HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS		QUICK SEARCH:   [advanced] Author: Keyword(s): Year:  Vol:  Page:

J. Biol. Chem., Vol. 282, Issue 41, 30189-30197, October 12, 2007

This Article Abstract Full Text (PDF) Supplemental Data All Versions of this Article: 282/41/30189    most recent M704138200v1 Submit a Letter to Editor Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Wu, H. Articles by Plotnikov, A. N. Search for Related Content PubMed PubMed Citation Articles by Wu, H. Articles by Plotnikov, A. N. Social Bookmarking                      What's this?

Structural Basis of Substrate-binding Specificity of Human Arylamine N-Acetyltransferases^*

Hong Wu, Ludmila Dombrovsky, Wolfram Tempel, Fernando Martin, Peter Loppnau, Geoffrey H. Goodfellow, Denis M. Grant, and Alexander N. Plotnikov^¶1

From the Structural Genomics Consortium, University of Toronto, Toronto, Ontario M5G 1L5, the Department of Pharmacology, University of Toronto, Toronto, Ontario M5S 1A8, and the ^¶Department of Physiology, University of Toronto, Toronto, Ontario M5S 1A8, Canada

Received for publication, May 18, 2007 , and in revised form, July 18, 2007.

	ABSTRACT

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS AND DISCUSSION REFERENCES

 
The human arylamine N-acetyltransferases NAT1 and NAT2 play an important role in the biotransformation of a plethora of aromatic amine and hydrazine drugs. They are also able to participate in the bioactivation of several known carcinogens. Each of these enzymes is genetically variable in human populations, and polymorphisms in NAT genes have been associated with various cancers. Here we have solved the high resolution crystal structures of human NAT1 and NAT2, including NAT1 in complex with the irreversible inhibitor 2-bromoacetanilide, a NAT1 active site mutant, and NAT2 in complex with CoA, and have refined them to 1.7-, 1.8-, and 1.9-Å resolution, respectively. The crystal structures reveal novel structural features unique to human NATs and provide insights into the structural basis of the substrate specificity and genetic polymorphism of these enzymes.

	INTRODUCTION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS AND DISCUSSION REFERENCES

 
Arylamine N-acetyltransferases (NATs, EC 2.3.1.5)² catalyze the acetyl-CoA-dependent N-acetylation of primary aromatic amines and hydrazines, as well as the O-acetylation of their N-hydroxylated metabolites, thereby influencing the biological activity and toxicity of this class of chemicals (1-3). NAT enzyme activity therefore plays an important role in determining the duration of action and pharmacokinetics of aromatic amine-containing drugs used in clinical therapy, as well as in influencing the balance between detoxification and metabolic activation of aromatic amine procarcinogens (4). In the latter regard, N-acetylation reactions are considered to be protective, because the resulting arylacetamide derivatives are chemically stable, whereas O-acetylation of hydroxylamines produces acetoxy esters that can spontaneously decompose to electrophilic, DNA-binding nitrenium ions (5). Thus a better understanding of the structural features of NATs that contribute to their relative abilities to catalyze such reactions for various amine substrates may be of considerable predictive value in optimizing drug development and biomedical toxicology.

Two NATs, NAT1 and NAT2, have been annotated in the human genome. Each of these enzymes is genetically variable in human populations, with variable activity of NAT2 being responsible for the classic acetylation polymorphism that was discovered over half a century ago with the advent of isoniazid therapy for tuberculosis (6-8). In addition to isoniazid, the disposition of several other therapeutically useful drugs is affected by defective NAT2 function, including hydralazine, phenelzine, procainamide, and some of the sulfonamide anti-bacterials. NAT1 is highly homologous to NAT2 yet kinetically distinct, such that certain aromatic amines (such as isoniazid and sulfamethazine) are preferentially acetylated by NAT2, whereas others (such as p-aminosalicylic acid and p-aminobenzoic acid) are selective substrates for NAT1. NAT1 is also genetically variable in human populations. Numerous epidemiological studies have reported associations between both NAT1 and NAT2 variation and the occurrence of cancers related to exposure to aromatic amines (such as 4-aminobiphenyl and 2-naphthylamine) in the environment, although in many instances such findings have been contradictory.

NAT enzyme orthologues are also present in most (but not all) mammalian species, as well as in prokaryotes. Although the crystal structures of some prokaryotic NAT enzymes have been reported, eukaryotic NATs have so far eluded any attempts at the solution of a crystal structure. Here we present high resolution crystal structures of both human NATs, including NAT1 in complex with the irreversible inhibitor 2-bromoacetanilide, a site-directed NAT1 mutant, NAT1_F125S, and a NAT2-CoA complex. By comparing our structures with known prokaryotic structures, we provide evidence for novel structural features of human NATs that are absent in bacterial enzymes, including an insertion that produces a significant difference in the structure of the carboxyl terminus of the eukaryotic enzymes. We also provide the first complete picture of the distinct substrate selectivity of human NAT1 and NAT2, as well as key features of the naturally occurring variants of NAT1 and NAT2 that may explain the structural basis of their defective function.

	EXPERIMENTAL PROCEDURES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS AND DISCUSSION REFERENCES

 
Protein Expression and Purification—DNA fragments encoding human NAT1 and NAT2 were amplified by PCR and subcloned into modified cloning vectors (pET28a-LIC and pET28a-MHL, respectively), downstream of the polyhistidine coding region. Human NAT1 and NAT2 proteins were expressed in Escherichia coli BL21(DE3) codon plus RIL strain (Stratagene). E. coli cells carrying the NAT1_F125S mutant expression plasmid were grown in M9 minimal medium supplemented with 50 mg/liter kanamycin and induced by isopropyl-1-thio-D-galactopyranoside, final concentration of 1 mM, in the presence of 50 mg/liter selenomethionine and incubated overnight at 15 °C. Cells carrying wild-type NAT1 and NAT2 expression plasmids were grown at 37 °C in Terrific Broth, induced by addition of isopropyl-1-thio-D-galactopyranoside, and incubated overnight at 15 °C.

For purification of NAT1_F125S, the cells were harvested by centrifugation and lysed by passing through a microfluidizer (Microfluidics Corp.). The lysate was loaded onto a HiTrap Chelating column (Amersham Biosciences) charged with Ni²⁺. The column was washed with 10 column volumes of 20 mM Tris-HCl buffer, pH 8.0, containing 500 mM NaCl, 50 mM imidazole, and 5% glycerol, and the protein was eluted with elution buffer (20 mM Tris-HCl, pH 8.0, 500 mM NaCl, 250 mM imidazole, 5% glycerol). Eluted protein was treated with iodoacetamide at room temperature for 30 min. The protein was then loaded onto a Superdex 200 column (26 x 600 mm, Amersham Biosciences) and equilibrated with 20 mM Tris-HCl buffer, pH 8.0, and 150 mM NaCl. Thrombin (Sigma) was added to combined fractions containing NAT1 to remove the His tag. The protein was further purified to homogeneity by ion-exchange chromatography. Wild-type NAT1 was purified as described above, except the eluted protein from the nickel column was treated with 2-bromoacetanilide. For purification of NAT2, the eluate from the nickel column was dialyzed against 20 mM Tris-HCl, pH 8.0, 250 mM NaCl, 5% glycerol in the presence of tobacco etch virus protease. The protein was further purified to homogeneity by ion-exchange chromatography.

Crystallization and Data Collection—Purified NAT1_F125S was crystallized using the hanging drop vapor diffusion method at 20 °C by mixing 1 µl of the protein solution with 1 µl of reservoir solution containing 26% polyethylene glycol 4000, 0.2 M sodium acetate, 0.1 M Tris-HCl, pH 7.8. Wild-type NAT1 protein was crystallized by the same method in a solution containing 30% polyethylene glycol 4000, 0.2 M sodium acetate, 0.1 M Tris-HCl, pH 8.5. Purified NAT2 protein was complexed with CoA at a 1:10 molar ratio of protein to CoA and crystallized using the sitting drop, vapor-diffusion method by mixing 1 µl of protein solution with 1 µl of reservoir solution containing 2.5 M ammonium sulfate, 0.1 M Tris-HCl, pH 8.5.

Diffraction data for crystallographic model refinement were collected as shown in Table 1. For an isomorphous crystal of the NAT1_F125S, data sets at wavelengths of 0.9791 and 0.9795 Å were also collected at the National Synchrotron Light Source, beamline X25. All diffraction images were reduced to structure factor amplitudes using the HKL2000 software suite (9).

View larger version (36K): [in this window] [in a new window]   FIGURE 1. Overall structure of human NAT1 and NAT2. Ribbon diagram showing the structure of the human NAT1 (left, with acetanilide covalently bound to catalytic cysteine), NAT2 (middle, in complex with CoA), and S. typhimurium NAT (right). The three structures are presented in the same orientation. All strands and helices are labeled. The amino and carboxyl termini are labeled as NT and CT, respectively. Acetamide covalently attached to Cys-68 in NAT1 and CoA bound to NAT2 are shown in sticks, with carbon atoms as gray. The insertions and carboxyl-terminal tails of human NATs, which are missing in NAT proteins from other species, are colored in magenta.In the S. typhimurium NAT structure, all the sub-domains are colored as described previously (20): domain I (helical bundle, residues 1-86) in blue, domain II (-barrel, residues 87-175) in yellow, domain III (/, residues 204-273) in green, and interdomain region (residues 176-203) in gray. The figures were generated using the program PyMOL (W. L. DeLano (2002) PyMOL, DeLano Scientific, San Carlos, CA.).

View larger version (45K): [in this window] [in a new window]   FIGURE 2. Multiple sequence alignment of representative members of the NAT family. Identical residues are colored in red in the alignment. Secondary structure elements of human NATs are assigned by the PROCHECK program (35) and are shown above the sequences and labeled: the helices are shown as cylinders, the strands are shown as arrow bars. Residues of the catalytic triad are labeled with stars; the residues interacting with CoA in NAT2 are labeled with red triangles; the residues involved in substrate binding are labeled with blue triangles. The alignment was generated using ClustalW (36) assisted with hand fittings.

View larger version (66K): [in this window] [in a new window]   FIGURE 3. Close-up stereo view of the active site of human NAT2 and NAT1. A, stereo view of CoA-binding site in human NAT2. CoA and the residues interacting with it are shown in stick-and-ball representation. The carbon atoms in CoA are gray, while the carbon atoms in NAT2 are colored in cyan. The residues of the catalytic triad (Cys-68, His-107, and Asp-122) are also shown in stick-and-ball representation, with the carbon atoms as gray. B, stereo view of substrate-binding site of human NAT1. The catalytic residues and the covalently bound acetanilide on Cys-68 are shown in stick-and-ball representation, with the carbon atoms as gray. The residues interacting with acetanilide are shown in stick-and-ball representation, with carbon atoms colored in green-cyan. The figures were generated using the program PyMOL.

	RESULTS AND DISCUSSION

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS AND DISCUSSION REFERENCES

The three-dimensional structures of NAT1 and NAT2 are similar to each other, and the C atoms of residues 1-290 can be superimposed with an r.m.s.d. of 0.7 Å. The fold of human NAT1 and NAT2 largely resembles the overall structure of their prokaryotic orthologues (Fig. 1), which is traditionally described in terms of three domains (20).

The amino-terminal domain (residues 1-83) consists of five helices (1-5) and one short -strand between helices 2 and 3. The second domain comprises residues 85-192 and consists of nine -strands (2-11 with two short helices 6 and 7). These two domains connect through the -helical inter-domain (helices 8-10) to the third domain, which has four anti-parallel -strands (12-15) and helix 11. The helix 11 precedes a stretch of well resolved residues that lead across the protein molecule's surface into a buried carboxyl terminus (residue 290). In human NATs, residues 167 and 183 bracket a 17-residue insertion, which is absent in the structures of prokaryotic NATs (Fig. 1 and 2). The insertion presented an obstacle in previous modeling attempts (21, 22) but is well resolved in the electron density of our crystal structures (supplemental Fig. S1). The function of the insertion in human NATs is not fully understood. However, a recent study has shown that the insertion contributes to the stability of human NATs (22). The first few residues of this insertion form a short -strand (10) and extend the previously mentioned 4-stranded anti-parallel -sheet (domain III) by parallel alignment to its fourth strand 15. The insertion also acts as a lid to bury the carboxyl terminus of the protein and makes several contacts with residues from 14 and 15 strands, 14 -15, 9-10, and 6-14 loops (not shown), and thus contributes to overall stability of the protein. Interestingly, the structures of NAT1 and NAT2 show a significant modification in the arrangement of their carboxyl-terminal residues when compared with prokaryotic NATs. Beginning at residue 274, the carboxyl-terminal helix observed in prokaryotic NATs is dissolved ("stretched out"). In contrast, the carboxyl terminus extends deep within the folded human NATs, into close proximity to the buried catalytic triad (Fig. 1) and capable of interacting with CoA (Fig. 3A), as described below. The human NAT1 and NAT2 fold shows no similarity to the GCN5-related N-acetyltransferase (23) and amino()-terminal acetyltransferase superfamilies (the crystal structure of NAT13 is available now, PDB code 2OB0).³

View larger version (29K): [in this window] [in a new window]   FIGURE 4. Binding of CoA to NAT2. Schematic diagram of the interaction between CoA (carbon atoms and bonds shown in gray) and human NAT2. The carbon atoms of residues that form hydrogen bonds with CoA are shown with bonds in cyan. Hydrogen bonds are drawn as dashed lines, and donor-acceptor distances are given. Residues involved in hydrophobic interaction are shown with arcs. The figure was generated by using the program LIGPLOT (37).

Substrate-binding Site—The crystal structure of NAT2 in complex with CoA contains two protein molecules per asymmetric unit that can be superimposed with an r.m.s.d. of 0.3 Å for C atoms. Each of the two polypeptide chains binds a CoA molecule in an equivalent position (Fig. 3A). Although the structures show no obvious similarity to acetyl-CoA-dependent N-acetyltransferases of the GCN5-related superfamily, the CoA molecules bind to the enzyme in a conformation similar to that found in GCN5-related acetyltransferases (23). The bound CoA is bent and wraps around the surface of the protein, positioning the sulfhydryl group of CoA in the cleft between the -helical inter-domain and the -barrel (domain II). The -mercaptoethylamine and pantothenate moieties are located 12 Å deep inside the cleft, placing the cofactor's sulfhydryl group close to that of the catalytic cysteine (Cys-68), with a sulfur-sulfur distance of nearly 2.74 Å (Fig. 4).

Hydrophobic residues Phe-37, Phe-93, Leu-209, Phe-217, and Leu-288 make an extensive set of van der Waals contacts throughout most of the length of the pantotheine arm of CoA and thus play a major role in orienting the reactive sulfhydryl group for acetyl transfer (Figs. 3A and 4). In addition, the carbonyl group of the pantothenate moiety establishes a hydrogen bond with the nitrogen atom of Ser-216. No other hydrogen or water-mediated hydrogen bonds are found between the pantotheine arm of CoA and the enzyme.

The pyrophosphate group of CoA makes a series of direct hydrogen bonds with the sequential amide nitrogens of residues Thr-103 and Gly-104 in the 3-4 loop and with the hydroxyl groups of Tyr-208 and Thr-214 (equivalent to Ser-214 in NAT1) from helix 9 and the 9-10 loop, respectively (Figs. 3A and 4). The crystal structure of the NAT2-CoA complex does not support previous indications that the pyrophosphate loop ("P-loop") starts at the Gly-126 residue of Salmonella typhimurium and proximal to the active site (20, 24).

The N6 of the adenine ring of CoA forms a single hydrogen bond with the side chain of Ser-287. The adenine ring also makes contacts with hydrophobic residues Pro-97 and Val-98. In NAT1, phenylalanine is in position 287, suggesting a mainly hydrophobic interaction of the adenine ring of CoA with NAT1 and NAT2.

Superimposition of the NAT2-CoA complex and NAT1_ F125S structures shows that CoA binding induces slight structural rearrangements that are confined to the 3-4 loop ("P-loop"). This loop moves by 1.8 Å toward the center of the cleft upon CoA binding.

The active site of NAT1 is clearly defined by the crystal structure of wild-type NAT1, which reacted with the irreversible inhibitor 2-bromoacetanilide, leading to acetanilide bound to the sulfur atom of the catalytic cysteine (Cys-68). The aromatic ring of the acetanilide molecule mimics substrates of NAT1. It stacks with the hydrophobic surface of the substrate-binding site formed by the aromatic ring of Phe-125 and the side chain of Val-93 (Fig. 5A). It also interacts with the side chains of other hydrophobic residues (Ile-106 and Phe-217). Phe-217 is highly conserved in many NATs, whereas residues Val-93, Ile-106, and Phe-125 in human NAT1 are replaced by Phe, Val, and Ser in NAT2. The hydrophobic environment is required for the binding of substrates. Our structure of the NAT1_F125S mutant with acetamide covalently bound to Cys-68 revealed a similar environment except for the substitution of Phe-125 with serine (Fig. 5B). As has been described previously (25), this substitution virtually abolishes NAT1 affinity preference for PAS over SMZ by reducing its affinity for the smaller PAS and increasing its affinity for the bulkier SMZ. Thereby, the substrate selectivity of NAT1_F125S resembles that of wild-type NAT2.

View larger version (84K): [in this window] [in a new window]   FIGURE 5. Substrate binding pockets of NAT1 and NAT2. A and B show active sites of wild-type and F125S mutant of NAT1, respectively. C and D show PAS modeled in the substrate binding pocket of NAT1 in surface and schematic representation, respectively. E and F show SMZ modeled in the substrate binding pocket of NAT2 in surface and schematic representation, respectively. The covalently bound acetanilide (in wild-type NAT1) and acetamide (in F125S mutant), PAS and SMZ are shown in stick-and-ball representation, with carbon atoms in gray, nitrogen in blue, oxygen in red, and sulfur in orange. The critical residues for substrate binding are shown in stick-and-ball representation, with carbon atoms colored either in green-cyan for NAT1 or cyan for NAT2 except the catalytic triad where the carbon atoms are gray. In schematic diagrams, PAS and SMZ are colored in red. The residues involved in substrate binding and specificity are labeled. Hydrogen bonds are drawn as dashed lines. The Van der Waals surfaces in the enzyme's pocket are shown as green-cyan (NAT1) or cyan (NAT2) planes.

First, the substrate binding pocket in NAT1 is smaller (162 ų) than that of NAT2 (257 ų). This is a consequence of two key residue substitutions at positions 127 and 129, namely Arg-127 and Tyr-129 in NAT1, whereas in NAT2, serine residues occupy these positions (Fig. 5, C and E). The effect of having two bulky groups protruding into the substrate-binding site is significant, in essence reducing the volume of the NAT1 pocket by almost 40% compared with that of NAT2 (Fig. 5, D and F). The second major substitution takes place at position 93, where NAT1 exhibits a valine and NAT2 a phenylalanine (Fig. 5, C and E). This substitution has the effect of introducing a "lip" in the Van der Waals surface of the binding pocket of NAT2, making it more selective for substrates that can accommodate this feature (Fig. 5, D and F). The F125S mutation does not change the volume of the substrate binding pocket to any significant measure, but rather alters the extent of the hydrophobic interactions with the pocket surface at this position.

Docking studies suggested a clear preference for substrate types as a function of the specific enzyme. Aminobenzyl compounds were identified to bind preferably to NAT1. In NAT1, the amino residue of PAS makes hydrogen bond contacts with His-107 and the backbone carbonyl of Ile-106. The acid motif in PAS was directed toward Arg-127, also making hydrogen bond interactions. The aromatic ring of PAS presented - staking interactions with the phenyl group of Phe-125 (Fig. 5D), much like the pose of the co-crystallized acetanilide fragment discussed above (Fig. 5A). In contrast, the sulfonamide class of compounds bound selectively to NAT2. This largely resulted from the presence of Phe-93 in NAT2, whose side chain introduces a bump in the Van der Waals surface of this enzyme's pocket (Fig. 5F). The sulfonamide series in the library of virtual substrates, by virtue of their hinged-like structure, were identified by the docking experiments as the set of compounds that can accommodate this substitution and capitalize on the most binding interactions. The results of these docking simulations are consistent with enzyme kinetic studies of wild-type and site-directed mutant forms of NAT1 and NAT2.⁴

View larger version (55K): [in this window] [in a new window]   FIGURE 6. Location of SNPs on human NATs. A, the overall fold of human NAT1 (left) and NAT2 (right) are shown in ribbon diagrams. The positions of unique amino acid substitutions in human NATs are shown in stick-and-ball representation with carbon atoms colored in yellow. The amino and carboxyl termini of both proteins are labeled as NT and CT, respectively. B, secondary structure and sequences of human NATs. The identical residues are shown as dots in NAT2. The SNPs are highlighted in red. The residue numbers are shown above the sequence, and the amino acid substitutions are shown below the sequences.

The crystal structures of NAT1 and NAT2 provide a rationale for the altered function of the NAT variants. Most of the human NAT variants have altered residues that are conserved between the two proteins, with a few exceptions (Fig. 6B). The positions of the point mutations in NAT1 and NAT2 are scattered throughout the entire primary structure, but with a majority of the substitution sites (Arg-117, Arg-166, Val-149, Ser-214, Met-205, and Glu-261 in NAT1; Ile-114, Glu-167, Arg-197, Lys-268, Lys-282, and Gly-286 in NAT2) located on the surfaces of the proteins and thus exposed to the cellular environment (Fig. 6A). Reduction in the enzymatic activity of several of these NAT variants is believed to be due to increased ubiquitinylation of the mutant proteins and their degradation by the 26 S proteasome, because activity reduction correlates with reduced protein level, while the transcription rate of the NAT genes and the enzyme kinetics are unchanged (4, 31).

The crystal structures of NAT1 and NAT2 show that few mutations are involved in intramolecular polar or hydrophobic interactions (R64W, E167Q, R187Q, and D251V in NAT1; R64Q or R64W, D122N, L137F, and Q145P in NAT2) (Fig. 7). In NAT1 and NAT2, Arg-64 is located on the 4-5 loop and makes hydrogen bond interactions with Glu-38 and Asn-41 (Fig. 7A). Substitutions of Arg-64 with Trp or Gln will destroy these interactions. The bulkier side chain of tryptophan will clash with the surrounding residues, thereby changing the local protein structure. Also, the surface-exposed hydrophobic side chain of tryptophan may lead to aggregation of the protein. This structural information provides the basis for previous observations that the recombinant protein product of NAT1^*17 (R64W) was found to be insoluble when expressed in E. coli and forms microaggregates in vivo (31).

The R187Q (NAT1^*14B) alloenzyme exhibits slow acetylation activity due to a substantially lower level of the expressed protein (4). Arg-187 forms a hydrogen bond with Glu-182, which is located in the 17-residue insertion unique to human NAT1 and NAT2 (Fig. 7B) and required for the stability of the human NATs (22). Therefore, the R187Q substitution most likely decreases the stability of the protein and lowers protein level thereafter. However, R187Q also decreases the affinity of NAT1 for PAS by 15-fold (32), suggesting that this mutation also alters the active site topology.

The D251V (NAT1^*22) substitution is located on strand 15, and Asp-251 makes contacts with Arg-242 and Asn-245 through hydrogen bonding (Fig. 7C). In D251V, changing the charged residue (Asp-251) to a hydrophobic residue (Val) would break the interactions and result in destabilization of the protein. This is in agreement with biochemical data that showed low catalytic activity associated with reduced protein expression in bacterial (33) and yeast (4) systems, which is apparently due to decreased stability of the protein.

View larger version (65K): [in this window] [in a new window]   FIGURE 7. Interaction network of particular mutations. All the residues that are involved in the interactions are shown in sticks, with the carbon atoms colored in green-cyan (in NAT1) and cyan (in NAT2). The carbon atoms in the mutated residues are in gray. The distance for hydrogen bonds are labeled. A, in both NAT1 and NAT2, Arg-64 is in contact with Glu-38 and Asn-41 through hydrogen bonding. B, Arg-187 of NAT1 forms hydrogen bond with Glu-182. C, Glu-167 of NAT1 is hydrogen bonding with Lys-185 and Asp-251. D, D251V (NAT1*22) makes contacts with Glu-167, Arg-242, and Asn-245 through hydrogen bonding. E, the catalytic residue Asp-122 of NAT2 forms hydrogen bonds with Asn-72 and His-107. F, Leu-137 of NAT2 is in contact with Trp-159 and Leu-194 through hydrophobic interactions. G, Gln-145 of NAT2 forms hydrogen bonds with Trp-132 and Gln-133.

The NAT2^*12D allele encodes a protein with two substitutions, D122N and K268R. Asp-122 is located on the 5-6 loop and is a member of the catalytic triad (Fig. 7F). Any changes here would be expected to affect the activity of the enzyme.

The NAT2^*17 (Q145P) alloenzyme exhibits slow acetylator phenotype due to reduced expression level rather than stability of the protein (4). Gln-145 is located in the 7-8 loop and makes interactions with Trp-132 and Gln-133 (Fig. 7G).

In conclusion, we have provided the first direct structural analysis of human NAT1 and NAT2, enzymes that play important roles in the biotransformation of drugs and environmental agents of the aromatic amine and hydrazine chemical classes. These structures should provide valuable tools for the further analysis of structural features that impart NAT catalytic selectivity and for the eventual identification and development of chemical agents that are predicted to be detoxified or bioactivated by these enzymes.

	FOOTNOTES

 
^* This work was supported by the Structural Genomics Consortium with funds from Genome Canada through the Ontario Genomics Institute, the Canadian Institutes for Health Research, the Canada Foundation for Innovation, the Ontario Challenge Fund, the Ontario Innovation Trust, the Wellcome Trust, GlaxoSmithKline, the Knut and Alice Wallenberg Foundation, and the Vinnova and Swedish Foundation for Strategic Research. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

The on-line version of this article (available at http://www.jbc.org) contains supplemental Fig. S1.

¹ To whom correspondence should be addressed: Structural Genomics Consortium, University of Toronto, 100 College St., Toronto, Ontario M5G 1L5, Canada. Tel.: 416-946-3868; Fax: 416-946-0588; E-mail: alexander.plotnikov{at}utoronto.ca.

² The abbreviations used are: NAT, arylamine N-acetyltransferase; r.m.s.d., root mean square deviation; SMZ, sulfathetamine; PAS, p-aminosalicylic acid.

³ A. Plontnikov, manuscript in preparation.

⁴ D. Grant, manuscript in preparation.

	ACKNOWLEDGMENTS

 
We thank Dr. Alexey Botchkarev for fruitful discussions and Dr. Cheryl Arrowsmith for critical reading of the manuscript. Data for this study were measured at beamline X25 of the National Synchrotron Light Source.

	REFERENCES

TOP ABSTRACT INTRODUCTION EXPERIMENTAL PROCEDURES RESULTS AND DISCUSSION REFERENCES

 

1. Riddle, B., and Jencks, W. P. (1971) J. Biol. Chem. 246, 3250-3258[Abstract/Free Full Text]
2. Weber, W. W., and Hein, D. W. (1985) Pharmacol. Rev. 37, 25-79[Medline] [Order article via Infotrieve]
3. Evans, D. A. (1989) Pharmacol. Ther. 42, 157-234[CrossRef][Medline] [Order article via Infotrieve]
4. Hein, D. W. (2002) Mutat. Res. 506-507, 65-77[Medline] [Order article via Infotrieve]
5. Guengerich, F. P. (1992) Pharmacol. Ther. 54, 17-61[CrossRef][Medline] [Order article via Infotrieve]
6. Grant, D. M., Goodfellow, G. H., Sugamori, K., and Durette, K. (2000) Pharmacology 61, 204-211[CrossRef][Medline] [Order article via Infotrieve]
7. Hein, D. W., McQueen, C. A., Grant, D. M., Goodfellow, G. H., Kadlubar, F. F., and Weber, W. W. (2000) Drug Metab. Dispos. 28, 1425-1432[Medline] [Order article via Infotrieve]
8. Hein, D. W., Doll, M. A., Fretland, A. J., Leff, M. A., Webb, S. J., Xiao, G. H., Devanaboyina, U. S., Nangju, N. A., and Feng, Y. (2000) Cancer Epidemiol. Biomarkers Prev. 9, 29-42[Abstract/Free Full Text]
9. Otwinowski, Z. M. W. (1997) Methods Enzymol. 276, 307-326[CrossRef]
10. Hendrickson, W. A., Horton, J. R., and LeMaster, D. M. (1990) EMBO J. 9, 1665-1672[Medline] [Order article via Infotrieve]
11. Schneider, T. R., and Sheldrick, G. M. (2002) Acta Crystallogr. D. Biol. Crystallogr. 58, 1772-1779[CrossRef][Medline] [Order article via Infotrieve]
12. Sheldrick, G. M. (2002) Zeitschrift für Kristallographie 217, 644-650[CrossRef]
13. McCoy, A. J., Grosse-Kunstleve, R. W., Storoni, L. C., and Read, R. J. (2005) Acta Crystallogr. D. Biol. Crystallogr. 61, 458-464[CrossRef][Medline] [Order article via Infotrieve]
14. Perrakis, A., Morris, R. M., and Lamzin, V. S. (1999) Nat. Struct. Biol. 6, 458-463[CrossRef][Medline] [Order article via Infotrieve]
15. Murshudov, G. N., Vagin, A. A., and Dodson, E. J. (1997) Acta Crystallogr. D. Biol. Crystallogr. 53, 240-255[CrossRef][Medline] [Order article via Infotrieve]
16. Davis, I. W., Murray, L. W., Richardson, J. S., and Richardson, D. C. (2004) Nucleic Acids Res. 32, 615-619[CrossRef]
17. Emsley, P., and Cowtan, K. (2004) Acta Crystallogr. D. Biol. Crystallogr. 60, 2126-2132[CrossRef][Medline] [Order article via Infotrieve]
18. Case, D. A., Cheatham, T. E., 3rd, Darden, T., Gohlke, H., Luo, R., Merz, K. M., Jr., Onufriev, A., Simmerling, C., Wang, B., and Woods, R. J. (2005) J. Comput. Chem. 26, 1668-1688[CrossRef][Medline] [Order article via Infotrieve]
19. Ewing, T. J., Makino, S., Skillman, A. G., and Kuntz, I. D. (2001) J. Comput. Aided Mol. Des. 15, 411-428[CrossRef][Medline] [Order article via Infotrieve]
20. Sinclair, J. C., Sandy, J., Delgoda, R., Sim, E., and Noble, M. E. (2000) Nat. Struct. Biol. 7, 560-564[CrossRef][Medline] [Order article via Infotrieve]
21. Zhang, N., Liu, L., Liu, F., Wagner, C. R., Hanna, P. E., and Walters, K. J. (2006) J. Mol. Biol. 363, 188-200[CrossRef][Medline] [Order article via Infotrieve]
22. Walraven, J. M., Trent, J. O., and Hein, D. W. (2007) Drug Metab. Dispos. 35, 1001-1007[Abstract/Free Full Text]
23. Dyda, F., Klein, D. C., and Hickman, A. B. (2000) Annu. Rev. Biophys. Biomol. Struct. 29, 81-103[CrossRef][Medline] [Order article via Infotrieve]
24. Sandy, J., Holton, S., Fullam, E., Sim, E., and Noble, M. (2005) Protein Sci. 14, 775-782[CrossRef][Medline] [Order article via Infotrieve]
25. Goodfellow, G. H., Dupret, J. M., and Grant, D. M. (2000) Biochem. J. 348, 159-166[CrossRef][Medline] [Order article via Infotrieve]
26. Lower, G. M., Jr., Nilsson, T., Nelson, C. E., Wolf, H., Gamsky, T. E., and Bryan, G. T. (1979) Environ. Health Perspect. 29, 71-79[Medline] [Order article via Infotrieve]
27. Okkels, H., Sigsgaard, T., Wolf, H., and Autrup, H. (1997) Cancer Epidemiol. Biomarkers Prev. 6, 225-231[Abstract]
28. Brockmoller, J., Cascorbi, I., Kerb, R., and Roots, I. (1996) Cancer Res. 56, 3915-3925[Abstract/Free Full Text]
29. Filiadis, I. F., Georgiou, I., Alamanos, Y., Kranas, V., Giannakopoulos, X., and Lolis, D. (1999) J. Urol. 161, 1672-1675[CrossRef][Medline] [Order article via Infotrieve]
30. Gu, J., Liang, D., Wang, Y., Lu, C., and Wu, X. (2005) Mutat. Res. 581, 97-104[Medline] [Order article via Infotrieve]
31. Liu, F., Zhang, N., Zhou, X., Hanna, P. E., Wagner, C. R., Koepp, D. M., and Walters, K. J. (2006) J. Mol. Biol. 361, 482-492[CrossRef][Medline] [Order article via Infotrieve]
32. Hughes, N. C., Janezic, S. A., McQueen, K. L., Jewett, M. A., Castranio, T., Bell, D. A., and Grant, D. M. (1998) Pharmacogenetics 8, 55-66[CrossRef][Medline] [Order article via Infotrieve]
33. Lin, H. J., Probst-Hensch, N. M., Hughes, N. C., Sakamoto, G. T., Louie, A. D., Kau, I. H., Lin, B. K., Lee, D. B., Lin, J., Frankl, H. D., Lee, E. R., Hardy, S., Grant, D. M., and Haile, R. W. (1998) Pharmacogenetics 8, 269-281[CrossRef][Medline] [Order article via Infotrieve]
34. Grant, D. M., Hughes, N. C., Janezic, S. A., Goodfellow, G. H., Chen, H. J., Gaedigk, A., Yu, V. L., and Grewal, R. (1997) Mutat. Res. 376, 61-70[Medline] [Order article via Infotrieve]
35. Laskowski, R. A., MacArthur, M. W., Moss, D. S., and Thornton, J. M. (1993) J. Appl. Crystallogr. 26, 283-291[CrossRef]
36. Thompson, J. D., Higgins, D. G., and Gibson, T. J. (1994) Nucleic Acids Res. 22, 4673-4680[Abstract/Free Full Text]
37. Wallace, A. C., Laskowski, R. A., and Thornton, J. M. (1995) Protein Eng. 8, 127-134[Abstract/Free Full Text]

CiteULike

Complore

Connotea

Del.icio.us

Digg

Reddit

Technorati

This article has been cited by other articles:

				P. J. Ballester, I. Westwood, N. Laurieri, E. Sim, and W. G. Richards Prospective virtual screening with Ultrafast Shape Recognition: the identification of novel inhibitors of arylamine N-acetyltransferases J R Soc Interface, July 8, 2009; (2009) rsif.2009.0170v1. [Abstract] [Full Text] [PDF]

				P. J. Murphy The Development of Drug Metabolism Research as Expressed in the Publications of ASPET: Part 3, 1984-2008 Drug Metab. Dispos., October 1, 2008; 36(10): 1977 - 1982. [Abstract] [Full Text] [PDF]

This Article Abstract Full Text (PDF) Supplemental Data All Versions of this Article: 282/41/30189    most recent M704138200v1 Submit a Letter to Editor Alert me when this article is cited Alert me when eLetters are posted Alert me if a correction is posted Citation Map Services Email this article to a friend Similar articles in this journal Similar articles in PubMed Alert me to new issues of the journal Download to citation manager Request Permissions Citing Articles Citing Articles via HighWire Citing Articles via Google Scholar Google Scholar Articles by Wu, H. Articles by Plotnikov, A. N. Search for Related Content PubMed PubMed Citation Articles by Wu, H. Articles by Plotnikov, A. N. Social Bookmarking                      What's this?