Human Molecular Genetics, 2001, Vol. 10, No. 18 1971-1982
© 2001 Oxford University Press
Structure–function analysis of phytanoyl-CoA 2-hydroxylase mutations causing Refsum’s disease
The Oxford Centre for Molecular Science and The Dyson Perrins Laboratory, South Parks Road, Oxford OX1 3QY, UK and 1Department of Chemical Pathology, King’s, Guy’s and St Thomas’ Medical School, St Thomas’ Hospital Campus, Lambeth Palace Road, London SE1 7EH, UK
Received May 10, 2001; Revised and Accepted June 25, 2001;
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTS AND DISCUSSIONCONCLUSIONSMATERIALS AND METHODSREFERENCES |
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Refsum’s disease is a neurological syndrome characterized by adult-onset retinitis pigmentosa, anosmia, sensory neuropathy and phytanic acidaemia. Many cases are caused by mutations in peroxisomal oxygenase phytanoyl-CoA 2-hydroxylase (PAHX) which catalyses the initial
-oxidation step in the degradation of phytanic acid. Both pro and mature forms of recombinant PAHX were produced in Escherichia coli, highly purified, and shown to have a requirement for iron(II) as a co-factor and 2-oxoglutarate as a co-substrate. Sequence analysis in the light of crystallographic data for other members of the 2-oxoglutarate-dependent oxygenase super-family led to secondary structural predictions for PAHX, which were tested by site-directed mutagenesis. The H175A and D177A mutants did not catalyse hydroxylation of phytanoyl-CoA, consistent with their assigned role as iron(II) binding ligands. The clinically observed P29S, Q176K, G204S, N269H, R275Q and R275W mutants were assayed for both 2-oxoglutarate and phytanoyl-CoA oxidation. The P29S mutant was fully active, implying that the mutation resulted in defective targeting of the protein to peroxisomes. Mutation of Arg-275 resulted in impaired 2-oxoglutarate binding. The Q176K, G204S and N269H mutations caused partial uncoupling of 2-oxoglutarate conversion from phytanoyl-CoA oxidation. The results demonstrate that the diagnosis of Refsum’s disease should not solely rely upon PAHX assays for 2-oxoglutarate or phytanoyl-CoA oxidation. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTS AND DISCUSSIONCONCLUSIONSMATERIALS AND METHODSREFERENCES |
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In humans the plasma level of the dietary-derived isoprenoid fatty acid, phytanic acid, is normally low (<30 µM) (1). However, high plasma levels are found in patients with adult Refsum’s disease (ARD) (OMIM 266500) and to a lesser extent in patients with other disorders of peroxisomal function (2). Patients with ARD present with a constellation of symptoms including retinitis pigmentosa, anosmia, deafness, peripheral polyneuropathy, cerebellar ataxia and ichthyoisis, and have increased cerebrospinal fluid (CSF) protein and a pathognomic severe plasma phytanic acidaemia (1–3). Around 45% of reported cases of ARD in the UK are associated with defects in the function of phytanoyl-CoA 2-hydroxylase (PAHX), while the remainder are associated with a second, as yet uncharacterized, locus (4,5). PAHX is an iron(II)- and 2-oxoglutarate-dependent oxygenase (6) that catalyses a key step in the metabolic degradation of phytanic acid. The presence of a 3-methyl group in phytanic acid prevents its degradation by the standard ß-oxidation pathway for saturated fatty acids. Instead, a preliminary four-step
-oxidation pathway occurring in peroxisomes excises a single carbon atom (1,2,7). The phytanic acid is activated to give its coenzyme A ester (8), which is then hydroxylated in the reaction mediated by PAHX (6,9–11). Lyase-catalysed fission of the C1–C2 bond of 2-hydroxyphytanoyl-CoA then occurs to give pristanal (12,13) and formyl-CoA (14). Pristinal is then oxidized in an NAD+-dependent reaction to pristanic acid (15). After re-esterification with coenzyme A and racemization (16–18) further metabolism occurs through the ‘normal’ ß-oxidation pathway initially within the peroxisome and later within mitochondria (19).
Since the identification of PAHX as the enzyme responsible for -oxidation of phytanic acid and its role in ARD, several deletion, truncation and frame-shift mutations giving rise to shortened PAHX enzymes have been identified. PAHX is initially produced in a pro-protein form with an N-terminal peroxisomal targeting sequence-2 (PTS-2) tag. Mature PAHX, found within peroxisomes, is produced by cleavage of the PTS-2 tag between residues Thr-30 and Ser-31 (9,11). Jansen et al. (20) have recently reported a mutation, P29S, located in the N-terminal peroxisomal targeting sequence. This mutant of PAHX has been produced as a maltose-binding protein (MBP)-fusion protein, and analysed for activity in vitro. We also report the purification and characterization of wild-type pro and mature PAHX, and the mature H175A, Q176K, D177A, G204S, N269H, R275W and R275Q functional mutants.
PAHX shows possible mechanistic similarities with related iron(II) and 2-oxoglutarate-dependent oxygenases. These are a heterogenous group of enzymes showing little primary sequence homology but considerable structural similarity when analysed by X-ray crystallography. The PAHX mutations are discussed in the light of sequence and structural comparisons of PAHX with related iron(II) and 2-oxoglutarate-dependent oxygenases for which structures have been reported.
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RESULTS AND DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTS AND DISCUSSIONCONCLUSIONSMATERIALS AND METHODSREFERENCES |
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Cloning, expression and purification of PAHX
Pro and mature forms of human PAHX were cloned into the MBP-fusion vector pMALc and the pET24a vector for expression in Escherichia coli. Both pro and mature forms of the enzyme were produced at
15% of the total soluble protein by sodium dodecylsulphate–polyacylamide gel electrophoresis (SDS–PAGE) analysis as MBP-fusion proteins using the pMALc vector (11). However, wild-type pro-PAHX protein was produced predominately as insoluble inclusion bodies using the pET24a vector. Thus, the pro-protein forms of wild-type and the P29S mutant were produced and characterized as MBP-fusion proteins. Mature wild-type and mutant PAHX enzymes were produced and characterized as soluble protein using the pET24a vector. Mature wild-type PAHX has a relatively high pI compared with other E.coli proteins [8.6 observed, compared with 8.4 (9) calculated], and a convenient purification using a carboxymethyl (CM)–Sepharose cation-exchange column was used. Small-scale purification yielded protein (<50 mg) of >95% purity (by SDS–PAGE analysis) in a single step. When larger quantities of enzyme were required, the pool from the CM–Sepharose column was further purified using gel filtration chromatography (Fig. 1). Di(thio-2-nitro-5-benzoate) (DTNB) titration of the mature wild-type enzyme showed that six of the seven cysteine residues were relatively exposed to solvent. The MBP-fusions of the pro-protein could also be conveniently purified by a protocol employing affinity and cation-exchange chromatography (Fig. 1). All mutant proteins were analysed by circular dichroism, which suggested that no gross distortion to the protein secondary structure had occurred compared with the wild-type enzyme.
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Assay conditions for pro and mature PAHX
Preliminary investigation of in vitro assay conditions showed that mature PAHX had an optimum pH for catalysis of
7.5, with maximum activity observed in 50 mM Tris–HCl [other buffers tested included 50 mM 3-N-(morpholino)propanesulphonic acid (MOPS)–NaOH pH 7.5 and HEPES–NaOH pH 7.5] (Table 1). The effects on PAHX of various co-factors, known to have a pronounced effect on the activity of other non-haem iron(II) and 2-oxoglutarate-dependent oxygenases (21), were investigated. Replacement of dithiothreitol (DTT), which had been previously used in PAHX assays (10), with tris(carboxyethyl)phosphine (TCEP) resulted in a >2-fold increase in activity, as has been observed for other 2-oxoglutarate-dependent oxygenases in vitro (21). Increasing the ascorbate concentration from 1 to 10 mM also enhanced activity of the recombinant enzyme, as had been observed for the native human liver enzyme (22). It has been observed previously that addition of ATP or GTP to the PAHX assay enhances PAHX activity (22,23). Consistent with the previous observations for PAHX, a 2-fold enhancement of activity of highly purified (>95% by SDS–PAGE analysis) recombinant mature PAHX was observed upon addition of 4 mM ATP to the standard co-factor mixture. The basis of this effect is unclear, although the observation that both GTP and ATP have similar effects suggests a non-specific process may be more likely than a specific allosteric activation. Mature PAHX apparently aggregates at concentrations of >5 mg/ml (as shown by gel filtration and dynamic laser light scattering analyses). Thus, it is important to choose appropriate PAHX concentrations (1 mg/ml in the assay) as aggregation resulted in lower activities.
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Iron(II) could not be effectively replaced by any of the alternative metal ions tested [Cd(II), Co(II), Cu(II), Mn(II), Ni(II), Pb(II), Zn(II)] at 2 mM final concentration, consistent with observations for the human liver enzyme (22). Significant activity (
40%) with iron(III) was observed, presumably due to reduction to iron(II) by ascorbate and TCEP under the assay conditions. Several metal ions at 2 mM (almost) completely inhibited PAHX activity in the presence of iron(II) at 2 mM. These were Cd(II), Cu(II), Pb(II), Zn(II), whereas partial inhibition (50%) was observed with Co(II), Fe(III), Mn(II) and Ni(II) at the same concentrations. This behaviour is apparently typical of 2-oxoglutarate-dependent oxygenases such as deacetoxycephalosporin C synthase (24). In the absence of ATP or GTP, the presence of Mg(II) also caused slight inhibition of activity (64 compared with 77 nmol/min/mg) (Table 1).
Several analogues of 2-oxoglutarate are known to inhibit iron(II) and 2-oxoglutarate-dependent oxygenases, e.g. prolyl 4-hydroxylase (25,26). The following analogues were tested for their ability to inhibit PAHX activity at 2–10 mM: N-oxaloglycine, N-oxalyl-2S-alanine, N-oxalyl-2R-alanine, pyridine-2,3-dicarboxylic acid, pyridine-2,6-dicarboxylic acid, N-(2-mercaptopropanoyl)-glycine and N-(3-mercaptopropanoyl)-glycine. N-(2-mercaptopropanoyl)-glycine and N-(3-mercaptopropanoyl)-glycine did not inhibit activity at concentrations up to 10 mM. Pyridine-2,3-dicarboxylic acid and pyridine-2,6-dicarboxylic acid were good inhibitors, and totally abolished PAHX activity at 2 mM. N-oxaloglycine was also a good inhibitor of PAHX, abolishing activity at 2 mM. However, the N-oxalo-2S-alanine, N-oxalo-2R-alanine analogues were less potent inhibitors than N-oxaloglycine, reducing PAHX activity by 50% at 10 mM. The weaker binding of these compounds is presumably due to the steric constraints imposed by accommodation of the alanine side-chain methyl group.
Stereochemical aspects of the PAHX reaction
Phytanic acid and phytanoyl-CoA exist in nature as a mixture of (3R) and (3S) epimers (7). Hydroxylation of the (3R,S)-phytanoyl-CoA by PAHX could give rise to four possible product stereoisomers. An early study by Tsai et al. (27) with rat liver extracts suggested that only one of the two possible diastereomeric pairs was formed from each of the C-3 epimers of phytanoyl-CoA in the enzymatic reaction. Subsequently, Croes et al. (28) demonstrated that both (3R)- and (3S)-3-methylhexadecanoic acid (an analogue of phytanic acid) were hydroxylated in vitro by rat liver extracts. These results indicated that the stereochemistry at C-3 of the substrate (i.e. the carbon bearing the methyl group) determined the product stereochemistry at C-2. It was found that the threo-isomer was exclusively formed from both epimers of the substrate, i.e. the absolute stereochemistry of the products for the (2S) and (2R) substrates were (2S,3R) and (2R,3S), respectively (28).
Extended incubation of an equimolar mixture of the (3R)- and (3S)-epimers of the natural substrate phytanoyl-CoA with highly purified recombinant mature PAHX resulted in >95% conversion to hydroxylated products as judged by high pressure liquid chromatography (HPLC) analysis demonstrating that both epimers are substrates for a single PAHX enzyme (29). Analysis of the time-course for this reaction revealed an apparent burst phase of 1 min under the optimized assay conditions, which may indicate that one epimer is preferentially oxidized, although other explanations for this observation are possible. The absolute stereochemistry of the hydroxylated product is unknown, but is anticipated to be the same as in the study by Croes et al. (28)
Kinetic analysis of wild-type mature PAHX
Initial kinetic analyses of PAHX in the absence of ATP gave an apparent Km for phytanoyl-CoA of 30 µM. However, marked deviation from Michaelis–Menten behaviour was observed upon more detailed analysis. Kinetic analysis of non-haem iron(II)-dependent oxygenases can be complex due to the presence of competing reactions including auto-catalytic inactivation (24,30). Current kinetic analyses of PAHX are further complicated by the use of a discontinuous assay (21) and by the use of a substrate, phytanoyl-CoA, which is a mixture of (3R)- and (3S)-epimers, which may not be oxidized with equal efficiency. Moreover, phytanoyl-CoA has a tendency to form micelles, and requires solubilization with ß-cyclodextrin (10) for incubation with PAHX. Thus, as for the reaction catalysed by the rat liver -methylacyl-CoA racemase (17) (where Nonidet P-40 was used to solubilize the pristanoyl-CoA substrate), it is difficult to determine accurately the effective substrate concentration. Phytanoyl-CoA also has the potential to form iron(II) complexes that may effect kinetic behaviour.
In contrast to phytanoyl-CoA, 2-oxoglutarate displayed the expected Michaelis–Menten behaviour, suggesting that the apparent non-Michaelis–Menten kinetics observed with phytanoyl-CoA are related to the properties of the substrate rather than of the enzyme. The following apparent kinetic parameters (±SEM) for 2-oxoglutarate were obtained: Km = 51.4 ± 15.6 µM; Vmax = 246 ± 17 nmol/min/mg; Kcat = 0.145 ± 0.010 S–1. The Km value compared well with the reported value (49 µM) for enzyme derived from human liver (22)
Structure–function motifs of the iron(II) and 2-oxoglutarate-dependent oxygenases
PAHX belongs to the ubiquitous super-family of 2-oxoglutarate-dependent and related oxygenases (6,31). All characterized members of this super-family have a requirement for iron(II) as a co-factor, dioxygen as a co-substrate, and all but two of the enzymes for 2-oxoglutarate as a co-substrate. Prior to crystallographic studies certain motifs including the HXD..H and RXS motifs, responsible for binding iron and 2-oxoglutarate, respectively, had been identified in many members of the family (6). However, since these motifs were not fully conserved and prior sequence analyses often failed to identify any sequence similarity between ‘sub-families’ it was unclear whether the super-family had evolved to a closely related mechanistic platform via divergent or convergent processes. Recent crystal structures have revealed that, at least for known members of the family, the former is probably the case. Thus, the crystal structures for deacetoxycephalosporin C synthase (DAOCS) (24,32), clavaminic acid synthase (CAS) (33) and proline 3-hydroxylase (P-3-H) (L.Hsueh et al., submitted) have revealed that, despite little initial apparent primary sequence similarity, they all contain an eight-stranded ß-barrel core (34). The CAS structure revealed variation in these motifs in that the iron is ligated by a HXE..H motif and the 5-carboxylate of the 2-oxoglutarate by arginine and threonine residues rather than the more common RXS motif (33,35). Furthermore, these structural studies revealed that the position of the ß-strand core within the overall primary sequence can change, and that there can be significant inserts within the loops linking the strands of the core.
This core forms a ‘platform’ for much of the catalytic machinery of the enzyme including the iron(II) and 2-oxoglutarate binding motifs (Fig. 2A and B), which appears to be conserved in PAHX. This analysis also revealed that secondary structure elements are apparently widely conserved within the family of iron(II) and 2-oxoglutarate-dependent oxygenases, even when the levels of primary sequence homology are low (Fig. 2C) and is limited to short highly conserved motifs, e.g. the HXD iron(II) binding motif.
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In the light of the structural studies, sequence analyses of 2-oxoglutarate-dependent oxygenases can be used to identify sub-families based on the presence of specific structural elements and sequences. Primary sequence analyses revealed at least 22 closely related sequences from a variety of organisms including the known mammalian PAHX enzymes (Fig. 3). Except for the putative hypophosphate dioxygenase (36) and the cis-propenylphosphonic acid epoxidase (37), no function for these proteins has been proposed, but it seems possible that many are involved in the degradation of fatty acid derivatives, including representatives from Mycobacteria. Since phytol displays antibacterial activity against Mycobacteria (38), iron(II) and 2-oxoglutarate-dependent oxygenases may be part of a detoxification mechanism for Mycobacteria and are potential targets for medicinal chemistry.
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Secondary structure predictions and the sequence order of proposed iron and 2-oxoglutarate binding motifs strongly imply that PAHX contains the conserved ß-strand core (Figs 2C and 3). All but one of the characterized, clinically observed PAHX point mutations (P29S) are contained within the proposed ß-strand core. We tested the structural predictions by investigating the in vitro effects of mutations (H175A, Q176K and D177A) to the predicted iron-binding HXD motif (residues 175–177) and a residue, Arg-275, thought to be important for binding the 2-oxoglutarate co-substrate. Following verification of the ‘gross’ structural model, we then investigated the effects of selected clinical mutations. Truncation and deletion mutants will clearly lead to major structural changes to PAHX, but the likely outcome of other mutations was less clear (e.g. P29S, G204S and N269H). The results imply that there are different categories of PAHX mutant, and thus have consequences for diagnosis of Refsum’s disease by enzyme assays and possibly for its treatment.
The P29S mutation
Proline-29 of the pro-form of PAHX is located within its N-terminal peroxisomal targeting sequence-2 (PTS-2) motif (20). Primary sequence comparisons with the targeting sequences of other peroxisomal and mitochondrial enzymes show that this proline is highly conserved, and is located close to the signal peptide cleavage point (between Thr-30 and Ser-31). Purification and characterization of the P29S pro-protein as its MBP-fusion showed that it had full in vitro activity compared with wild-type mature PAHX as measured by both 2-oxoglutarate and phytanoyl-CoA conversion (Table 2).
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The in vivo effect of the P29S mutant is unclear. It may affect transport into the peroxisome, cleavage of the targeting sequence within the peroxisome and/or in vivo solubility of PAHX. Cleavage of the PTS-2 sequence is not required for transport into the peroxisome (39,40). Proline-29 is relatively remote from the PTS-2 signal (residues 9–16), but is close to the cleavage site of the PTS-2 protease (between residues Thr-30 and Ser-31) (9,11), suggesting that the functional effect of the P29S mutation is to hinder cleavage of the 30 residue PTS-2 pro-sequence. Unlike the mature PAHX that is produced in a soluble form, the pro-protein is produced as inclusion bodies in E.coli. Cleavage of the pro-protein may result in a conformational change and possibly activation. Thus, failure to cleave the pro-protein in the peroxisome may result in aggregation and consequent degradation or expulsion of PAHX.
Mutations affecting iron and 2-oxoglutarate binding
Analysis of the primary sequences for PAHX enzymes revealed the presence of a conserved arginine residue, Arg-275, which was predicted to bind the 5-carboxylate of the 2-oxoglutarate co-substrate (Figs 2C and 3). Arg-275 is not part of the RXS motif common in some sub-families of 2-oxoglutarate-dependent oxygenases. However, this motif is absent in many members of the family, including CAS (33) and mammalian prolyl-4-hydroxylase (41). Arg-275 of PAHX of the eighth strand of the conserved ß-sheet core, consistent with its proposed function. Mutations to Arg-275 observed in Refsum’s disease patients include R275W and R275Q (11,20), the former of which has already been reported to be inactive (11).
The efficiency of 2-oxoacid utilization by PAHX depends on both the production of the high-energy iron species that performs substrate oxidation (Scheme 1), and the efficiency of substrate oxidation by this species. Mutations to iron or 2-oxoglutarate binding ligands have previously been shown to uncouple 2-oxoglutarate utilization from substrate oxidation in other oxygenases, e.g. prolyl 4-hydroxylase (41) and DAOCS (42) and this possibility was considered when interpreting the results for the Arg-275 PAHX mutants.
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The R275W, R275Q and R275A mutants were constructed, and shown to have reduced activity (43), consistent with the proposed role of this residue (the R275W and R275Q mutants retained
0.1 and 0.3% of wild-type activity, respectively, with 2-oxoglutarate). However, the effect of these and other mutations to Arg-275 can be partially rescued by the use of alternative 2-oxoacid co-substrates with hydrophobic side-chains, such as 2-oxo-5-thiahexanoic acid (43). This phenomenon is an example of ‘chemical co-substrate rescue’ as reported for DAOCS (42), and strongly implies that Arg-275 is involved in ligating the 5-carboxylate of 2-oxoglutarate. It proved more difficult to rescue the activity of the R275W and R275Q mutants than the R275A mutant. The small methyl side-chain of alanine allows the accommodation of several alternative 2-oxoacids, and it is possible that the introduction of the side-chains of glutamine and tryptophan perturb the eighth conserved strand of the ß-strand core.
An additional mutant with conservation of charge, R275K, was also produced. Unlike wild-type PAHX, the R275K mutant was unable to utilize 2-oxoadipic acid (activity <0.2 nmol/min/mg) but retained 1% of wild-type activity with 2-oxoglutarate (2 compared with 266 nmol/min/mg). This may be a reflection of differences in the catalytic efficiency with different 2-oxoacids. The activity of the R275K mutant could be partially rescued with 2-oxoacids with hydrophobic side-chains, although the level of rescue was low (<1 nmol/min/mg at 2 mM, and 14–23 nmol/min/mg at 60 mM). As with the other Arg-275 mutants the best levels of rescue were obtained with 2-oxo-5-thiahexanoate (43), although several other 2-oxoacids gave significant levels of rescue at 60 mM (data not shown).
The primary sequence similarity analyses (Fig. 3) implied that His-175, Gln-176 and Asp-177 comprise an HXD sequence, which is part of the 2His-1-carboxylate iron(II)-binding motif (44) of PAHX. In order to test the functional effects of these residues, the H175A and D177A mutations were constructed, and the recombinant proteins purified (<95% purity by SDS–PAGE analyses) as mature proteins. The H175A and D177A mutations have not been isolated from patients with Refsum’s disease, although the Q176K and D177G mutations have been reported to be present (20). Assays for both 2-oxoglutarate and phytanoyl-CoA conversion showed that, as expected, the activities of the H175A and D177A mutants were severely impaired (Table 2), with no hydroxylation of phytanoyl-CoA detected. A low level (2–6% of wild-type activity) of 2-oxoglutarate conversion was observed, suggesting that substrate oxidation was significantly uncoupled from 2-oxoglutarate conversion by these mutations. These results confirm the importance of His-175 and Asp-177 for iron(II) binding, and suggest that the D177G mutant of PAHX is inactive due to defective iron binding and, possibly, uncoupling of 2-oxoglutarate conversion from substrate oxidation. Mutations of analogous HXD motif residues in DAOCS (His-143 and Asp-185) (45,46), CAS (His-144 and Glu-146) (47,48) and flavanone 3ß-hydroxylase (His-220 and Asp-222) (49) to other residues also resulted in inactivation.
Following ‘verification’ of His-175 and Asp-177 as iron(II) binding ligands, the clinically observed Q176K mutant was made. The molecular basis for putative inactivation by this mutation is less clear, since the crystal structures of DAOCS (24,32) and CAS (33) imply that it is not involved in iron(II) binding. Activity assays on the Q176K mutant revealed that 2-oxoglutarate conversion was reduced to 57% and substrate oxidation to 16% of the levels of wild-type enzyme, i.e. significant uncoupling of substrate oxidation from 2-oxoglutarate utilization is caused by this mutation in vitro. The coupling ratio between 2-oxoglutarate and phytanoyl-CoA conversion is 3.5:1, compared with 1:1 for the wild-type enzyme. Previous studies on DAOCS have demonstrated that distortion of iron-binding ligands by changing the conformation of a neighbouring residue can result in uncoupling of 2-oxoglutarate conversion from prime substrate oxidation (50).
The identity of the second histidine residue in the 2His-1-carboxylate motif is uncertain, but possible candidates are His-213, His-220, His-259 and His-264. Sequence similarity–structure correlation studies suggest that the most likely candidate is His-264 [probably located on the seventh strand of the conserved core as for DAOCS (32) and CAS (33)] which is in close proximity to Arg-275 (Fig. 3). In some 2-oxoglutarate-dependent oxygenases, e.g. prolyl 4-hydroxylase, a ‘third’ histidine is located in the active site and is apparently involved in 2-oxoglutarate binding (41). His-213 of PAHX, which is conserved throughout the PAHX sub-family, is a candidate for this role. His-220, is mutated to a tyrosine in some patients with Refsum’s disease (20). However, this residue is not highly conserved within related sequences, and thus it is unlikely to be an iron(II) binding histidine.
Further mutations uncoupling 2-oxoglutarate oxidation from substrate conversion
In addition to the R275W mutation (9,20), the initial cloning and expression of PAHX from patients with Refsum’s disease revealed two point mutations, G204S and N269H (9). In order to examine the functional effects of these mutations, the required recombinant mutant enzymes were purified in the mature form.
Activity analyses in vitro demonstrated that both the G204S and N269H mutants were able to convert 2-oxoglutarate to succinate and CO2 at 40% of the level of the wild-type enzyme (Table 2). These results were somewhat surprising as the native G204S mutant has been reported previously to be inactive (20). However, no hydroxylation of phytanoyl-CoA with either mutant was observed within the limits of detection (<0.5% of the conversion observed for the wild-type enzyme), demonstrating that in these mutants substrate oxidation has been completely uncoupled from that of 2-oxoglutarate.
The sequence–structure analyses for PAHX indicated that both Gly-204 and Asn-269 are highly conserved (Fig. 3). Secondary structure predictions indicate that Gly-204 is located prior to or at the start of ß4 of the ß-strand core, and is probably involved in a ß-turn. Thus, its mutation to serine has disturbed the fourth strand of the conserved core and neighbouring secondary structures. Asn-269 is located between ß7 and ß8 of the ß-strand core, probably immediately after a turn (residues 265–267).
Since the presence of substrate significantly stimulates conversion of 2-oxoglutarate, the presence of the G204S and N269H mutations does not prevent binding of phytanoyl-CoA. It seems that these residues play an important role in co-ordinating substrate and co-substrate reactions (Scheme 1), possibly by orienting the substrate correctly with respect to the reactive ferryl intermediate during catalysis (24). Together with the results for the Q176K mutant, the results for the G204S and N269H mutants reveal that assays for PAHX activity should not be solely based on 2-oxoglutarate conversion.
Mutations affecting gross structure
Further known mutations to PAHX can be classified into two other different types: those that cause gross structural defects, and those that effect catalytic function without affecting the gross structure (Table 2). Examples of mutations probably causing gross structural changes include the frame-shift, truncation and exon-deletion mutants (9,11,20). The frame-shift mutation at Leu-55 (20) will result in truncation at residue 66, which will delete the entire conserved ß-strand core of PAHX. The insertion mutation between Ala-192 and Trp-193 will disturb the third strand of the conserved core. The exon-deletion mutations, removing exon 3 (Tyr-46 to Arg-82), will result in the deletion of a large number of amino acids before the conserved core.
Several clinically observed point mutations may also give rise to disturbances in conformation of the core (based on primary sequence homology and secondary structure analyses) and include P173S (which is located at the beginning of the second strand), R245Q (located in the fifth strand) and F257S (located in the sixth strand). In addition, the W193R, E197Q and I199F mutations occur in a hydrophobic region partially occupied by the third strand, with residues 197 and 198 being the only polar/charged residues. The functional effects of these mutations are unclear.
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONRESULTS AND DISCUSSIONCONCLUSIONSMATERIALS AND METHODSREFERENCES |
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Primary sequence alignment based on identity (of amino acids or nucleotides) or similar physiological properties of amino acids often fail to reveal structural or functional relationships between enzymes. A key element in our PAHX work has been the interpretation of primary sequence alignments in the light of known structures for related enzymes and secondary structure predictions. This approach has led to successful prediction testing of the structure–function effects of clinical mutations to PAHX, and should be applicable to the study of other enzymes in the 2-oxoglutarate-dependent oxygenase super-family. Indeed, a recent paper has proposed roles for three putative 2-oxoglutarate-dependent oxygenases using primary sequence homologies in conjunction with known structural motifs (51).
From a diagnostic perspective the results suggest that assays for Refsum’s disease should not be based on PAHX activity alone. The work also suggests that at least four different types of mutation cause loss of PAHX activity in vivo. Firstly, mutations to the peroxisomal targeting sequence do not affect catalytic function but probably affect targeting or degradation of the enzyme. A second group of mutations, including truncation and missense mutations, result in total loss of PAHX activity, i.e. no catalytic conversion of 2-oxoglutarate or phytanoyl-CoA. A third group of mutations results in uncoupling of substrate oxidation from that of 2-oxoglutarate. In the case of two of the mutants (G204S and N269H) falling into this category, this is not due to ‘simple’ hindrance of substrate binding, but is due to incorrect co-ordination during the mechanistic process. A fourth group of mutations causes (partial) inactivation by hindering binding/utilization of the 2-oxoglutarate co-substrate and/or iron(II) co-factor. From a therapeutic and biochemical view-point the latter two groups are particularly interesting since it may be possible to restore their activity with modified co-substrates. In the case of one set of mutations, i.e. to Arg-275, we have already shown that activity can be restored by use of alternative and bio-available co-substrates (43). Diet therapy is already used to treat ARD by limiting phytanic acid intake. It is possible that this therapy may be enhanced in the future by the addition of appropriate precursors, which will ‘restore’ the activity of PAHX by ‘chemical co-substrate rescue’, thus allowing more successful treatment of some cases of this chronic disabling disorder.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTS AND DISCUSSIONCONCLUSIONSMATERIALS AND METHODSREFERENCES |
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All chemicals were supplied by Sigma-Aldrich Chemical (Poole, Dorset, UK), unless otherwise stated, and were of analytical grade or higher. Chromatography systems and columns were obtained from Amersham Pharmacia Biotech (Little Chalfont, Bucks, UK) or Bio-Rad (Hemel Hempstead, Herts, UK). Protein purification was performed on an fast protein liquid chromatography Bio-pilot system. CM–Sepharose was packed into a XK50/20 column (5 x 10 cm, 200 ml) and Superdex-75 was packed into a G32 x 1000 column (Amicon, 3.2 x 89 cm, 716 ml). HPLC columns were obtained from Phenomenex (Macclesfield, Cheshire, UK). (3R,S, 7R, 11R)-Phytanoyl-CoA was synthesized from phytanic acid by known methods (29). N-oxaylglycine and N-oxalyl-alanines were synthesized by Dr E.J.Gibson (Dyson Perrins Laboratory, Oxford, UK) using standard methods (26). Oligonucleotides were synthesized by V.Cooper (Dyson Perrins Laboratory) using Applied Biosystem DNA synthesizers (Models 380B and 394). Other Molecular biology reagents were obtained from Stratagene (Europe) (Amsterdam Zuidoost, The Netherlands) or New England Biolabs (Hitchen, Herts, UK).
Cloning and expression of PAHX
Human cDNA liver library in 
Uni-ZAP XR vector was used to infect XL 1-Blue MRF’ cells. After complete cell lysis, phage DNA was prepared by the standard protocol (52). The complete PAHX gene was amplified from the cDNA library using primers 5'P(E)a forward (5'-CCGGAATTCGTGGGGGTTCCCCGCGCCGCAGCCAT GGAG-3') and 3'P(SNH) reverse (5'-AAGCTTGCTAGCGTCGACTCAAAGATTGGTTCT TTC-3'), cloned into the pGEM-T vector and identified by DNA sequencing. Mature PAHX was amplified with primers 5'PM(KEN)a forward (5'-GGTACCGAATTCCATATGTCAG GGACTATTTCCTCTGCC-3') and 3'P(SNH), cloned into the pGEM-T vector and sequenced.
Pro-PAHX was amplified from the pGEM-T construct using forward primer 5'P(KEN)a, 5'-GGTACCGAATTCCATATGGAGCAGCTTCGC-3' and reverse primer 3'P(SNH), 5'-AAGCTTGCTAGCGTCGACTCAAAGATTGGTTCTTTC-3'. The required fragment was cloned into pGEM-T vector and sequenced. Pro- and mature PAHX were sub-cloned into the pMALc vector using 5' EcoRI and 3' HindIII restriction sites for expression, whilst for the pET24a vector they were cloned using 5' NdeI and 3' HindIII restriction site. Clones were checked for expression by induction with 0.5 mM isopropyl-ß-D-thiogalactoside (IPTG) for 4 h at 27°C. In the pMALc vector, both fragments, with N-terminal MBP-fusion, expressed as 15–20% of total soluble protein. In pET24a the native pro-PAHX expressed predominantly as insoluble inclusion bodies, whilst mature PAHX was expressed as 15% soluble protein.
Site-directed mutagenesis
Point mutations were created in the pGEM-T/ mature PAHX vector by PCR using an oligonucleotide-mediated double primer method (QuikChange), according to the manufacturers’ instructions (Table 3). The presence of the mutation was confirmed by automated DNA sequencing.
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Fermentation of recombinant E.coli cells
Escherichia coli BL21 (DE3)/pET24a expressing mature PAHX was inoculated from a glycerol freeze into 4 x 100 ml of 2TY medium supplemented with 25 µg/ml of kanamycin and grown overnight at 37°C. This was used to inoculate 30 l of the same medium supplemented with 5 ml of polypropylene glycol 2025 in a New Brunswick Scientific MPP40 fermenter. The culture was grown at 27°C and 300 r.p.m. with 30 l/min air for 4 h. Cells were induced at OD600 = 0.8 with 0.5 mM IPTG and fermented for a further 4 h. Cells were concentrated to
3 l using a tangential flow concentrator, centrifuged (17 500 g, 15 min) and stored at –80 °C.
Enzyme purification
All manipulations were performed on ice or at 4°C. Fractions were analysed by SDS–PAGE using 12.5% (w/v) resolving and 3% (w/v) stacking gels. Protein concentrations were determined by the method of Bradford (53). Iso-electric points of pro- and mature PAHX were determined with a Bio-Rad analytical flat bed IEF using pre-cast PhastGelTM IEF 3–9 gels (Amersham Pharmacia Biotech). Pro- and mature PAHX from the pMALc vector (with N-terminal MBP-fusion) were purified by affinity chromatography using MBP binding protein columns (New England Biolabs), according to the manufacturer’s instructions, followed by cation-exchange chromatography (vide infra). The molecular weights of all enzymes were determined by ESI MS, and were within experimental error of the calculated molecular weight.
Cells expressing wild-type or mutant mature PAHX [E.coli BL21 (DE3)/ pET 24a, 50 g] were thawed and resuspended in 500 ml of 20 mM MOPS-NaOH, 10% (v/v) glycerol pH 7.2, 1 mM benzamidine-HCl, 1 mM phenylmethylsulphonyl fluoride (PMSF), 0.5 mM 1,10 o-phenanthroline and 2 mM DTT. PAHX aggregated when cells were suspended at >10% (w/v) and did not bind to the CM–Sepharose column. The solution was stirred for 15 min and sonicated for 5 x 30 s at power 5 with 60 s cooling (W-380 sonicator, Ultrasonic). DNA was precipitated with 1% (w/v) streptomycin sulphate and 0.1% (w/v) polyethyleneimine. After 10 min stirring the extract was centrifuged (JA-14 rotor, 22 000 g, 10 min), loaded onto a CM–Sepharose column equilibrated in 20 mM MOPS-NaOH, 10% (v/v) glycerol pH 7.2, 0.2 mM benzamidine-HCl, 0.2 mM PMSF, 0.1 mM 1,10 o-phenalthraline and 2 mM DTT (low salt buffer) and washed with 1000 ml of same buffer at 10 ml/min. Protein was eluted with a 0–700 mM NaCl gradient in the same buffer over 900 ml, followed by 100 ml of 1 M NaCl in buffer. Fractions (25 ml) 12–21 (330–580 mM NaCl) were pooled and concentrated in 300 and 50 ml Amicon-stirred cells with a YM-30 membrane and exchanged into 10 mM Tris–HCl, 10% (v/v) glycerol pH 7.5 with an Econo-Pak column. Protein was concentrated to 30 mg/ml and stored at –80°C. Wild-type enzyme was further purified by concentrating to 30 ml and loading 4 x 7.5 ml onto Superdex-75 column (716 ml) equilibrated in 100 mM Tris–HCl, 50 mM NaCl, 10% (v/v) glycerol pH 7.5. Protein was eluted at 2 ml/min and fractions (5 ml) collected between 270–470 ml. Fractions 4–10 (290–320 ml) were pooled, concentrated and exchanged as above.
The apparent molecular mass of native mature PAHX was determined by exchanging into gel filtration buffer and loading 5 mg (in 2 ml) onto a Superdex-75 column (716 ml) which had been equilibrated in gel filtration buffer at 2 ml/min. The column was calibrated using the following proteins (5 mg of each) BSA (66 kDa); ovalbumin (45 kDa); carbonic anhydrase (29 kDa); cytochrome C (12.4 kDa).
DTNB titration of PAHX
Pro- and mature PAHX (1 mg) in 200 µl of Tris–HCl pH 7.5 was incubated in ice with 2.5 mM DTT for 135 min. The samples were diluted to 500 µl and exchanged into 50 mM Tris–HCl pH 8.0 with a NAP-5 column (Amersham Pharmacia). The sample was diluted to 8 ml and 0.7 ml was assayed at A412 in the presence and absence of 2% (w/v) SDS with 1 mM DTNB (54).
Dynamic laser light scattering
Dynamic laser light scattering measurements were performed in a DynaPro-801 TC instrument at 20°C using various concentrations of mature PAHX in 10 mM Tris–HCl pH 7.5.
Circular dichroism studies
Enzymes were exchanged into 20 mM NaH2PO4-NaOH buffer (pH 7.0) using Bio-Rad Econo-Pak columns. CD spectroscopic analyses between 190 and 300 nm (50 nm/min) were performed using a Jasco J-720 spectropolarimeter with a 1 mm cell at room temperature. Protein concentration was measured by amino acid analysis and values adjusted in the final equation.
Enzyme assays
Assays were performed (at least in triplicate) in a final volume of 100 µl with final concentrations of: 50 mM Tris–HCl pH 7.5, 1 mM iron(II), 2 mM 2-oxoglutarate, 50 µM synthetic (3R,S, 7R, 11R)-phytanoyl-CoA, 0.44 mM ß-cyclodextrin, 100 µM TCEP, 10 mM ascorbate, 4 mM ATP and 10 µg wild-type or mutant enzyme for 5 min. Analyses of 2-oxoglutarate inhibitors (2–10 mM) were performed using the same assay mixture. ‘Chemical co-substrate rescue’ experiments were performed as above except using 20 µg enzyme (2 mM 2-oxoacid, 60 min incubation) or 10 µg enzyme (60 mM 2-oxoacid, 5 min incubation). The reaction was quenched with 250 mM ethylenediamine-tetraacetic acid [(EDTA) final concentration] [Quenching with 100 mM EDTA and 40% (v/v) acetonitrile resulted in multiple product peaks]. Samples were centrifuged and analysed by HPLC with a Hypersil C18 column (4.6 x 250 mm) at 254 nm (10). 2-Oxoglutarate conversion assays were performed under similar conditions in a total volume of 200 µl using the reported assay (50) and quenched after 30 min.
Kinetic parameters for Phytanoyl-CoA and 2-oxoglutarate were determined using assays at 30°C for 5 min. Mature PAHX (10 µl, 88 µg/ml), 10–200 µM phytanoyl-CoA and 20–2000 µM 2-oxoglutarate were used. Kinetic parameters (± SE) were determined using SigmaPlot, assuming a mass of 35 452 ± 3 Da for wild-type mature PAHX.
Multiple sequence alignments and secondary structure predictions
Sequences were identified with a blastp search using the NCBI website (www.ncbi.nlm.gov) with default parameters. Unfinished sequences were obtained from the Institute for Genome Research Website (http://www.tigr.org). Multiple sequence alignment was performed using ClusterW, and the secondary structure predicted from the consensus sequence using Jpred2 and Prof programmes with minor manual refinements.
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ACKNOWLEDGEMENTS |
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We thank Professor R.J.A.Wanders (AMC, University of Amsterdam) for helpful discussions, Dr R.T.Aplin for ESI MS analyses, and A.C.Willis for protein sequencing and amino acid analyses. The BBSRC, EPSRC, MRC, Wellcome Trust, the E.U. Biotechnology Project and the Felix Foundation (scholarship to M.M.) funded this work. Preliminary sequence data was obtained from the Institute for Genome Research Website (http://www.tigr.org). The NIDR and the Burroughs Wellcome Fund funded Candida albicans sequencing; Beowulf Genomics funded Bordetella pertussis sequencing. MAFF and the Wellcome Trust fund Mycobacterium bovis sequencing. The Wellcome Trust funds Myobacterium tuberculosis (H37rv) sequencing. NIAID funds Mycobacterium avian sequencing.
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FOOTNOTES |
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+ To whom correspondence should be addressed. Tel: +44 1865 275677; Fax: +44 1865 275674; Email: matthew.lloyd@chem.ox.ac.uk
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REFERENCES |
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