Cytochrome P450 168A1 from Pseudomonas aeruginosa is involved in the hydroxylation of biologically relevant fatty acids

The cytochrome P450 CYP168A1 from Pseudomonas aeruginosa was cloned and expressed in Escherichia coli followed by purification and characterization of function. CYP168A1 is a fatty acid hydroxylase that hydroxylates saturated fatty acids, including myristic (0.30 min-1), palmitic (1.61 min-1) and stearic acids (1.24 min-1), at both the ω-1- and ω-2-positions. However, CYP168A1 only hydroxylates unsaturated fatty acids, including palmitoleic (0.38 min-1), oleic (1.28 min-1) and linoleic acids (0.35 min-1), at the ω-1-position. CYP168A1 exhibited a catalytic preference for palmitic, oleic and stearic acids as substrates in keeping with the phosphatidylcholine-rich environment deep in the lung that is colonized by P. aeruginosa.


Introduction
Pseudomonas aeruginosa is an opportunistic pathogen, and the leading cause of chronic lung infection in cystic fibrosis patients [1]. P. aeruginoisa uses the lung surfactant (which is essential for normal breathing, preventing alveoli collapse and acting as a system of lung defense in the lungs) as a source of nutrients allowing it to colonize a large portion of the lungs. This causes airway plugging and surface damage to epithelial cells [1]. The lung surfactant largely consists of a class of phospholipids called phosphatidylcholine. This phospholipid is a source of fatty acids, which are released during degradation by lipases and phospholipases, which are excreted by P. aeruginosa [2][3][4]. These released fatty acids could be broken down through the fatty acid degradation pathway via the β-oxidation cycle [5] to be used as a source of energy or they could be taken up by the cell to be used for other cellular processes.
For instance, upon cellular uptake in other organisms, fatty acids can be metabolized by cytochrome P450 enzymes (CYPs) to produce hydroxy fatty acids, which, in turn, can be used for a variety of physiological functions, including diacid formation [5,6], sophorolipid a1111111111 a1111111111 a1111111111 a1111111111 a1111111111 production [7], and the synthesis of cutin and suberin in plants [8]. While cytochromes P450 (CYPs) are not directly involved in the β-oxidation cycle of fatty acid degradation, they are essential in forming or initiating the formation of hydroxy fatty acids and/or diacids. These molecules can then act as an energy source as they can be degraded by the β-oxidation cycle [5,6].
It is unlikely, because of their cellular localization, that fatty acid hydroxylating CYPs in P. aeruginosa will interact directly with the lung surfactant in order to have access to phosphatidylcholine. Therefore, in order for these CYPs to access the fatty acids released from phosphatidylcholine, P. aeruginosa expresses all the relevant genes involved in phosphatidylcholine degradation (lipases: LipA and LipC, and phospholipases: PlcH and PlcR). The expression of these genes results in fatty acids, glycerol and phosphorylcholine being released. The individual constituents are further imported and degraded, via high levels of expression of many genes involved in the fatty acid degradation pathway suggesting that P. aeruginosa may utilize phosphatidylcholine as one of the major nutrient sources in vivo [2] and could also be made available for interaction with the CYPs.
In this study, we have investigated CYP168A1 from P. aeruginosa, the sole member of its cytochrome P450 family, to establish whether this enzyme is able to catalyze the hydroxylation of fatty acids. We have cloned, expressed and purified CYP168A1 and have successfully demonstrated the enzyme's ability to hydroxylate biologically relevant fatty acids at the sub-terminal carbons.

Chemicals
Growth media, ampicillin, 5-aminolevulenic acid and isopropyl-β-D-thiogalactopyranoside (IPTG) were purchased from Formedium, Ltd. (Hunstanton, UK). Chemicals used in the preparation of phosphate buffers were purchased from Fisher Scientific (Loughborough, UK). Voriconazole was purchased from Discovery Fine Chemicals (Dorset, UK). Palmitoleic acid (C16:1) was purchased from Tokyo Chemical Industry UK Ltd (Oxford, UK). All other fatty acids and chemicals were purchased from Sigma-Aldrich (Poole, UK), unless otherwise stated.

Heterologous expression and purification of CYP168A1 protein
The CYP168A1 gene (UniProt accession number Q9I107) was synthesized by Eurofins MWG Operon (Ebersberg, Germany) including nucleotide sequence optimization for expression in E. coli. The gene was designed to contain the triplet GCT, coding for alanine as the second amino acid, to aid expression in E. coli and a C-terminus hexahistidine tag to facilitate purification by affinity chromatography using Ni 2+ -NTA agarose. In addition an NdeI restriction site was incorporated at the 5' end and a HindIII restriction site at the 3' end of the gene. The gene was cloned using the NdeI and HindIII restriction sites into pET17b and transformed into BL21(DE3)pLysS cells under ampicillin and chloramphenicol selection. Transformants were used to inoculate Terrific broth containing ampicillin and grown at 37˚C and 180 rpm for 6 hours. 1 mM IPTG and 1 mM 5-aminolevulenic acid were added for induction prior to expression. CYP168A1 was expressed at 25˚C and 130 rpm for 20 hours. Cells were harvested (10 min at 3000 x g), re-suspended in 0.1 M potassium phosphate buffer (pH 7.4) and stored at -80˚C overnight. Samples were thawed and spun at 140000 x g for 1 hour at 4˚C to recover the solubilized protein in the supernatant. CYP168A1 was purified using Ni 2+ -NTA agarose (Qiagen) and eluted in 0.1 M Tris-HCl (pH 8.1) containing 25% (w/v) glycerol and 1% (w/v) L-histidine. SDS-polyacrylamide gel electrophoresis was undertaken to assess protein purity.

Determination of cytochrome P450 protein concentration
Reduced carbon monoxide difference spectroscopy was performed (light path, 10 mm) according to the method of Estabrook et al., 1972 [9] to determine cytochrome P450 protein concentration using an extinction coefficient of 91 mM -1 cm -1 at 450 nm [10]. Absolute spectra were determined from 700 nm to 250 nm (light path, 4.5 mm). The heme concentration of the purified CYP168A1, diluted with 10 mM potassium phosphate (pH 7.4), was determined by measuring the Soret peak at 417 nm using an extinction coefficient of 125 mM -1 cm -1 [11] and the total protein concentration was determined by measurement of the absorbance at 205 nm using an extinction coefficient of 31 ml mg -1 cm -1 [12] from which the percentage heme incorporation was calculated. The Reinheitszahl (Rz) ratio of absorbance due to the heme Soret peak at 417 nm and that due to the absorbance of the apoprotein was also determined as a primary indicator of enzyme purity and heme incorporation [13]. All spectral determinations were made using a Hitachi U-3310 UV-visible spectrophotometer (San Jose, CA).

Fatty acid binding studies
Fresh supplies of fatty acids were purchased prior to commencing ligand binding and catalysis studies. Stock solutions containing 0.25 mg ml -1 myristic acid (C14:0), palmitic acid (C16:0), stearic acid (C18:0) and oleic acid (C18:1) were prepared in dimethylformamide along with 0.5 mg ml -1 palmitoleic acid (C16:1), 0.1 mg ml -1 linoleic acid (C18:2) and 10 mg ml -1 arachidonic acid (C20:4). These stock fatty acid solutions were progressively titrated against 5 μM of CYP168A1 protein in 0.1 M Tris-HCl (pH 8.1) buffer containing 25% (w/v) glycerol using quartz semi-micro cuvettes with equivalent volumes of dimethylformamide added to the cytochrome P450-containing reference cuvette. The absorbance difference spectra from 500 nm to 350 nm were determined after each incremental addition of fatty acid. Ligand saturation curves were constructed from the change in absorbance (ΔA peak-trough ) against fatty acid concentration. The dissociation constant for the fatty acid-CYP168A1 complex (K s ) was determined by nonlinear regression (Levenberg-Marquardt algorithm) using a rearrangement of the Morrison equation [14] and the Michaelis-Menten equation. The magnitude of the spin state change for type I difference spectra was calculated from ΔA 390-420 using an extinction coefficient of 100 mM -1 cm -1 [15]. All fatty acid binding experiments were undertaken in quadruplicate.
In between ligand binding determinations, the quartz cuvettes were washed with deionized water and then soaked for 30 min in 2-propanol at room temperature to desorb any residual fatty acids from the cuvette surfaces, followed by rinsing a further three-times with 2-propanol and then deionized water prior to drying.

Fatty acid reconstitution assays
The reconstitution assay system contained 0.25 μM CYP168A1, 2.5 μM spinach ferredoxin (Sigma-Aldrich F3013), 0.25 μM spinach ferredoxin-NADP + reductase (Sigma-Aldrich F0628), 50 μM dilaurylphosphatidylcholine (DLPC), 100 μM fatty acid, 4 mM glucose-6-phosphate, 3 U/ml yeast glucose-6-phosphate dehydrogenase and 0.1 M potassium phosphate (pH 7.4). Assay mixtures were incubated at 37˚C for 5 min prior to initiation with 4 mM β-NADPH-Na 4 and then incubated for a further 2.5 hours at the same temperature. Fatty acids and their hydroxylated products were recovered by extraction with dichloromethane and dried in a vacuum centrifuge. TMS derivatisation of the samples and analysis by GCMS were performed as previously described [16]. Percentage product formation was calculated from the GC peak areas of the fatty acid and the hydroxylated metabolites, with compound identities confirmed by mass fragmentation patterns of fatty acid standards.
To test the effect of the azole antifungal drugs miconazole, tebuconazole and voriconazole on CYP168A1 turnover, the reconstitution assays were repeated as described above using oleic acid as the substrate, except 2 μM of azole dissolved in DMF was also present. Control assays contained no azole, but an equivalent volume of DMF. Experiments were undertaken in duplicate.
CYP168A1 reconstitution assays were also performed using 50 μM cholesterol, cholesta-4-ene-3-one, progesterone and testosterone as potential substrates. For these assays the CYP168A1 concentration was increased to 2 μM in the presence of 2.5 μM spinach ferredoxin, 0.25 μM spinach ferredoxin-NADP + reductase, 50 μM DLPC, 4 mM glucose-6-phosphate, 3 U/ml yeast glucose-6-phosphate dehydrogenase, 0.1 M potassium phosphate (pH 7.4) and 4 mM β-NADPH-Na 4 , followed by 3 hours incubation at 37˚C. Steroid and sterol compounds were extracted with ethyl acetate (2 x 3 ml), dried using a vacuum centrifuge and derivatized firstly with methoxamine followed by silyation using BSTFA-TMCS and analyzed by GCMS (57). Metabolites were identified from GC traces and MS fragmentation patterns compared against positive controls for CYP mediated hydroxylation reactions performed in our laboratory.

Azole binding studies
The azole antifungal drugs miconazole, tebuconazole and voriconazole were used in binding studies with CYP168A1 in accordance with previously described methods [17,18]. Stock solutions of these azoles (0.75-10 mg ml -1 ) were prepared in DMF and progressively titrated against 2 μM CYP168A1 in 0.1 M Tris-HCl (pH 8.1) buffer containing 25% (w/v) glycerol. Equivalent volumes of DMF were added to a reference cuvette containing 2 μM of CYP168A1. The difference spectrum from 500 nm to 350 nm was determined after each incremental addition of azole. Binding saturation curves were constructed from the ΔA peak-trough against azole concentration. The dissociation constant (K d ) of the enzyme-azole complex was determined by nonlinear regression (Levenberg-Marquardt algorithm) using a rearrangement of the Morrison equation [14]. All binding experiments were undertaken in duplicate.

MIC determinations
P. aeruginosa strains DSMZ 22644 and ATCC 39324 were grown in LB media overnight. Optical density at 520 nm of the overnight cultures was measured using a spectrophotometer. Cultures were adjusted to give an optical density of approximately 0.2, which is equivalent to a cell count of approximately 1 x 10 9 . The cells were pelleted and resuspended in the same volume of water. Further dilutions were made in M9 media with oleic acid used as the carbon source (instead of glucose) to give approximately 2 x 10 5 cells. Stock concentrations of tebuconazole, miconazole and voriconazole were prepared in dimethyl sulfoxide (12.8, 6.4, 3.2, 1.6, 0.8, 0.4, 0.2, 0.1, 0.05 and 0.025 mg ml -1 ). These stock azole solutions were diluted ten-fold in fresh LB media and then diluted a further ten-fold with inoculum in microtiter plate wells. This gave final azole concentrations of 128, 64, 32, 16, 8, 4, 2, 1, 0.5 and 0.25 μg ml -1 . The microtiter plates were incubated at 37˚C for 24 hours. Following this initial incubation period, 20 μl of 0.2% Resazurin was added to each microtiter plate well. The plates were incubated for a further 48 hours at 37˚C before being read. A color change from purple to pink indicated the presence of respiring cells. Each azole MIC determinations were performed in triplicate and scored manually.

Data analysis
Curve-fitting of ligand binding data were performed using the computer program Quantum-Soft ProFit (version 6.1.12). Phylogenetic analysis of CYP168A1 was performed using the Uni-Prot BLAST online software resource (http://www.uniprot.org/blast/). Amino acid sequence alignments were performed using ClustalX version 1.81 software (http://www.clustal.org/).

Heterologous expression of CYP168A1
Overexpression of CYP168A1 in E. coli resulted in protein yields of~900 nmol per liter of culture. SDS-polyacrylamide gel electrophoresis, following Ni 2+ -NTA agarose purification, confirmed that CYP168A1 was over 90% pure as judged by staining intensity with Coomassie brilliant blue R-250. The absolute spectrum of CYP168A1 ( Fig 1A) was characteristic of a ferric cytochrome P450 enzyme that had been isolated predominately in the low-spin state with a Soret peak at 417 nm [19,20]. The A 393-470 /A 417-470 value was 0.412, where 0.4 is indicative of 100% low-spin occupancy and 2.0 indicative of 100% high-spin occupancy [21], confirming CYP168A1 to be over 95% low-spin in the oxidized resting state. The observed A 417 /A 280 for CYP168A1 was 1.26, which was within the expected range of 1 and 2 for a cytochrome P450 [13]. Heme incorporation was 77 to 81% calculated from measurements at A 205 and A 417 [11,12] with a specific heme content of 15.3 nmol mg -1 protein comparable to the expected value of 17 to 20 nmol mg -1 for cytochrome P450 enzymes [13]. The dithionite-reduced carbon monoxide difference spectrum for CYP168A1 (Fig 1B) was characteristic of CYPs isolated in their native state exhibiting a red-shifted Soret peak at 450 nm [9,10].

Fatty acids are able to bind to CYP168A1
The heme prosthetic group of CYPs exists in equilibrium between the low-spin (hexacoordinate) form, characterized by a heme Soret peak at~418 nm, and the high-spin (pentacoordinate) form, characterized by a heme Soret peak at~393 nm. The resting state of most CYPs is predominantly the low-spin ferric form [23]. Ligand binding can perturb the spin state equilibrium in two main ways. Substrates and other ligands can bind to the CYP (though the ligands themselves do not directly coordinate to the heme) resulting in the displacement of the axial-ligated water molecule from the heme ferric ion and a change in spin state from low-to high-spin. The magnitude of the spin state change observed is dependent on both the CYP and the ligand. For example, lauric acid binding to Streptomyces peucetius CYP147F1 causes 95% of the enzyme molecules to undergo a low-to high-spin state transition [24], whereas for eukaryotic CYP51 enzymes the spin state changes associated with substrate binding rarely exceed 10% [25]. This low-to high-spin transition gives rise to a type I difference spectrum that is characterized by a spectral peak at~390 nm and trough at~420 nm. Other ligands can directly coordinate to the heme ferric ion (commonly through an aromatic nitrogen atom) [19,26], for example the binding of azole antifungals to CYP51 enzymes [27]. The resultant complexes are low-spin (hexacoordinate) and often result in a red-shift of the Soret peak from 418 nm to 425-434 nm [19]. This direct coordination of the ligand to the heme ferric ion gives rise to a type II difference spectrum [19]. The spectral peak varies from 425 nm (if the CYP was 100% in the high-spin state before ligand binding) to 432 nm (if the CYP was 100% in the low-spin state) with respective troughs of 390 nm and 410 nm. For CYPs of mixed spin state the peaks and troughs will be at intermediate wavelengths.
It was important to optimize the stock fatty acid concentrations so as to obtain sufficient ascendant points on the titration curve before reaching ligand saturation to facilitate curve fitting of data and accurate K s determination, especially as the type I difference spectra for some fatty acids (notably stearic, palmitoleic and oleic acids) started to dissipate at higher ligand concentrations (Fig 3). This may be caused by either slow aggregation of the CYP168A1-fatty acid complex (although no visible precipitation was observed) or a perturbation of the spinstate equilibrium of the CYP168A1-substrate complex so that some molecules transition to the low-spin state by partial coordination of a water molecule as the sixth axial heme ligand. This phenomenon was evident in the ligand saturation curve for palmitic acid with CYP168A1 reported by Tooker et al [28].
This phenomenon of partial dissipation of type I binding spectra at higher ligand concentrations had previously been observed for cholesterol (but not 4-cholesten-3-one) binding to CYP125 from Mycobacterium tuberculosis [29]. Capyk et al [29] demonstrated this phenomenon was due to the ligand solvent (10% w/v 2-hydroxypropyl-β-cyclodextrin), as when the solvent was changed (to 25 mM EDTA-bridged β-cyclodextrin dimer) the premature dissipation of the type I difference spectrum was no longer observed with cholesterol. As the partial dissipation of type I binding spectra appears to be both ligand-and solvent-specific, both molecules must interact with the CYP protein to cause this effect. When the fatty acid solvent was changed from DMF to ethanol the partial dissipation of the type I binding spectra still occurred at higher ligand concentrations.
The binding saturation curves for all the fatty acids, except arachidonic acid, were best fit using a rearrangement of the Morrison equation [14] ( Table 1, Fig 3) to calculate the dissociation constant of the substrate-CYP168A1 complex (K s ), indicating tight binding to CYP168A1. The Michaelis-Menten equation gave the best fit for arachidonic acid binding (Fig 4). Stearic and linoleic acids bound the tightest to CYP168A1 with apparent K s values under 0.05 μM, followed by palmitic acid (K s 0.12 μM), then oleic and palmitoleic acids (K s 0.15 μM) and myristic acid (K s 0.25 μM), with arachidonic acid binding having the lowest affinity (K s~4 60 μM). The fold-difference in apparent K s values compared to stearic and linoleic acids were~2-,~3-,~3-~5-, and~9160-fold for palmitic, oleic, palmitoleic, myristic, and arachidonic acids, respectively. The high K s value for arachidonic acid suggested it would be a poor CYP168A1 substrate. Therefore, based on apparent ligand binding affinities, CYP168A1 exhibits a preference for C16 to C18 fatty acids. The magnitude of the low-to high-spin transitions induced were 23, 58, 37, 66, 55, 28 and 32% for myristic, palmitic, palmitoleic, stearic, oleic, linoleic and arachidonic acids, respectively, based on the observed ΔA max values (Table 1).
CYP168A1 was also able to hydroxylate the unsaturated fatty acids palmitoleic acid, oleic acid and linoleic acid, but only at the ω-1-carbon. Catalytic turnover with oleic acid was similar to that of palmitic acid in terms of production of the ω-1-hydroxy metabolite ω-1-hydroxyoleic acid (1.28 min -1 ) (Fig 5D, Table 2) and was 3.4-and 3.7-fold greater than the relative ω-1-hydroxy metabolites produced when palmitoleic and linoleic acids were used as substrates. Lower catalytic turnovers were observed with palmitoleic and linoleic acids (0.38 and 0.35 min -1 , respectively) although both these fatty acids bound tightly to CYP168A1 (K s values of 0.155 μM and >0.05 μM, respectively). Reconstitution assays with arachidonic acid showed no detectable product formation and a large K s value of 458 μM was obtained for arachidonic acid with CYP168A1 (Table 1). No C-C cleavage of fatty acyl chains was detected in the assay products, suggesting that CYP168A1 was not a CYP107H homolog.
Catalytic turnover established the order of substrate preference to be palmitic acid (highest turnover) followed by oleic acid, stearic acid, palmitoleic acid, linoleic acid and finally myristic acid with the lowest turnover (5.3-fold lower than palmitic), whilst arachidonic acid was catalytically inactive. In contrast, the fatty acid K s data suggested the order of substrate preference would have been stearic and linoleic acids, with the lowest K s values, followed by oleic acid, palmitoleic acid, myristic acid, and finally arachidonic acid with a 9000-fold larger K s value than stearic acid. Both stearic and linoleic acids gave similar apparent K s values with CYP168A1 and yet the catalytic turnover observed with stearic acid was 3.5-fold greater than that for linoleic acid. Further investigations are required in order to determine the mechanisms responsible for the observed differences in CYP168A1 catalytic turnover between substrates.
CYP168A1 reconstitution assays using cholesterol, cholesta-4-ene-3-one, progesterone and testosterone as potential substrates gave no oxygenated products, suggesting CYP168A1 is a fatty acyl hydroxylase. Type I difference spectra were observed for all seven fatty acids with 5 μM CYP168A1. Mean K s values from four replicates were calculated using a rearrangement of the Morrison equation [14] and are shown ± standard deviations, except for arachidonic acid where the Michaelis-Menten equation was used. a The K s values for these two fatty acids were below the lower accuracy limit of the Morrison equation of 0.05 μM (1% the concentration of the enzyme) [61]. b Path length of cuvettes used with arachidonic acid were 4.5 mm compared to 10 mm used with the other fatty acids.
Binding saturation was not achieved at 660 μM arachidonic acid.
https://doi.org/10.1371/journal.pone.0265227.t001 Gas chromatograms for TMS-derivatized assay metabolites obtained when palmitic acid (C) and oleic acid (D) were used as substrates are shown. Peak (i) corresponds to palmitic acid, peak (ii) to ω-2-hydroxypalmitic acid, peak (iii) to ω-1-hydroxypalmitic acid, peak (iv) to oleic acid and peak (v) to ω-1-hydroxyoleic acid (all TMS-derivatives). Mass fragmentation patterns of the palmitic acid products can be found in the supporting information (S1 Fig). Other minor GC peaks were identified as impurities present in the source fatty acids. https://doi.org/10.1371/journal.pone.0265227.g005 Azoles are able to bind to CYP168A1, but they have no effect on P. aeruginosa growth CYP168A1 was titrated against the azole antifungal drugs tebuconazole, voriconazole and miconazole. The enzyme was able to bind each of the three azoles, eliciting type II difference spectra (Fig 6), with apparent dissociation constants (K d ) for the azole-CYP168A1 complexes of 0.18 ±0.01, 2.05 ±0.54 and 0.19 ±0.06 μM for tebuconazole, voriconazole and miconazole,  6. Binding of azole antifungals to CYP168A1. Miconazole (0.75 mg ml -1 ), tebuconazole (5 mg ml -1 ) and voriconazole (10 mg ml -1 ) were progressively titrated against 2 μM CYP168A1 in quartz semi-micro cuvettes of path length 4.5 mm. After each 1 μl addition of azole solution the difference spectrum was measured against a CYP168A1-containing reference cuvette in which an equivalent volume of DMF was added. The cumulative type II difference spectra are shown along with the ligand saturation curves which were fitted using a rearrangement of the Morrison equation [14].
https://doi.org/10.1371/journal.pone.0265227.g006 respectively, calculated from the ligand saturation curves (Fig 6). This was in contrast to K d values for tebuconazole, voriconazole and miconazole obtained with Candida albicans CYP51 of 0.036, 0.010 and 0.026 μM, respectively [27,31]. The addition of 2 μM azoles to reconstitution assays containing 0.25 μM CYP168A1 showed they elicited no effect on the catalytic activity towards oleic acid when compared against the DMF control. However, the presence of 0.5% DMF in the CYP168A1 assay system caused a 10-fold reduction in oleic acid turnover. Tebuconazole, voriconazole and miconazole were used in minimum inhibitory concentration (MIC) determinations with P. aeruginosa strains DMZ 22644 and ATCC 39324 grown on oleic acid. In all cases, the azoles at concentrations up to 128 μg ml -1 had no inhibitory effect on P. aeruginosa growth as the Resazurin indicator turned pink in all wells, indicating cell respiration.

Discussion
Fatty acids are essential for cell life. Hydroxylation of these molecules can result in the formation of hydroxy fatty acids that can act as signaling molecules and can be used to produce more complex molecules, such as diacids [7,32]. Therefore, identification and characterization of enzymes involved in fatty acid metabolism is important, particularly in pathogens, such as P. aeruginosa, where insight into their function can be used to identify new potential targets for novel inhibitors and teach us more about the mechanism of fatty acid degradation.
CYP168A1 was able to bind a range of biologically relevant fatty acids. The K s range of <0.05 to 0.25 μM for fatty acids that were catalytically active with CYP168A1 were similar to those observed previously for Streptomyces peucetius CYP147F1 [24], Sorangium cellulosum CYP267A1 [33], Streptomyces coelicolor A3(2) CYP105D5 [34], and some studies with Bacillus megaterium CYP102A1 [35]. The K s values in this study were similar to those recently reported for P. aeruginosa CYP168A1 [28], with the exception for stearic acid which was over 6-fold lower (<0.05 μM compared to 0.327 μM) and arachidonic acid which was over 400-fold higher (458 μM compared to 0.96 μM) with the latter mainly due to the atypical difference spectrum observed with arachidonic acid in this study. Higher K s values have been observed (8 to 30 μM) with Bacillus subtilis CYP107H1 [36] and substantially higher K s values (19 to 1065 μM) observed with Bacillus subtilis CYP102A2 and CYP102A3 [37].
CYP168A1 fatty acid turnover rates were similar to those obtained with Sphingomonas paucimobilis CYP152B1 [38], with both CYPs giving highest turnovers with palmitic acid, and with Sorangium cellulosum CYP267A1 [33], except the highest turnover was obtained for capric acid with CYP267A1. Fatty acid turnover rates with CYP168A1 were~10-fold higher than those observed with Mycobacterium marinum CYP153A16 and Marinobacter aquaeolei CYP153A [39] and turnover of palmitic acid was 23-fold greater than observed with Mycobacterium tuberculosis CYP124 [40]. The CYP168A1 turnover numbers obtained in this study, although being relatively low for cytochrome P450 monooxygenases, were similar to those reported by Tooker et al [28] of 0.138 min -1 with lauric acid and 0.222 min -1 with arachidonic acid, however, in this study no catalytic activity was observed with arachidonic acid. At present we cannot account for this difference, although solubility / bioavailability of arachidonic acid in the in vitro reconstitution assay may be a contributory factor. Interestingly, Tooker et al [28] CYP168A1 K m for lauric acid was 25-fold higher than the K s value, suggesting that other parameters besides substrate binding affinity were significant contributors to the observed K m value.
Not all bacterial fatty acid hydroxylating CYPs only hydroxylate unsaturated fatty acids at one position. For instance, CYP105D5 can hydroxylate oleic acid at multiple positions with a preference for the ω-1-position [34]. CYP102A1 can also cause the epoxidation of unsaturated fatty acids, but no epoxidation was observed when CYP168A1 was used in reconstitution assays with unsaturated fatty acids, such as oleic acid and linoleic acid. In the presence of arachidonic acid, CYP102A1 is able to both hydroxylate at the ω-2-carbon and cause epoxidation between carbons 14 and 15 [46], whereas CYP168A1 showed no turnover of arachidonic acid in this study.
In all cases, CYP168A1 was only able to hydroxylate the fatty acids at the sub-terminal carbons, with no activity observed at the terminal, ω-carbon, unlike other bacterial fatty acid hydroxylating CYPs, such as CYP124 (36) and CYP119 from Sulfolobus acidocaldarius [47], and those from eukaryotes, such as members of the CYP52 family in yeast [32,41] and the CYP4 family in mammals [48][49][50][51]. Therefore, CYP168A1 is unlikely to be involved in the production of α,ω-diacids. In order to hydroxylate fatty acids at the ω-carbon, CYP4B1 requires an unusual heme-polypeptide ester in the active site to mediate this energetically disfavored process [52].
As CYP168A1 does not catalyze the terminal ω-hydroxylation of fatty acids, it will not be involved in the production of α,ω-diacids. However, ω-1-fatty acids can be further converted to ω-1-oxo fatty acids. In Legionella, long chain ω-1-oxo fatty acids, also hydroxylated at the αcarbon (thus producing α-hydroxy, ω-1-oxo fatty acids), are constituents of the cell wall [53]. Also ω-1-oxo fatty acids can be further converted to ω-1-oxo dicarboxylic acids [54]. CYP168A1 does not metabolize sterols or steroids, which suggests the true substrates of CYP168A1 (if not fatty acids) are unlikely to be larger molecules. Alternatively, as in the case of CYP107H1, the fatty acids used in this study may reflect part of the true CYP168A1 substrate(s). Nevertheless, whilst the exact biological function of CYP168A1, like many other bacterial fatty acid hydroxylating CYPs, is yet to be determined, and requires further study. However, it can be hypothesized that CYP168A1 is involved in the degradation of fatty acids. Release of a high concentration of fatty acids (through high levels of phosphatidylcholine degradation) can be toxic to cells as they can cause the inhibition of enzymes involved in fatty acid degradation/β-oxidation [55,56]. By hydroxylating these fatty acids through an enzyme, such as CYP168A1, the toxic effect can be lessened, and fatty acids can be stored.
It may be possible to use CYP168A1 as a potential target for novel anti-Pseudomonas drugs. Azole antifungals are a class of inhibitor that target CYP51 enzymes, presently optimized to inhibit fungal orthologs, through the direct coordination of the imidazole N-3 or the triazole N-4 nitrogen to the CYP51 heme ferric cation as the sixth axial ligand [19]. This mode of action also means that these azole antifungals are also able to bind to the heme iron of other CYPs, such as CYP52A21 from Candida albicans [41], CYP124 from Mycobacterium tuberculosis [40] and CYP164A2 from M. smegmatis [57].
CYP168A1 bound miconazole with greater affinity than CYP164A2 [57] and CYP124 [40], whilst CYP168A1 bound voriconazole with similar affinity to human CYP51 [58] and with greater affinity than M. smegmatis CYP51 [57]. However, CYP168A1 bound azole antifungals relatively poorly compared to fungal CYP51 enzymes, which typically gave K d values of 0.004 to 0.05 μM [58][59][60]. When added to reconstitution assays, where oleic acid was used as the substrate, these azoles had no effect on the catalytic activity of CYP168A1 despite the azole concentration being eight times the concentration of the enzyme used, suggesting that once the substrate is bound to CYP168A1 it is not readily displaced by the azole antifungals investigated in this study. The azole ligand binding studies were performed using pure CYP168A1 enzyme in the absence of substrate and redox partners and that azole binding properties may differ in the CYP168A1 reconstitution assays. MIC experiments with the P. aeruginosa strains DSMZ 22644 and ATCC 39324, grown on oleic acid, agreed with this observation as tebuconazole, miconazole and voriconazole had no inhibitory effect on growth at concentrations up to 128 μg ml -1 . By contrast, Tooker et al [28] reported that CYP168A1 was inhibited by ketoconazole when arachidonic acid was the substrate, resulting in a~70% reduction in enzyme activity with a CYP168A1:ketoconaozle ratio of 5μM:10μM, indicating screening a wider range of azole compounds may identify further CYP168A1 inhibitors. Currently available azole antifungal drugs would need to be redesigned and optimized to target CYP168A1 for such drugs to be considered effective inhibitors of P. aeruginosa.
This study has shown that the previously uncharacterized CYP168A1 from P. aeruginosa is involved in the sub-terminal hydroxylation of biologically relevant fatty acids. It is able to hydroxylate saturated fatty acids at both the ω-1-and ω-2-positions, but it is only able to hydroxylate unsaturated fatty acids at the ω-1-carbon.