:10.1002/adsc.200505044

Cytochrome P450 monooxygenases are biocatalysts that hydroxylate or epoxidise a wide range of hydrophobic organic substrates. Their technical application is, however, limited to a small number of whole-cell processes. The use of the isolated P450 enzymes is believed to be impractical due to their low stability, stoichiometric need of the expensive cofactor NAD(P)H and low solubility of most substrates in aqueous media. We investigated the behaviour of an isolated bacterial monooxygenase (mutants of CYP102A1) in a biphasic reaction system supported by cofactor recycling with the NADP ⁺-dependent formate dehydrogenase from Pseudomonas sp 101. Using this experimental set-up cyclohexane, octane and myristic acid were hydroxylated. To reduce the process costs a novel NADH-dependent mutant of CYP102A1 was designed. For recycling of NADH an NAD⁺-dependent FDH was used. The stability of the monooxygenase mutants under the reaction conditions in the biphasic system was quite high as revealed by total turnover numbers of up to 12,850 in the NADPH-dependent cyclohexane hydroxylation and up to 30,000 in the NADH-dependent myristic acid oxidation.

Full text of DOI:10.1002/adsc.200505044

FULL PAPERS
Catalytic Hydroxylation in Biphasic Systems using
CYP102A1 Mutants
Steffen C. Maurer, Katja Kꢀhnel, Leonard A. Kaysser, Sabine Eiben,
Rolf D. Schmid, Vlada B. Urlacher*
Institute of Technical Biochemistry, Stuttgart University, Allmandring 31, 70569 Stuttgart, Germany
Fax: (þ49)-711-685-3196, e-mail: itbvur@itb.uni-stuttgart.de
Received: January 24, 2005; Revised: March 11, 2005; Accepted: April 5, 2005
Abstract: Cytochrome P450 monooxygenases are bio- myristic acid were hydroxylated. To reduce the proc-
catalysts that hydroxylate or epoxidise a wide range of ess costs a novel NADH-dependent mutant of
hydrophobic organic substrates. Their technical appli- CYP102A1 was designed. For recycling of NADH
cation is, however, limited to a small number of an NADþ-dependent FDH was used. The stability
whole-cell processes. The use of the isolated P450 en- of the monooxygenase mutants under the reaction
zymes is believed to be impractical due to their low conditions in the biphasic system was quite high as re-
stability, stoichiometric need of the expensive cofac- vealed by total turnover numbers of up to 12,850 in
tor NAD(P)H and low solubility of most substrates the NADPH-dependent cyclohexane hydroxylation
in aqueous media. We investigated the behaviour of and up to 30,000 in the NADH-dependent myristic
an isolated bacterial monooxygenase (mutants of acid oxidation.
CYP102A1) in a biphasic reaction system supported
by cofactor recycling with the NADPþ-dependent for- Keywords: biotransformations; biphasic system; co-
mate dehydrogenase from Pseudomonas sp 101. Us- factor recycling; cofactor specificity; green chemistry;
ing this experimental set-up cyclohexane, octane and hydroxylation; P450 monooxygenase
Introduction
covery of products from a complex fermentation broth.
Considering these problems, the use of isolated oxygen-
Cytochromes P450 belong to the heme-containing en- ases in enzyme reactors can be advantageous.[6–9]
zyme class of monooxygenases (EC 1.14.x.y). They are
From a technical point of view, microbial P450s are
widely distributed in nature and play an important role easier to handle than P450 enzymes from plants and an-
in primary and secondary metabolism as well as in de- imals. They are not membrane-associated and exhibit a
toxification of xenobiotic compounds.[1] P450 enzymes relatively high stability and activity.[10]
catalyse the introduction of molecular oxygen into acti-
Particularly appropriate for cell-free processes are so-
vated and even non-activated aliphatic or aromatic XH called “self-sufficient” P450 monooxygenases. They
bonds (X¼C, N, S). Moreover, a remarkable number of consist of a heme domain fused to an FAD- and FMN-
¼
P450 enzymes are capable of epoxidising C C double containing P450 reductase domain and, in contrast to
bonds.[2] Common to both reaction types is the incorpo- the majorityofP450s, do not have tobe supplied with ad-
ration of one atom of dioxygen into the substrate, while ditional redox partners (reductases) apart from
the other atom is reduced to water by electrons ultimate- NADPH. The best characterised fusion protein
ly originating from the nicotinamide cofactor CYP102A1 from Bacillus megaterium has been inten-
NAD(P)H. Electrons are transferred from NAD(P)H sively studied during the last two decades.[11,12] This en-
to the P450 heme iron via a flavin reductase or/and an zyme converts saturated and unsaturated fatty acids
iron-sulphur protein.[3]
with chain lengths of C12 to C22 with high activity (up
Currently, the use of P450 enzymes in industrial proc- to 4000 minꢀ1) and partially also high stereoselectivity
esses is restricted to whole-cell biotransformations.[4,5] to their subterminally oxygenated derivatives.[13,14] Re-
Advantages of such systems are stabilisation of the com- cently, cloning and characterisation oftwo further fusion
plex P450 systems with up to three individual proteins enzymes CYP102A1 and CYP102A3 from Bacillus sub-
and simultaneous regeneration of cofactors in the cellu- tilis, exhibiting high homology to CYP102A1, was re-
lar metabolism. Disadvantages of whole-cell processes ported.[15–17] While a large number of publications exist
might be a further degradation of products, toxicity of describing novel evolved CYP102A1 mutants with al-
the educts or products for the cell, and complicated re- tered substrate specificity,[18–26] there are only very few
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Catalytic Hydroxylation in Biphasic Systems
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reports about the use of these biocatalysts in preparative
organic synthesis.[7,27] The main drawback for in vitro ap-
plications of P450s is the dependence on the highly ex-
pensive cofactor NAD(P)H, which has to be added in
stochiometric or even higher amounts (if uncoupling is
taken into account). Several possibilities exist to over-
come this problem, for example, the use of peroxides
(so-called shunt-pathway)[28] or cathodic reduction of
the heme iron.[29–31] Other approaches are electrochem-
ical[32] or enzymatic reduction of the oxidised cofactor
NAD(P)þ during biotransformations. Dehydrogenases
are currently utilised for cofactor recycling in various bi-
ocatalytic processes.[7,33,34]
Figure 1. Results of screening a CYP102A1 mutant library
for hydroxylation activity towards cyclohexane. NADPH
consumption in the presence of 470 mM cyclohexane was
tracked for 3 minutes.
In a previous paper we reported a cofactor recycling
system for CYP102A1 by means of an NADPþ-depend-
ent formate dehydrogenase (FDH, EC 1.2.1.2) from
Pseudomonas sp. 101. It has been shown that both en-
zymes can be used in combination.[34]
Using this approach we have developed a process for pocket[39] also showed rather high activity of 38 eq
oxidation of cyclohexane in a biphasic (cyclohexane/ eqꢀ1 minꢀ1, however lower than the double mutantꢂs ac-
aqueous buffer) system. Industrial oxidation of cyclo- tivity.
hexane is a process still open to improvements. Thus,
Uncoupling of NADPH oxidation and substrate hy-
on one hand, this reaction serves as a model for activa- droxylation resulting in reduction of oxygen to hydro-
tion of an inert hydrocarbon, on the other hand there gen peroxide or water is an issue frequently encountered
might be interest in an alternative approach for cyclo- during monooxygenase catalysis.[38,40] Therefore product
hexane oxidation.
formation and its coupling to NADPH consumption was
In order to prove the general applicability of investigated by GC/MS analysis (data not shown). This
CYP102A1 mutants in biphasic reaction systems, the hy- revealed coupling efficiencies of 25% for the R47L,
droxylation of octane and myristic acid was also investi- Y51F mutant, 19% for R47L, Y51F, F87V, L188Q,
gated.
16% for A74G, L188Q and only 6% for wild-type
Further reduction of the process costs was achieved by CYP102A1. Generally we found correlation between
switching the cofactor specificity of CYP102A1 from NADPH oxidation rates and corresponding coupling ef-
NADPH to NADH. The advantage of NADH is its low- ficiencies. The faster the NADPH oxidation was the
er price and higher stability.[35] Besides that, NADþ-de- higher was the coupling efficiency. All mutants, for
pendent dehydrogenases are more abundant in nature which coupling efficiency was measured, produced cy-
than their NADPþ-dependent analogues, opening up clohexanol as a single product (GC/MS data not shown).
the possibility to combine CYP102A1-catalysed reac- The double mutant R47L, Y51F was revealed as the
tions with a large variety of dehydrogenases.
most active (cyclohexane hydroxylation activity:
14 minꢀ1 for R47L, Y51F compared to 0.75 minꢀ1 for
the wild-type enzyme) among the mutants investigated
and therefore used in further experiments with this sub-
strate. As activity of the wild-type enzyme in this reac-
tion was almost negligible it was not investigated for
preparative purposes.
Results and Discussion
Identification of CYP102A1 Mutants Hydroxylating
Cyclohexane
A set of CYP102A1 mutants constructed by site-direct- Kinetic Characterisation
ed mutagenesis was screened for activity towards cyclo-
hexane by monitoring NADPH oxidation at 340 nm dur- Kinetic data for the purified mutant R47L, Y51F were
ing the hydroxylation reaction. The results are summar- determined by monitoring of cyclohexane-dependent
ised in Figure 1.
NADPH oxidation at a saturating concentration of the
The highest activity (56 eq eqꢀ1 minꢀ1) was detected cofactor.
for the CYP102A1 mutant R47L, Y51F. Both amino
The plot of NADPH oxidation rate versus cyclohex-
acid substitutions render the substrate access channel ane concentration shown in Figure 2 has non-hyperbolic
more hydrophobic causing easier access of cyclohexane characteristics. This indicates cooperativity in substrate
to the active centre of the enzyme.[36–38] The mutant binding according to the Hill kinetic model. Even at the
R47L, Y51F, F87V, L188Q with an enlarged binding maximum cyclohexane concentration (ꢁ500 mM) that
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These measurements confirmed the steady-state ki-
netic data. This is, however, atypical for CYP102A1
that usually corresponds to Michaelis–Menten kinetics.
The results summarised in Figures 2 and 3 clearly
demonstrate that application of the CYP102A1 mutant
R47L, Y51F would benefit from an increase in cyclohex-
ane concentration in the aqueous phase. Different deter-
gents such as Tween 20, Tween 80, cholate, CHAPS, a-
cyclodextrin, b-cyclodextrin and co-solvents such as
methanol, ethanol, DMSO, acetone, butanol, THF
were tested with CYP102A1 R47L, Y51F in order to in-
crease substrate solubility. Only Tween 20 and 80 ap-
plied at 1% (v/v) displayed a positive effect leading to
1.5- and 2-fold higher cyclohexane hydroxylation activ-
ity. However, these beneficial effects could not be trans-
ferred to the two-phase reaction system. While Tweens
enhanced the activity of the monooxygenase by increas-
ing the cyclohexane concentration, the stability of mon-
ooxygenase and/or dehydrogenase was negatively
affected.
Figure 2. Steady-state NADPH turnover rates by purified
CYP102A1 mutant R47L, Y51F at various cyclohexane con-
centrations. The plot demonstrates sigmoidal dependency of
the reaction rate on substrate concentration. This indicates
non-Michaelis–Menten kinetics.
is limited by its solubility in the aqueous phase, full sat-
uration of the active centre of the enzyme was not
reached.
Biphasic Reaction System Cyclohexane/Aqueous
Substrate binding to P450 usually causes a heme spin- Buffer
state shift from low to high, which can be monitored as
changes in P450 absorbance spectra (Figure 3). Cyclo- We investigated the optimal conditions for hydroxyla-
hexane binds to R47L, Y51F leading to an increase in tion of cyclohexane by CYP102A1 in a biphasic reaction
the difference between the absorbance at 417 nm system (Figure 4). In this system cyclohexane acts as a
(trough) and that at 387 nm (peak). However, the max- substrate pool and simultaneously as an organic solvent
imum heme spin-state shift was not reached at a concen- for extraction of the reaction product. Thus, the catalytic
tration of cyclohexane of 500 mM.
system is retained in the aqueous phase while the prod-
The plot of absorption differences at ~387 nm and uct may continuously be isolated from the organic
417 nm versus cyclohexane concentration indicates ex- phase. The cofactor recycling system is based on a mu-
ponential growth of the spectral changes in the concen- tant of FDH from Pseudomonas sp. which has been pro-
tration range inferred through the solubility limit (small ven to be highly stable in organic solvents.[8,41]
graph in Figure 3).
Figure 3. Cyclohexane binding difference spectra of CYP102A1 R47L, Y51F. Difference spectra were obtained by subtracting
reference spectra of enzyme (8.6 mM) without substrate from spectra of samples containing the indicated cyclohexane concen-
trations and exactly the same enzyme concentration as the reference. The small chart at the right upper corner was generated
by plotting A387 minus A417 data from the difference spectra versus the relevant cyclohexane concentrations.
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Figure 5. Production of cyclohexanol during reaction in the
biphasic system. The reaction was performed in a stirred
20-mL flask containing 5 mL cyclohexane and 5 mL aqueous
reaction medium. Squares: Negative control containing E.
coli cell extract; circles: CYP102A1 mutant A74G, F87V,
L188Q; triangles: CYP102A1 mutant R47L, Y51F.
Figure 4. Scheme for CYP102A1-mediated hydroxylation of
cyclohexane in a biphasic system. The organic phase serves
as substrate pool and extracts the products from the aqueous
phase.
the mutant R47L, Y51F was active for at least four
days (~100 h) when stabilised by addition of BSA and
catalase (Figure 5).
Limitations in transfer of substrate from the organic to
the aqueous phase can decrease reaction rates, at least if
The stability of this dehydrogenase in a 1:1 mixture of phase transfer is slower than the rate of hydroxylation.
FDH reaction buffer and cyclohexane under vigorous
The triple mutant displaying a lower reaction rate
stirring was tested. Studies on FDH activity revealed a than the double mutant (Figure 1; 7 eq eqꢀ1 minꢀ1 com-
half-life of at least one day for this cofactor regenerating pared to 56 eq eqꢀ1 minꢀ1, respectively) also showed
enzyme in this system.
lower cyclohexanol production in the biphasic reaction
The effect of different protein stabilising agents – glyc- system (Figure 5). For the best mutant R47L, Y51F
erol, bovine serum albumin (BSA), catalase, PEG deriv- the total turnover number (ttn) was 9620 (Table 1). An
atives – was studied. Addition of 10 mg mLꢀ1 BSA and increase in ttn to 10280 was achieved by changing the re-
0.2 mg mLꢀ1 (600 U mLꢀ1) catalase from horse liver sta- action vessel from a stirred flask at room temperature to
bilised the CYP102A1 monooxygenases in the biphasic a flask shaken vigorously at 160 rpm and 188C.
emulsion and led to higher hydroxylation activity for an
extended period of time (data not shown).
In a further experiment, the reaction was scaled up to
one litre volume (1.3 mmol of CYP102A1 R47L, Y51F)
Concentration of cyclohexanol in the organic phase and was carried out in a fermenter under continuous aer-
measured during the biotransformation indicates that ation. On-line monitoring of oxygen partial pressure
Table 1. Comparison of the biotransformation reactions in biphasic reaction media.
Substrate
CYP102A1 mutant Reaction Reaction volume Amount
ttn for
ttn for
Volumetric
productivity
time [h] [mL] of CYP102A1 CYP102A1 cofactor
[b]
[nmol]
mutant
NAD(P)þ[a] (mg Lꢀ1 hꢀ1
)
Cyclohexane R47L,Y51F
Cyclohexane A74G,F87V,L188Q 100
Cyclohexane R47L,Y51F
Cyclohexane R47L,Y51F
100
10 (stirred flask)
10 (stirred flask)
40 (shaking flask) 180
1000 (fermenter) 1300
10 (stirred flask)
8 (stirred flask)
10 (stirred flask)
66
66
9620
2370
10280
12850
2200
1270
313
925
334
280
825
300
63.5
15.6
46.3
16.7
18.6
100
100
Octane
A74G,F87V,L188Q 100
65
100
5
Myristic acid A74G,F87V,L188Q 80
Myristic acid A74G,F87V,L188Q, 24
R966D,
3300
30000
101
153
W1046S (NADH-
dependent)
[a]
NAD(P)H concentration was 0.1 mM in the aqueous phase.
Calculated for the whole reaction time. Higher volumetric activities can be calculated for the beginning of the reactions.
[b]
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and pH allowed us to control the reaction progress. The droxylated myristic acid derivatives were produced. A
increase in oxygen partial pressure reflects a decrease in hydroxylation of 12-hydroxy- and 13-hydroxymyristic
oxygen consumption during the hydroxylation reaction. acid has previously been reported, even though with
As protons are stoichiometrically consumed during the low reaction rate.[42] Totally, 92.9 mg of myristic acid
reaction (Figure 4), no further increase in pH indicates and derivatives were extracted. This mixture consisted
loss in activity.
of 41.4 mg of myristic acid (predominantly found in
The total turnover number (ttn) in the fermenter the dodecane phase) and of 51.5 mg (55%) of hydroxy-
reached 12,850, yielding 1.67 g of cyclohexanol lated products less soluble in dodecane as revealed by
(16.7 mmol). The course of product development indi- GC/MS. Hydroxylated products comprised 48% mono-
cates that the monooxygenase was active for at least hydroxy- (w-1 to w-7) and 52% dihydroxymyristic acid
100 h. During the first 53 h productivity (12.6 mmol cy- (mixture of 7 regioisomers). As stated in Table 1this cor-
clohexanol) was higher than during the second half of responds to ttns of 3300 for the P450 enzyme and 825 for
the reaction (4.1 mmol). The space-time yield during the cofactor NADPþ.
the first 53 h of the reaction was 23.8 mg Lꢀ1 hꢀ1 (Ta-
These results indicate the applicability of the biphasic
ble 1). This result can be significantly improved by ap- reaction system for conversion of different substrates by
plying higher concentrations of CYP102A1 and FDH. CYP102A1.
In this case only substrate phase transfer will be the lim-
iting factor.
Conversion of Myristic Acid by an NADH-Dependent
CYP102A1 Mutant
Conversion of Octane and Myristic Acid
Multiple sequence alignment of NADPH-dependent
In order to demonstrate the general applicability of cytochrome P450 reductases, which supply the P450
CYP102A1 in biphasic systems, the hydroxylation of oc- monooxygenases with electrons, revealed 5 highly con-
tane and myristic acid were also investigated. The served positions important for cofactor specificity.[43,44]
CYP102A1 mutant A74G, F87V, L188Q hydroxylates These positions in CYP102A1 are S965, R966, K972
both substances with NADPH turnover rates of and Y974, which bind the 2’-phosphate group of
1760 minꢀ1 for octane[19] and 2100 minꢀ1 for myristic NADPH and W1046, the so-called gate-keeping amino
acid.
acid, involved in electron transfer. Dçhr et al. changed
Similar to the experiments with cyclohexane, octane the cofactor specificity of the human P450 reductase
was used as substrate and as organic phase. GC/MS anal- from NADPH to NADH via substitution of the gate-
ysis revealed a product mixture consisting of 2-, 3- and 4- keeping tryptophan to alanine.[44]
octanol as well as the corresponding octanones. The for-
The same effect was observed for the CYP102A1 mu-
mation of octanones by CYP102A1 was previously re- tant W1046A, that could accept NADH as cofactor [KM
ported.[20] A possible mechanism for the formation of (NADH)¼14 mM and kcat (NADH)¼6300 minꢀ1 com-
the ketones is a second hydroxylation of the alcohol to pared to CYP102A1 wild-type reductase: KM
generate a gem-diol, which dehydrates to the corre- (NADPH)¼2.5 mM, kcat (NADPH)¼7930 minꢀ1, KM
sponding ketone. Ketones might alternatively originate (NADH)¼1430 mM and kcat (NADH)¼2810 minꢀ1
]
from vic-diols via pinacol rearrangement. In any case (Table 2) as revealed by reduction of cytochrome C.
double hydroxylation is a prerequisite for ketone forma- However, in our investigation serine at this position
tion. For calculation of total turnover numbers the GC/ caused a stronger effect on the switch in cofactor specif-
MS signals of all reaction products were calibrated for icity [KM (NADH)¼4.4 mM und kcat (NADH)¼
quantification using authentic standards of known con- 4710 minꢀ1]. A further improvement in terms of catalyt-
centration. For octane hydroxylation a ttn of 2200 was ic activity of the CYP102A1 reductase with NADH was
measured.
achieved by substitution of the phosphate binding argi-
For hydroxylation of myristic acid in the biphasic sys- nine 966 by aspartate. Kinetic constants for the reduc-
tem, the substrate was dissolved in dodecane. In prelimi- tase double mutant R966D, W1046S were KM
nary investigations dodecane was proven not tobe a sub- (NADH)¼12 mM, kcat (NADH)¼14601 minꢀ1. Thus,
strate of CYP102A1 A74G, F87V, L188Q. Fatty acid hy- the catalytic efficiency (kcat/KM) of the CYP102A1 re-
droxylation was performed under constant air bubbling. ductase in NADH-dependent cytochrome C reduction
The total reaction volume of 8 mL consisted of 4 mL of was improved 640-fold. Further details on reductase
the aqueous catalyst system and 4 mL 100 mM myristic mutants will be described elsewhere.[45]
acid (91 mg) in dodecane.
The two substitutions R966D and W1046S were intro-
With myristic acid as a substrate for CYP102A1, usu- duced into the reductase domain of the CYP102A1
ally a mixture of the subterminally hydroxylated prod- A74G, F87V, L188Q monooxygenase. The new
ucts is obtained.[13,14] In the biphasic system, however, NADH-dependent mutant was tested with myristic
additionally to the monohydroxylated products dihy- acid in the biphasic system. The NADþ-dependent
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FDH from Pseudomonas sp.[46] was used for cofactor re- dissolve the hydrophobic substrate using a large volume
cycling. Extraction of the reaction mixture and subse- of aqueous buffer. 0.74 mmol of product were obtained
quent GC/MS analysis revealed 20% substrate conver- from 4-L reaction mixture, resulting in a ttn of approxi-
sion to hydroxylated products. The hydroxy- and dihy- mately 1500. Our results revealed not only technical ad-
droxymyristic acid products identified were the same vantages of the biphasic system, but also its high stability
and at approximately the same ratio as in the expressed in a ttn of 12800 with cyclohexane and 30000
NADPH-driven reaction. The lower yield of products with myristic acid as substrate. The total turnover num-
(20% compared to 55% yield in the NADPH-depend- bers of the cofactor ranging from ~300 up to ~1300 are
ent reaction) is most likely a result of the lower amount among the highest reported for a preparative applica-
of P450 enzyme added to the reaction mixture (5 nmol tion of isolated oxygenases.
compared to 100 nmol). The ttn calculated for this reac-
The total turnover numbers, calculated for
tion reached 30000. To our knowledge this represents CYP102A1 are higher than the value of up to 2867 re-
the highest value reported for P450 monooxygenases ported for the asymmetric epoxidation of styrene by
so far (Table 1). Further experiments on use of the styrene monooxygenase (StyAB) in a two-phase dodec-
NADH-dependent system in preparative organic syn- ane/aqueous buffer system.[8] In this system a maximum
thesis are currently being performed.
turnover number of 87 was determined for the cofactor
NADþ.
Lutz et al. investigated the preparative application of
2-hydroxybiphenyl 3-monooxygenase coupled to enzy-
matic cofactor regeneration. The authors reported a
Conclusions
The exploitation of isolated P450 monooxygenases in bi- ttn for NADþ of maximally 503.[47]
ocatalysis is a field of ongoing research. However, until
Zambianchi et al. used isolated cyclohexanone mono-
now it is not clear if the properties of these enzymes al- oxygenase for Baeyer–Villiger oxidation of 10 g Lꢀ1 bi-
low their in vitro use. Although a few reports concerning cyclo[3.2.0]hept-2-en-6-one. The process was running
the use of oxygenases (for a compilation see[47]) in syn- under enzymatic NADPH regeneration and continuous
thetic applications do exist, bioengineering-related substrate feeding.[48]
data are still scarce.
While volumetric productivities reported in our study
We investigated the enzymatic hydroxylation of dif- are quite low (maximum 0.15 g Lꢀ1 hꢀ1), higher values
ferent substrates of CYP102A1 in a biphasic system. found for other oxygenases and reported in literature
[8]
The proposed biphasic reaction scheme (Figure 4) is es- (up to 1 g Lꢀ1 hꢀ1
)
are usually calculated for 10 h or
peciallyfitted to the requirementsof the catalytic system less reaction time. The values were determined for the
under debate: the aqueous phase dissolves – and retains whole reaction time (up to 100 h).
– cofactor and proteins without need for expensive
Switching the cofactor specificity of CYP102A1 from
membrane technology. The organic phase acts as sub- NADPH to NADH allows us to reduce the cofactor
strate pool dissolving substrates or representing the sub- costs to about 20%. The new NADH-dependent mutant
strate itself. Also a substantial amount of reaction prod- demonstrated a comparable stability to that of its
ucts accumulates in the organic phase. These can be con- NADPH-dependent homologue.
tinuously removed using extraction, distillation or ad-
sorption to resins.
The summarised results and the comparison with oth-
er oxygenases pointed out the potential of P450 enzymes
Our experiments (Table 2) can be compared to the for cell-free applications. Further investigations are,
first report of preparative in vitro use of a bacterial however, needed and will address problems of low sub-
P450 monooxygenase.[7] The authors reported a syn- strate solubility, low enzyme activity and low regio- and/
thesis of 14,15-epoxyeicosatrienoic acid from arachi- or stereoselectivity. The CYP102A1 monooxygenase is
donic acid (500 mg, 1.64 mmol) using CYP102A1 a highly “evolvable” biocatalyst. Protein engineering
F87V (600 U, 500 nmol) and glucose 6-phosphate dehy- by directed evolution and site-directed mutagenesis
drogenase for NADPH recycling. Their attempt was to has generated many mutants with versatile properties,
Table 2. Comparison of KM and kcat values for reductase mutants. Values were calculated for NADPH- or NADH-dependent
cytochrome C reduction. The monooxygenase domain in all cases carried mutations A74G, F87V and L188Q.
WT-reductase
W1046A
W1046S
R966D, W1046S
KM (NADPH)
kcat (NADPH)
KM (NADH)
kcat (NADH)
(2.5ꢂ0.2) mM
(0.82ꢂ0.08) mM
(524ꢂ5) minꢀ1
(14.0ꢂ0.75) mM
(6300ꢂ100) minꢀ1
(0.78ꢂ0.05) mM
(366ꢂ4) minꢀ1
(4.4ꢂ0.1) mM
(1.6ꢂ0.2) mM
(7930ꢂ430) minꢀ1
(1430ꢂ70) mM
(5410ꢂ430) minꢀ1
(11.6ꢂ0.7) mM
(2810ꢂ100) minꢀ1
(4710ꢂ40) minꢀ1
(14600ꢂ1200) minꢀ1
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ture for only 4 h, using terrific broth (TB)-medium for expres-
sion and prolonging cultivation time after induction to 16 h at
258C, 800 to 1200 nmol (96 to 144 mg) of soluble CYP102A1
per litre of cell culture were obtained. Aliquots of the crude
cell lysates containing 25 to 50 mM CYP102A1 were stored at
ꢀ208C or directly used for preparative reactions.
including the activity towards a tremendous number of
substrates. Application of these CYP102A1 variants in
biphasic systems represents an efficient and feasible ap-
proach for synthesis of many useful compounds.
For kinetic measurements purified enzyme was used. Purifi-
cation was performed according to a two-step strategy using a
Ni-sepharose HP (Amersham Biosciences, Sweden) column
with elution by 100 mM imidazole first. For further purification
and to remove imidazole, the eluate was loaded onto a fracto-
gel EMD DEAE 650S (Merck KgaA, Darmstadt, Germany)
column and eluted by applying a linear sodium chloride gradi-
ent.
Experimental Section
Chemicals, Enzymes and Strains
All chemical reagents were of analytical grade or higher and
¨
purchased from Fluka, Aldrich, Sigma or Riedel-de-Haen.
NADPH tetrasodium salt and FDH from Pseudomonas sp.
101 were procured from Jꢀlich Fine Chemicals (Jꢀlich, Germa-
ny). E. coli strain BL21 (DE3) and vector pET28aþwere ob-
tained from Novagen (Madison, Wisconsin, USA).
Cyclohexane was distilled prior to use in order to remove
traces of cyclohexanol and cyclohexanone.
Saturated solutions of substrates in aqueous buffer were
prepared by adding 5 mL of organic substrate to 50 mL of
50 mM potassium phosphate buffer (KPi), pH 7.5. The bipha-
sic mixture was vigorously stirred overnight, the substrate-sa-
turated aqueous phase was separated using a separatory funnel
and used immediately.
Activity Assays and Characterisation of Mutants
Substrate binding difference spectra, NADPH oxidation and
CO difference spectra were recorded on an Ultrospec
3000 UV/vis spectrometer (Amersham Biosciences, Sweden).
Concentration of correctly folded P450 enzymes was deter-
mined from the CO-binding difference spectra of the reduced
heme iron using an extinction coefficient of 91 mMꢀ1 cmꢀ1 as
reported elsewhere.[52]
For selection of mutants with activity towards cyclohexane
160 mL cyclohexane-saturated 50 mM KPi, pH 7.5 (0.59 mM
cyclohexane) per well (96-well microtitre plate) was mixed
with 20 mL of cell lysate and the reaction was started by addi-
tion of 20 mL 1 mM NADPH stock solution in 50 mM KPi. Re-
action progress was monitored by measuring the absorption of
NADPH at 340 nm for 5 minutes. For each mutant a blank con-
taining no substrate was recorded and subtracted from the
slope recorded in the presence of substrate. Each of these
measurements was repeated eight times and the average slope
during the first 3 min of the reaction was calculated. For calcu-
lations of turnover numbers an extinction coefficient for
NADPH of 6.22 mMꢀ1 cmꢀ1 at 340 nm was used.
Design of NADH-Dependent Reductase Mutants
For creation of an NADH-dependent mutant of CYP102A1 a
sequence alignment with the rat cytochrome P450 reductase
was performed and two positions important for cofactor specif-
icity of CYP102A1 were identified. The corresponding mu-
tants were constructed by site-directed mutagenesis using the
Stratagene QuickChangeTM Kit (Stratagene, La Jolla, Ca,
USA) according to the manufacturerꢂs protocol. Mutations
were introduced to plasmid pT-USC1BM3[49] using the follow-
ing oligonucleotide primers:
Kinetic measurements were performed in sealed cuvettes at
least in triplicates to reduce standard errors.
Primer W1046S forward: 5’-gatacgcaaaagacgtgTCGgctggg-
taagaattc-3’
Coupling efficiency was measured by incubation of purified
CYP102A1 (3 mM), 50 U mLꢀ1 catalase with 0.4 mM cyclohex-
ane and 0.2 mM NADPH in a sealed cuvette. After total con-
sumption of NADPH the reaction mixture was extracted three
times with 300 mL 1-butanol or diethyl ether. The combined or-
ganic layers were dried over MgSO4 and analysed by quantita-
tive GC/MS. From the amount of substrate converted, the cou-
pling of NADPH oxidation to product formation can be de-
duced.
Primer W1046S reverse: 5’-gaattcttacccagcCGAcacgtctttt-
gcgtatc-3’
Primer W1046A forward: 5’-gatacgcaaaagacgtgGCGgct-
gggtaagaattc-3’
Primer W1046A reverse: 5’-gaattcttacccagcCGCcacgtctttt-
gcgtatc-3’
Primer R966D forward: 5’-gcttcataccgctttttctGACatgccaaa-
tcagccg-3’
Primer R966D reverse: 5’-cggctgatttggcatGTCagaaaaagc-
ggtatgaagc-3’
Substrate binding difference spectra were recorded using
standard procedures.[13,53]
Reductase activity was measured by standard cytochrome C
reduction assay as described previously.[50]
Myristic acid was oxidised in aqueous buffer by adding
20 mL of 5 mM myristic acid dissolved in DMSO to 930 mL of
1 mM CYP102A1 in 50 mM KPi, pH 7.5. The reaction was
started by addition of 50 mL 10 mM NADPH in 50 mM KPi,
pH 7.5. After 15 min the reaction mixture was acidified using
dilute HCl and extracted thrice with diethyl ether and analysed
as described in the section on product identification.
FDH activity assays were performed to estimate the stability
of FDH when stirred with cyclohexane. The same assay was
used during reactions to measure residual FDH activity. The
assay was performed by adding 50 mL FDH-containing solu-
Expression and Purification of CYP102A1 Mutants
The expression system pET28aþCYP102A1 was described
previously.[34] Mutations were introduced by PCR using the
QuikChangeTM site-directed mutagenesis kit from Stratagene
according to the manufacturerꢂs protocol.[51] High level protein
expression was achieved using a slightly modified version of
the protocol already published.[34] By incubating the starter cul-
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Adv. Synth. Catal. 2005, 347, 1090–1098

Catalytic Hydroxylation in Biphasic Systems
FULL PAPERS
tion to a cuvette containing 950 mL FDH reaction buffer
(50 mM KPi, 300 mM Naþ HCOOꢀ, 0.1 mM NADPþ,
pH 7.0) and measuring NADPH development at 340 nm.
decane. Analysis of this solid revealed it consisted of 90%
mono- and dihydroxymyristic acid. In contrast the dodecane
phase contained 95% myristic acid and only traces of hydroxy-
lated compounds.
Preparative Incubation in Biphasic Systems
Product Identification
The aqueous phase consisted of 250 mM sodium formate,
50 mM KPi, 10 mg mLꢀ1 BSA for stabilisation, 600 U mLꢀ1
catalase to destroy traces of hydrogen peroxide, 0.1 mM
NADPþ. A 1.5-fold excess of FDH over CYP102A1 was
used in the reaction to avoid limitation due to insufficient co-
factor recycling. Then the organic substrate was added. In the
cases of cyclohexane and octane as substrates the distilled sol-
vents were used as organic phase. In the case of myristic acid
conversion, the organic phase consisted of a 100 mM solution
of myristic acid in dodecane. Additionally 1% DMSO was add-
ed to the aqueous phase to enhance solubility of the organic
substrate. To start the reactions the respective CYP102A1 mu-
tant was added. For cyclohexane and octane conversions the
amount of CYP102A1 added is given in the results section. Hy-
droxylation of myristic acid was performed by 100 nmol
CYP102A1 A74G, F87V, L188Q (NADPH dependent) and al-
ternatively by 5 nmol CYP102A1 A74G, F87V, L188Q,
R966D, W1046S (NADH dependent).
Products were identified on a Shimadzu GC/MS-QP2010
equipped with a 30 m FS-Supreme column (internal diameter
0.25 mm, film thickness 0.25 mM) using helium as carrier gas
at linear velocity of 30 cm sꢀ1
.
Cyclohexane: 1) 508C for 5 min; 2) 50 to 2008C at 208C minꢀ1
;
3) 2 min at 2008C. The GC/MS signals of cyclohexane and cy-
clohexanol were calibrated for quantification using 10 stand-
ard concentrations ranging from 0.05 to 15 mM and fitted to
a quadratic function.
Octane: 1) 408C for 1 min; 2) 40 to 808C at 28C minꢀ1; 3)
1 min at 808C; 4) 80 to 2008C at 308C minꢀ1. Linear calibration
curves using in each case 10 standard concentrations of 2-, 3-
and 4-octanol and 2-, 3- and 4-octanone ranging from 0.01 to
1 mM were created.
Myristic acid: 1) 1758C for 1 min; 2) 175 to 2758C at 58C
minꢀ1; 3) 1 min at 2758C. Trimethylsilylated hydroxy deriva-
tives of myristic acid were identified by their characteristic
MS fragmentation patterns.
For hydroxylation of cyclohexane in a Labfors 3 l fermenter
(Infors AG, Bottmingen, Switzerland) the following param-
eters were used: 1.3 mmol CYP102A1 R47L, Y51F, stirrer
speed 350 rpm, aeration 0.5 L minꢀ1 and readjustment of pH
by 1 M KPi, 400 mM NaþHCOOꢀ pH 6. The biphasic reaction
emulsion consisted of 500 mL of aqueous phase and 500 mL of
cyclohexane. For online monitoring of the reaction parameters
IRIS software was applied. To collect evaporated educt and
product the fermenter was equipped with a cryo trap charged
with dry ice. To prevent foam formation, 100 mL of antifoam
(silicone oil) were added.
References
[1] Cytochrome P450: Structure, Mechanism and Biochemis-
try, (Ed.: P. R. E. Ortiz de Montellano), Plenum Press,
New York, 1995.
[2] D. F. Lewis, in: Cytochromes P450 Structure, Function
and Mechanism, (Ed.: D. F. Lewis), Taylor and Francis
Ltd., London, 1996, pp 115–167.
During the reaction course FDH activity assays were per-
formed. When no activity could be detected any more, FDH
was added (maximally once per reaction batch; half the
amount of FDH as in the beginning of the reaction).
[3] D. W. Nebert, F. J. Gonzalez, Annu. Rev. Biochem. 1987,
56, 945–993.
Monitoring of the reaction progress was achieved by taking
samples from the organic phase, drying them over MgSO4 and
analysis by quantitative GC/MS. In the case of myristic acid,
samples were taken from the organic and the aqueous phase
(1:1), acidified by addition of dilute HCl, extracted with dieth-
yl ether and evaporated to dryness. Trimethylsilylation was
achieved by dissolving the solid in N,O-bis(trimethylsilyl)tri-
fluoroacetamide and 1% trimethylchlorosilane and incubation
at 608C for 30 min in a sealed vial.
For quantitative determination of total products, the reac-
tion mixture was extracted three times with diethyl ether, dried
and analysed by GC/MS. For cyclohexanol the phase partition-
ing between cyclohexane and the aqueous reaction medium
was determined. Below 20 g Lꢀ1 a linear relation of cyclohex-
anol in aqueous and organic phase was found: [cyclohexanol in
cyclohexane]¼(0.936 þ/- 0.011) [cyclohexanol in aqueous
phase].
[4] K. Petzoldt, K. Annen, H. Laurent, R. Wiechert, (Scher-
ing Aktiengesellschaft, Berlin, Germany), US Patent
4,353,985, 1982.
[5] S. Picataggio, T. Rohrer, K. Deanda, D. Lanning, R. Rey-
nolds, J. Mielenz, L. D. Eirich, Biotechnology (NY) 1992,
10, 894–898.
[6] V. Reipa, M. P. Mayhew, V. L. Vilker, Proc. Natl. Acad.
Sci. USA 1997, 94, 13554–13558.
[7] J. R. Falck, Y. K. Reddy, D. C. Haines, K. M. Reddy,
U. M. Krishna, S. Graham, B. Murry, J. A. Peterson, Tet-
rahedron Lett. 2001, 42, 4131–4133.
[8] K. Hofstetter, J. Lutz, I. Lang, B. Witholt, A. Schmid,
Angew. Chem. Int. Ed. 2004, 43, 2163–2166.
[9] V. B. Urlacher, S. Lutz-Wahl, R. D. Schmid, Appl. Micro-
biol. Biotechnol. 2004, 64, 317–325.
[10] V. Urlacher, R. D. Schmid, Curr. Opin. Biotechnol. 2002,
13, 557–564.
[11] L. O. Narhi, A. J. Fulco, J. Biol. Chem. 1986, 261, 7160–
7169.
[12] L. P. Wen, A. J. Fulco, J. Biol. Chem. 1987, 262, 6676–
6682.
For myristic acid the acidified (pH 2) reaction mixture was
extracted thrice with diethyl ether. The combined organic lay-
ers were evaporated under vacuum until only high-boiling do-
decane was left. At this stage a white solid had formed. The sol-
id was separated by centrifugation and washed twice with do-
Adv. Synth. Catal. 2005, 347, 1090–1098
asc.wiley-vch.de
ꢁ 2005 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
1097

Steffen C. Maurer et al.
FULL PAPERS
[13] G. Truan, M. R. Komandla, J. R. Falck, J. A. Peterson,
Arch. Biochem. Biophys. 1999, 366, 192–198.
[14] T. W. Ost, C. S. Miles, J. Murdoch, Y. Cheung, G. A.
Reid, S. K. Chapman, A. W. Munro, FEBS Lett. 2000,
486, 173–177.
[35] J. T. Wu, L. H. Wu, J. A. Knigh, Clin. Chem. 1986, 32,
314–319.
[36] H. Li, T. L. Poulos, Nat. Struct. Biol. 1997, 4, 140–146.
[37] M. A. Noble, C. S. Miles, S. K. Chapman, D. A. Lysek,
A. C. MacKay, G. A. Reid, R. P. Hanzlik, A. W. Munro,
Biochem. J. 1999, 339, 371–379.
[38] A. B. Carmichael, L. L. Wong, Eur. J. Biochem. 2001,
268, 3117–3125.
[39] Q. S. Li, U. Schwaneberg, M. Fischer, J. Schmitt, J. Pleiss,
S. Lutz-Wahl, R. D. Schmid, Biochim. Biophys. Acta
2001, 1545, 114–121.
[40] L. D. Gorsky, D. R. Koop, M. J. Coon, J. Biol. Chem.
1984, 259, 6812–6817.
[41] V. I. Tishkov, A. G. Galkin, V. V. Fedorchuk, P. A. Savit-
sky, A. M. Rojkova, H. Gieren, M. R. Kula, Biotechnol.
Bioeng. 1999, 64, 187–193.
[42] F. Ahmed, E. H. Al-Mutairi, K. L. Avery, P. M. Cullis,
W. U. Primrose, G. C. K. Roberts, C. L. Willis, Chem.
Commun. 1999, 2049–2050.
[43] C. L. Elmore, T. Porter, J. Biol. Chem. 2002, 50, 48960–
48964.
[44] O. Dçhr, M. J. I. Paine, T. Friedberg, G. C. K. Roberts,
C. R. Wolf, Proc. Natl. Acad. Sci. USA, 2001, 98, 81–86.
[45] L. A. Kaysser, S. Eiben, R. D. Schmid, V. B. Urlacher, in
preparation.
[46] V. I. Tishkov, A. G. Galkin, G. N. Marchenko, O. A.
Egorova, D. V. Sheluho, L. B. Kulakova, L. A. Demen-
tieva, A. M. Egorov, Biochem. Biophys. Res. Commun.
1993, 192, 976–981.
[47] F. Zambiachini, P. Pasta, G. Carrea, S. Colonna, N. Gag-
gero, J. M. Woodley, Biotechnol. Bioeng. 2002, 78, 489–
496.
[48] J. Lutz, V. V. Mozhaev, Y. L. Khmelnitsky, B. Witholt, A.
Schmid, J. Mol. Catal. B: Enzymatic. 2002, 19–20, 177–
187.
[49] U. Schwaneberg, A. Sprauer, C. Schmidt-Dannert, R. D.
Schmid, J. Chromatogr. 1999, 848, 149–159.
[50] A. H. Phillips, R. G. Langdon, J. Biol. Chem. 1962, 237,
2652–2660.
[51] M. A. Vandeyar, M. P. Weiner, C. J. Hutton, C. A. Batt,
Gene 1988, 65, 129–133.
[15] O. Lentz, V. Urlacher, R. D. Schmid, J. Biotechnol. 2004,
108, 41–49.
[16] M. Budde, S. C. Maurer, R. D. Schmid, V. B. Urlacher,
Appl. Microbiol. Biotechnol. 2004, 66, 180–186.
[17] M. C. Gustafsson, O. Roitel, K. R. Marshall, M. A. No-
ble, S. K. Chapman, A. Pessegueiro, A. J. Fulco, M. R.
Cheesman, C. von Wachenfeldt, A. W. Munro, Biochem-
istry 2004, 43, 5474–5487.
[18] Q. S. Li, U. Schwaneberg, P. Fischer, R. D. Schmid,
Chemistry 2000, 6, 1531–1536.
[19] D. Appel, S. Lutz-Wahl, P. Fischer, U. Schwaneberg,
R. D. Schmid, J. Biotechnol. 2001, 88, 167–171.
[20] E. T. Farinas, U. Schwaneberg, A. Glieder, F. H. Arnold,
Adv. Synth. Catal. 2001, 343, 601–606.
[21] O. Lentz, Q. S. Li, U. Schwaneberg, S. Lutz-Wahl, P.
Fischer, R. D. Schmid, J. Mol. Cat. B 2001, 15, 123–133.
[22] Q. S. Li, J. Ogawa, R. D. Schmid, S. Shimizu, Appl. Envi-
ron. Microbiol. 2001, 67, 5735–5739.
[23] Q. S. Li, J. Ogawa, R. D. Schmid, S. Shimizu, FEBS Lett.
2001, 508, 249–252.
[24] A. Glieder, E. T. Farinas, F. H. Arnold, Nat. Biotechnol.
2002, 20, 1135–1139.
[25] M. W. Peters, P. Meinhold, A. Glieder, F. H. Arnold, J.
Am. Chem. Soc. 2003, 125, 13442–13450.
[26] T. Seng Wong, F. H. Arnold, U. Schwaneberg, Biotech-
nol. Bioeng. 2004, 85, 351–358.
[27] S. Schneider, M. G. Wubbolts, G. Oesterhelt, D. San-
glard, B. Witholt, Biotechnol. Bioeng. 1999, 64, 333–341.
[28] P. C. Cirino, F. H. Arnold, Angew. Chem. Int. Ed. 2003,
42, 3299–3301.
[29] R. W. Estabrook, M. S. Shet, K. Faulkner, C. W. Fisher,
Endocr. Res. 1996, 22, 665–671.
[30] B. D. Fleming, Y. Tian, S. G. Bell, L. L. Wong, V. Urlach-
er, H. A. Hill, Eur. J. Biochem. 2003, 270, 4082–4088.
[31] A. K. Udit, F. H. Arnold, H. B. Gray, J. Inorg. Biochem.
2004, 98, 1547–1550.
[32] F. Hollmann, B. Witholt, A. Schmid, J. Mol. Cat. B 2002,
791, 1–10.
[52] T. Omura, R. J. Sato, J. Biol. Chem. 1964, 239, 2370–
2378.
[33] S. B. Lamb, D. C. Lamb, S. L. Kelly, D. C. Stuckey, FEBS
Lett. 1998, 431, 343–346.
[53] K. C. Leibman, A. G. Hildebrandt, R. W. Estabrook, Bio-
chem. Biophys. Res. Commun. 1969, 36, 789–794.
[34] S. Maurer, V. Urlacher, H. Schulze, R. D. Schmid, Adv.
Synth. Catal. 2003, 345, 802–810.
1098
ꢁ 2005 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim
asc.wiley-vch.de
Adv. Synth. Catal. 2005, 347, 1090–1098