Crystallization and preliminary X-ray analysis of the reductase component of p-hydroxyphenylacetate 3-hydroxylase from Acinetobacter baumannii

The reductase component of p-hydroxyphenylacetate 3-hydroxylase from A. baumannii was overexpressed, purified and crystallized. X-ray diffraction data were collected and processed to 2.3 Å resolution.

Abstract

p-Hydroxyphenylacetate 3-hydroxylase (HPAH) from Acinetobacter baumannii catalyzes the hydroxylation of p-hydroxyphenylacetate (HPA) at the ortho position to yield 3,4-dihydroxyphenylacetate (DHPA). HPAH from A. baumannii is a two-component flavoprotein consisting of a smaller reductase (C₁) component and a larger oxygenase (C₂) component. The C₁ component supplies a reduced flavin in its free form to the C₂ counterpart for hydroxylation. In addition, HPA can bind to C₁ and enhance the flavin-reduction rate without becoming hydroxylated. The recombinant C₁ component was purified and crystallized using the microbatch method at 295 K. X-ray diffraction data were collected to 2.3 Å resolution using synchrotron radiation on the BL13B1 beamline at NSRRC, Taiwan. The crystal belonged to the orthorhombic space group P2₁2₁2₁, with unit-cell parameters a = 47.78, b = 59.92, c = 211.85 Å, and contained two molecules of C₁ per asymmetric unit.

1. Introduction  

p-Hydroxyphenylacetate 3-hydroxylase (HPAH; EC 1.14.13.3) is a two-component flavoprotein that catalyzes the hydroxylation of p-­hydroxyphenylacetate (HPA) to form 3,4-dihydroxyphenylacetate (DHPA). HPAH has been identified and studied in the bacteria Pseudomonas putida (Arunachalam et al., 1992 ▶, 1994 ▶), Escherichia coli W (Galán et al., 2000 ▶), Acinetobacter baumannii (Chaiyen et al., 2001 ▶) and Pseudomonas aeruginosa (Chakraborty et al., 2010 ▶), as well as in the archaea Sulfolobus tokodaii strain 7 (Okai et al., 2006 ▶) and Thermus thermophilus HB8 (Kim et al., 2008 ▶). HPAH comprises a smaller reductase and a larger oxygenase on separate polypeptide chains. The reductase functions to generate a reduced flavin, which is able to diffuse to the oxygenase component for the hydroxylation reaction without a requirement for protein–protein interaction (Sucharitakul et al., 2005 ▶, 2006 ▶, 2007 ▶). The dihydroxylated product DHPA is ultimately converted to intermediate metabolites in the TCA cycle (Sparnins et al., 1974 ▶). Therefore, the enzyme is important for the bacterial biodegradation of lignin and phenolic compounds. Additional biological functions of HPAH in A. baumannii (which is an opportunistic pathogen) may also exist and are currently being investigated by our group.

HPAH from A. baumannii has been isolated and extensively investigated (Chaiyen et al., 2001 ▶; Thotsaporn et al., 2004 ▶, 2011 ▶; Sucharitakul et al., 2005 ▶, 2006 ▶, 2007 ▶; Ruangcharn et al., 2011 ▶; Tongsook et al., 2011 ▶). The enzyme possesses interesting properties which differ from those of other previously studied HPAH systems. The reductase component (C₁) of A. baumannii was purified with a tightly bound FMN molecule in each subunit. The enzyme is a homodimer with a subunit mass of about 35 kDa, which is similar to that of the reductase component of P. putida HPAH but larger than that of E. coli W HPAH. Its function is to generate reduced FMN for HPA hydroxylation by the larger oxygenase component (C₂). Studies of its kinetic mechanism have revealed that C₁ is also able to bind HPA without catalyzing its hydroxylation (Sucharitakul et al., 2005 ▶). Such HPA binding enhances the rates of both the reduction of FMN and the release of reduced FMN from C₁ (Sucharitakul et al., 2005 ▶, 2007 ▶). This effect has not been observed for any of the known flavin reductases from various organisms apart from the reductase component of P. putida HPAH (Arunachalam et al., 1992 ▶, 1994 ▶) and component B of nitrilotriacetate (NTA) monooxygenase from Chelatobacter strain ATCC 29600 (Uetz et al., 1992 ▶). Sequence analysis also revealed that the N-terminal half of C₁ is similar to the flavin-binding domains of other reductases (Thotsaporn et al., 2004 ▶). In contrast, its C-terminal half, which is absent in the other reductases, is similar to the corresponding sequences of the reductase components of Chelatobacter NTA monooxygenase and P. putida HPAH (13 and 47% identity over 144 amino acids, respectively). Therefore, these C-terminal sequences may be involved in HPA binding and may be responsible for substrate stimulation in the reaction.

The crystal structures of flavin reductases from various organisms have been determined, including phenol 2-hydroxylase (PheA2) from Bacillus thermoglucosidasius A7 (PDB entry 1rz0; van den Heuvel et al., 2004 ▶), HpaC of HPAH from S. tokodaii strain 7 (PDB entry 2d36; Okai et al., 2006 ▶) and HpaC from T. thermophilus HB8 (PDB entry 2ecr; Kim et al., 2008 ▶). All of these structures share the same protein fold and lack the extra C-terminal domain found in C₁. Recently, the structure of NTA monooxygenase component B from Mycobacterium thermoresistibile (PDB entry 3nfw; Zhang et al., 2011 ▶) has been determined. This enzyme possesses a similar but much smaller C-­terminal extension that is expected to regulate its function, as in C₁. At around 130 residues shorter in its C-terminal sequence in comparison with C₁, the C-terminal structure of this enzyme is likely to differ significantly from those of C₁ and the reductases of P. putida HPAH and Chelatobacter NTA monooxygenase. Aiming to use C₁ from A. baumannii HPAH as a representative structure of this group of reductases, we present its crystallization and preliminary X-ray analysis.

2. Materials and methods  

2.1. Expression and purification  

Recombinant C₁ was overexpressed and purified as previously reported (Chaiyen et al., 2001 ▶; Thotsaporn et al., 2004 ▶) with the following modifications. Briefly, cells were grown in Luria–Bertani medium supplemented with 100 µg ml⁻¹ ampicillin at 310 K until the OD₆₀₀ reached 1.0–1.5. Protein expression was induced with 1 mM isopropyl β-d-1-thiogalactopyranoside at 289 K. The cells were harvested after 8 h of induction or when the OD₆₀₀ reached 4.0 by centrifugation at 5500g for 10 min at 277 K. The cell paste was resuspended in a lysis buffer consisting of 100 mM sodium phosphate pH 8.0, 5 mM ethylenediaminetetraacetic acid (EDTA), 5 mM dithiothreitol (DTT), 60 µM phenylmethylsulfonyl fluoride. The cell suspension was disrupted on ice by ultrasonication and the lysate was centrifuged at 3500g for 1 h at 277 K. All subsequent purification steps were performed at 277 K. The supernatant was precipitated by 40–80% ammonium sulfate fractionation. The yellow pellet was resuspended and concentrated in buffer A (10 mM sodium phosphate buffer pH 7.0, 0.5 mM EDTA, 1 mM DTT). The concentrated enzyme was centrifuged at 3500g for 10 min before applying it onto a DEAE-Sepharose Fast Flow column (GE Healthcare) that had been pre-equilibrated and was subsequently washed with the same buffer. The enzyme was eluted with a gradient of 0–250 mM NaCl in buffer A. The enzyme fractions were pooled based on the OD₄₅₈ and then precipitated by 80% ammonium sulfate saturation. The yellow pellet was dissolved in a minimal volume of 15% saturated ammonium sulfate in buffer A. The dissolved enzyme solution was centrifuged at 3500g for 10 min and loaded onto a Phenyl-Sepharose column (GE Healthcare) that had been pre-equilibrated and was subsequently washed with 15% saturated ammonium sulfate in buffer A. The enzyme was eluted with a gradient of 15–0% ammonium sulfate and 0–40%(v/v) ethylene glycol in buffer A. Fractions containing C₁ were pooled based on the OD₄₅₈/OD₂₈₀ ratio. The purity of the enzyme was judged by 12.5% SDS–PAGE. The C₁ pool was concentrated using Amicon Ultra-4 centrifugal filter units with 10 kDa molecular-weight cutoff (Millipore) and was loaded onto a G-25 column for buffer exchange to 50 mM MOPS pH 7.0, 0.3 mM EDTA, 1 mM DTT. It was then concentrated to 10 mg ml⁻¹ based on its known extinction coefficient at 458 nm of 12 800 M ⁻¹ cm⁻¹ (Sucharitakul et al., 2005 ▶). HPA (10 mM in 50 mM MOPS pH 7.0) was added to an aliquot of the concentrated C₁ to give a final concentration of 1 mM for subsequent crystallization experiments.

2.2. Crystallization  

The purified C₁ was freshly prepared and filtered through 0.22 µm Amicon Ultrafree-MC centrifugal filter devices (Millipore) prior to crystallization setup. Crystallization of C₁ was carried out using the microbatch method (Chayen et al., 1992 ▶; D’Arcy et al., 2004 ▶) in 60-­well Mini Tray plates (Nunc) at 295 K. A droplet was set up under oil (Babi Mild Natural N’Mild baby oil; Chitnumsub et al., 2004 ▶) by mixing equal volumes (1–3 µl) of protein and crystallization solutions. Yellow single crystals were obtained using 20%(m/v) PEG 400, 0.1 M sodium acetate pH 4.6 as a precipitant. After optimization by varying the concentration of PEG 400 and glycerol, crystals were produced in a clear droplet from conditions consisting of 20–25%(m/v) PEG 400, 0.1 M sodium acetate pH 4.6, 10%(v/v) glycerol within 1 d and reached approximate dimensions of 0.15 × 0.15 × 0.3 mm in 3 d.

2.3. X-ray data collection  

The crystals were quickly dipped in Paratone-N as a cryoprotectant before vitrification and data collection. Initial X-ray diffraction experiments were performed at the Center for Excellence in Protein Structure and Function, Faculty of Science, Mahidol University, Thailand. The crystals were mounted on a nylon cryoloop (0.2–0.3 mm) and exposed to X-ray radiation using an in-house rotating-anode X-ray generator (Rigaku RU-H3R; Cu Kα; λ = 1.5418 Å) operating at 50 kV and 100 mA with an R-AXIS IV⁺⁺ detector system (Rigaku/MSC) for preliminary diffraction analysis. X-ray diffraction data were collected using an ADSC Quantum-315 CCD detector under a nitrogen cryostream (100 K) on beamline BL13B1 at the National Synchrotron Radiation Research Center (NSRRC), Taiwan. The crystal-to-detector distance and the offset of the detector position in the vertical direction were optimized to 400 and 25 mm, respectively, in order to collect X-ray reflections at higher resolution since the crystal was able to diffract beyond 2.6 Å resolution. A total of 360 frames with 0.5° oscillation width and 5 s exposure time per frame were collected and processed to 2.3 Å resolution. Data were processed using the CrystalClear program suite (Pflugrath, 1999 ▶) and the HKL-2000 package (Otwinowski & Minor, 1997 ▶) for the in-house and synchrotron data sets, respectively. X-ray data-collection statistics for the synchrotron data set are summarized in Table 1 ▶.

Table 1. Data-collection and processing statistics.

Values in parentheses are for the highest resolution shell.

X-ray source	Beamline BL13B1, NSRRC
Wavelength (Å)	1.0000
Resolution (Å)	50.00–2.30 (2.38–2.30)
Space group	P212121
Unit-cell parameters (Å)	a = 47.78, b = 59.92, c = 211.85
Total reflections	143944
Unique reflections	27013 (2261)
Completeness (%)	95.7 (81.6)
Multiplicity	5.3 (3.1)
〈I/σ(I)〉	40.3 (17.1)
Rmerge†	0.035 (0.064)

^† R _merge = , where I_i(hkl) is the intensity of the ith measurement of an equivalent reflection with indices hkl.

3. Results and discussion  

The recombinant C₁ component of A. baumannii HPAH was overexpressed in E. coli and purified to >95% homogeneity as judged by SDS–PAGE and native PAGE. The method produced approximately 150 mg pure protein from 3.6 l cell culture. The purified C₁ was crystallized using the microbatch method. Single rod-shaped crystals suitable for X-ray data collection were reproducibly obtained (Fig. 1 ▶). Their yellow colour indicated the presence of oxidized flavin molecules in the crystals. The crystals diffracted X-rays to beyond 2.6 Å resolution using our home-source X-ray equipment. Data processing showed that this crystal form belonged to the ortho­rhombic space group P2₁2₁2₁, with a long unit-cell length in the c dimension. Overlapping reflections were observed, which limited the usable data to only low resolution, despite the fact that the crystals diffracted much further. To overcome this problem, the orientation of the crystal was optimized by aligning its c axis parallel to the spindle angle of the goniometer. Practically, this was very difficult as the c axis was along one of the widths of the thin rod-shaped crystals, while the viscous cryoprotectant tended to pull the crystal back along the oval-shaped nylon cryoloop in its typical geometry parallel to the pin. Moreover, prolonged soaking (>10 s) in the cryoprotectant caused a dramatic decrease in diffraction, limiting the time that could be used for crystal orientation. Therefore, an L-shaped nylon cryoloop was made to speed up the mounting process, which allowed successful vitrification of a crystal in the desired orientation with high-quality X-­ray diffraction (Fig. 2 ▶). In addition to the optimization of data collection by crystal orientation, the detector was also offset to maximize the collection of higher angle diffraction. A complete data set was collected to 2.3 Å resolution using synchrotron radiation. The data were processed in the orthorhombic space group P2₁2₁2₁, with unit-cell parameters a = 47.78, b = 59.92, c = 211.85 Å. The calculated molecular volume (Matthews coefficient; Matthews, 1968 ▶) of 2.14 ų Da⁻¹ and solvent content of 42.7% suggested that there were two molecules of C₁ in the asymmetric unit.

Figure 1.

A crystal of the C₁ component of A. baumannii HPAH. This orthorhombic crystal grew to dimensions of 0.15 × 0.15 × 0.5 mm in 3 d.

Figure 2.

X-ray diffraction pattern of the C₁ crystal. The crystal was mounted such that its long axis was nearly parallel to the spindle axis to minimize reflection overlap. This was accomplished by the use of an L-shaped nylon loop (inset). While maintaining the crystal-to-detector distance of 400 mm to prevent spot overlaps, the CCD detector was offset to allow the collection of higher resolution data at the upper side of the detector.

The self-rotation function calculated in AMoRe (Navaza, 1994 ▶; Winn et al., 2011 ▶) showed a significant peak (62.3% of the self-peak) representing a twofold rotation axis located around 13° from the c axis, while no outstanding peak was found in the native Patterson. This suggested a dimeric structure for C₁, which is consistent with the previously reported results of gel-filtration chromatography (Chaiyen et al., 2001 ▶). We attempted to determine the structure of C₁ using the molecular-replacement method. The coordinates of the PheA2 component of phenol hydroxylase from B. thermoglucosidasius A7 (PDB entry 1rz0; 37% sequence identity over 161 residues) was used as a template to construct search models using either all atoms or polyalanines and conserved residues. However, no reasonable solution could be obtained using either monomeric or dimeric structures of PheA2. Since the PheA2 templates would account for only 25–50% of the molecular size of the dimeric C₁ in the asymmetric unit, they may be too small to provide sufficient phases for C₁ structure determination. In addition, monomeric or dimeric structures of NTA monooxygenase component B from M. thermoresistibile (PDB entry 3nfw; 29% sequence identity over 210 residues) were similarly used as search templates with a lack of success. This may suggest that the structure of the short C-terminal extension of 3nfw differs significantly from that of C₁. Hence, other phasing methods such as MIR or MAD are required for phase determination of the C₁ structure.