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Keywords:

  • drug metabolism;
  • arylamine N-acetyltransferase;
  • NAT;
  • crystal structure;
  • drug discovery

Abstract

  1. Top of page
  2. Abstract
  3. Introduction
  4. NAT Structure
  5. Catalytic mechanism
  6. Structure-activity relationships for NAT substrates
  7. Structure–activity relationships for NAT inhibitors
  8. NAT Polymorphisms
  9. Conclusion
  10. Acknowledgement
  11. Conflict of interest
  12. References

Arylamine N-acetyltransferase (NAT) plays an important role in metabolism and detoxification of many compounds including drugs and environmental carcinogens through chemical modification of the amine group with an acetyl group. Recent studies have suggested that NATs are also involved in cancer cell growth and inhibition of the enzymes may be a potential target for cancer chemotherapy. Three-dimensional (3D) structures are available for NATs from both prokaryotes and eukaryotes. These structures provide valuable insights into the acetylation mechanism, features of the active site and the structural determinants that govern substrate/inhibitor-binding specificity. Such insights allow a more precise understanding of the structure–activity relationships for NAT substrates and inhibitors. Furthermore, the structural elucidation of NATs has generated powerful tools in the design of small molecule inhibitors that should alleviate cancer, based on the important role of the enzyme in cancer biology.


Abbreviations
3D

three-dimensional

5-AS

5-aminosalicylic acid

ANS

anisidine

CoA

Coenzyme A

HDZ

hydrazines

NAT

Arylamine N-acetyltransferase

PABA

p-aminobenzoic acid

PAS

p-aminosalicylic acid

SMZ

sulfamethazine

TZD

thiazolidinedione

Introduction

  1. Top of page
  2. Abstract
  3. Introduction
  4. NAT Structure
  5. Catalytic mechanism
  6. Structure-activity relationships for NAT substrates
  7. Structure–activity relationships for NAT inhibitors
  8. NAT Polymorphisms
  9. Conclusion
  10. Acknowledgement
  11. Conflict of interest
  12. References

Arylamine N-acetyltransferases (NATs; EC 2.3.1.5) catalyse the acetylation reaction (generally classified as a Phase II process) by which an acetyl group from the cofactor acetyl coenzyme A (acetyl CoA) is transferred to the substrates, including drugs and carcinogens. NATs are present in most of living organisms (Butcher et al., 2002; Vagena et al., 2008) and NAT proteins have a characteristic motif – (Pro/Ala)-Phe-Glu-Asn-Leu [(P/A)FENL] (Payton et al., 2001). Although its endogenous functions remain largely unclear, NAT is essential for synthesis of cell wall in mycobacteria (Sim et al., 2008a). Human NATs include two isoenzymes, namely, NAT1 and NAT2. NAT1 is found in many tissues, particularly in the colon, whereas NAT2 is found mostly in the liver and intestine and is predominantly responsible for drug metabolism (Hickman et al., 1998; Sim et al., 2000). It acetylates many therapeutic agents such as isoniazid, dapsone and the sulphonamides.

In recent years, acetylation has been intensively studied due largely to its role in the carcinogenic activation of aromatic amines (Hein et al., 1993). These acetylated derivatives of arylamines and N-hydroxylated arylamines can undergo further oxidation or other reactions to form highly reactive cytotoxic and carcinogenic species (Hein et al., 1993; Hanna, 1994). Moreover, there is increasing evidence that NAT has an important role in folate catabolism and cancer cell biology (particularly for breast cancer) (Adam et al., 2003; Butcher et al., 2007; Sim et al., 2008b; Butcher and Minchin, 2012). In fact, human NAT1 has emerged as a new diagnostic marker or drug target for breast cancer (Laurieri et al., 2010; Russell et al., 2009; Tiang et al., 2010).

The first X-ray crystal structure for a NAT enzyme (from Salmonella typhimurium) was reported by Sinclair et al. (2000). To date, there are 11 crystal structures for NATs from human and five bacterial species (Table 1). This structural bank has provided valuable insights into the acetylation mechanisms and active site features. Importantly, it helps to explain why NATs selectively catalyse aromatic amines and exhibit overlapping yet distinct substrate specificity. Both human NAT1 and NAT2 are highly polymorphic enzymes. Many of the NAT polymorphisms translate to mutations in particular protein residues. The crystal structures have suggested individual function of the residues in the protein that has greatly improved the understanding of the phenotypes associated with NAT polymorphisms. The crystal structures represent not only a tool for predicting substrate selectivity but also a powerful tool in rational design of selective inhibitors of NATs for therapeutic purposes.

Table 1. List of available crystal structures for NATs (the structures can be found at http://www.pdb.org)
NATPDB entriesIn complex withReference
S. typhimurium NAT1E2T Sinclair et al., 2000
M. smegmatis NAT1GX3 Sandy et al., 2002
M. smegmatis NAT1W6FIsoniazidSandy et al., 2005b
M. smegmatis NAT (C70Q)1W5R Sandy et al., 2005a
Pseudomonas aeruginosa NAT1W4T Westwood et al., 2005
Mesorhizobium loti NAT12BSZ Holton et al., 2005
M. marinum NAT2VFB Fullam et al., 2008
M. marinum NAT2VFCCoenzyme AFullam et al., 2008
Human NAT1 (F125S)2IJAIodoacetamideWu et al., 2007
Human NAT12PQT2-BromoacetanilideWu et al., 2007
Human NAT22PFRCoenzyme AWu et al., 2007

NAT Structure

  1. Top of page
  2. Abstract
  3. Introduction
  4. NAT Structure
  5. Catalytic mechanism
  6. Structure-activity relationships for NAT substrates
  7. Structure–activity relationships for NAT inhibitors
  8. NAT Polymorphisms
  9. Conclusion
  10. Acknowledgement
  11. Conflict of interest
  12. References

The secondary and tertiary structures of the NATs are highly conserved among the enzymes from prokaryotes and eukaryotes (Westwood et al., 2006; Sim et al., 2008a). The NAT fold is usually described in terms of three domains (Sinclair et al., 2000; Wu et al., 2007). For example, in human NAT, the N-terminal domain [or the helical bundle in Sinclair et al., (2000)] consists of five helices (α1–α5) and one short β-strand between helices α2 and α3 (Figure 1). The second domain [or the β-barrel in Sinclair et al., (2000)] consists of 10 β-strands (β2–β11) and two short helices α6 and α7 (Figure 1). The two domains connect through the α-helical interdomain (helices α8–α10) to the third domain [or the α/β lid in Sinclair et al., (2000)], which has four anti-parallel β-strands (β12–β15) and helix α11 (Figure 1). The helix α11 precedes a stretch of residues that lead across the protein molecule's surface into a buried C-terminus (Figure 1).

figure

Figure 1. The tertiary and secondary structures of NAT. Panel A: The tertiary structure of NAT (using human NAT1 (PDB code: 2PQT) as an example), depicting the overall fold of a NAT structure. Panel B: Structure-based sequence alignment of human NATs (NAT1 and NAT2). The secondary structure elements are shown above the alignment. Conserved residues are highlighted in colour. The Protein Data Bank (PDB) codes for the structures are shown in parentheses. The triangles indicate the catalytic residues (please see text for details). This figure was produced using ENDscript (Gouet and Courcelle, 2002).

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Although a highly similar structural fold is shared by bacterial and human NATs, structural differences are evident. First, human NATs contain a 17-residue insertion (i.e. residues 167–183) between β9 and β11, which is absent in the structures of prokaryotic NATs (Wu et al., 2007) (Figure 2). The insertion includes the secondary structure elements β10 and α7. Although the function of this insertion in human NATs is not fully understood, the insertion does contribute to the stability of the protein by interacting with surrounding residues (Walraven et al., 2007). Second, human NATs show a significant structural divergence in the arrangement of their C-terminal residues compared with prokaryotic NATs (Figure 2). In human NAT, the C-terminal residues form a full coil and the terminus extends deep within the folded protein into close proximity to the buried catalytic triad and is capable of interacting with CoA (Wu et al., 2007) (Figure 2). By contrast, the C-terminus in prokaryotic NATs is in a helical configuration, and the helix is positioned away from the protein core (Figure 2).

figure

Figure 2. Structural comparison of bacterial NAT (panel A) and human NAT (panel B). The structures of NAT from Salmonella typhimurium (PDB code: 1E2T) and human NAT1 (PDB code: 2PQT) are shown. The insertion in the human NAT is indicated in purple. The structures of the C-terminal residues in both enzymes are shown in red.

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Active sites for cofactor and substrate binding

The crystal structures, in complex with CoA, for Mycobacterium marinum NAT and human NAT2 reveal that CoA binds differently to these two enzymes (Wu et al., 2007; Fullam et al., 2008). In human NAT, the CoA-binding site is located in a deep cleft formed between the interdomain (connecting domain II to domain III) helices α8–10 and the β-barrel (domain II) (Wu et al., 2007) (Figure 3A). CoA is bound to the active site in a bent conformation, positioning the sulfhydryl group of CoA towards the protein core and the 3′-phospho-ADP group on the surface of the protein (Figure 3A). The β-mercaptoethylamine and pantothenate moieties are located deep inside the cleft, placing the cofactor's sulfhydryl group close to that of the catalytic cysteine (C68), with a sulfur–sulfur distance of ∼3 Å (Wu et al., 2007). Binding of CoA to the active site is mainly mediated by hydrophobic and hydrogen-bond interactions (Figure 3B). The pantotheine arm of CoA makes an extensive set of van der Waals contacts with the non-polar residues F37, F93, L209, F217 and L288 (Figure 3B and C). The adenine ring also makes contacts with the hydrophobic residues P97 and V98. In addition, the carbonyl group of the pantothenate moiety forms a hydrogen bond with the nitrogen atom of S216. The pyrophosphate group of CoA makes a series of hydrogen bonds with the residues T103, G104, Y208 and T214. The N6 of the adenine ring of CoA forms a hydrogen bond with the side chain of S287 (Wu et al., 2007) (Figure 3B).

figure

Figure 3. The CoA binding site in human NAT (PDB code: 2PFR). Panel A: A full view of CoA binding to human NAT. Panel B: Molecular interactions of CoA with the binding site residues. Panel C: Chemical structure of acetyl CoA, showing its three components.

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Unexpectedly, a different CoA-binding site is present in the prokaryotic NAT (Fullam et al., 2008). Here, CoA binds to the deep cleft formed by domains II and III of the enzyme (Figure 4A). The binding site is located ∼30 Å from that in human NAT (Figure 4). Recognition of the pantotheinate arm of CoA is mainly through hydrophobic interactions with residues F38, Y69, Y71, H110, F130 and F204. The phosphate groups of CoA are hydrogen-bonded to residues such as W97 and K236. The adenine moiety of CoA interacts with the active site residues via a mixture of hydrophobic and polar contacts. Specifically, it forms hydrophobic, hydrogen-bond and ring-stacking interactions with residues V169, E152, H229 respectively (Fullam et al., 2008). The observation that human and prokaryotic NATs bind CoA in markedly different ways led Fullam et al. (2008) to hypothesize that the cofactor binding site may be a promising novel target for selectively inhibiting pathogenic prokaryotic NAT enzymes.

figure

Figure 4. Comparison of the CoA-binding site in bacterial NAT from M. marinum (PDB code: 2VFC; panel A) with that in human NAT2 (PDB code: 2PFR; panel B).

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The acetyl acceptor (substrate) binding site overlaps to a great extent with the CoA-binding site and this finding is consistent with the fact that the cofactor and the substrate bind to the enzymes in a sequential manner, a feature of the Ping Pong kinetic mechanism (see discussion below). Similar to the pantotheine arm of CoA, the entire substrate molecule binds to the deep position of the cleft formed between the helical interdomain and domain II (β-barrel) (Wu et al., 2007). The substrate-binding sites deduced from either prokaryotic or eukaryotic NATs consist mainly of hydrophobic residues (Figure 5A). This feature agrees with the fact that NATs have a preference for hydrophobic substrates.

figure

Figure 5. The substrate-binding site in human NAT. Panel A: A full view of the substrate-binding site (indicated as a blue surface) in human NAT (PDB code: 2PQT). Panel B: Comparison of the substrate-binding site residues of human NAT1 (in white; PDB code: 2PQT) with those of NAT2 (in green; PDB code: 2PFR). Residues at positions 127 and 129 play a critical role in determining the pocket size and the substrate specificity. Please see the text for details. The binding site is indicated as a yellow surface.

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Catalytic mechanism

  1. Top of page
  2. Abstract
  3. Introduction
  4. NAT Structure
  5. Catalytic mechanism
  6. Structure-activity relationships for NAT substrates
  7. Structure–activity relationships for NAT inhibitors
  8. NAT Polymorphisms
  9. Conclusion
  10. Acknowledgement
  11. Conflict of interest
  12. References

There is strong evidence that the acetylation reaction mediated by NATs proceeds via the double displacement (‘Ping Pong Bi Bi’) mechanism (Andres et al., 1983; 1988; Sinclair and Sim, 1997). There are two sequential steps to the reaction: firstly, the acetyl group is moved from acetyl CoA to form an acetylated enzyme intermediate, then the substrate is acetylated and CoA is released (Figure 6). Structural analysis reveals that the acetyl transfer in NATs is facilitated by a Cys-His-Asp catalytic triad (Cys68-His107-Asp122 in human NAT), which is strictly conserved in all known NATs and in identical positions in all NAT structures (Sinclair et al., 2000). The triad renders a stable thiolated form of cysteine that initiates the reaction by nucleophilic attack on the carbonyl of the acetyl moiety of acetyl CoA (Figure 6) (Wang et al., 2004; Sandy et al., 2005a). Consistent with their critical role in catalysis, mutations of any of the triad residues abolishes the enzyme activity (Dupret and Grant, 1992; Watanabe et al., 1992; Wang et al., 2004). Interestingly, the latest NAT structure (from M. marinum) presents an extended binding interface between CoA and the protein (Fullam et al., 2008). The authors propose that the nucleoside-phosphate moiety of CoA can remain associated with the protein, subsequent to acetylation of the catalytic cysteine, thereby protecting the acetylated enzyme intermediate from hydrolysis.

figure

Figure 6. The proposed catalytic mechanism of NAT showing the importance of the catalytic triad Cys68-His107-Asp122 (human NAT numbering) in initiating the acetylation reaction. The insert depicts the location of the triad residues according to the crystal structure of human NAT2 (PDB code: 2PFR).

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Wang et al. have proposed a model for the reaction chemistry in the NAT active site (Wang et al., 2004; 2005). In this model, a thiolate–imidazolium ion pair is formed between Cys68 and His107 with a pKa of 5.2. Asp122 is involved in the ionic interaction with His107 and is critical for optimal catalysis and structural integrity. Upon acetylation of the thiolate, half of the ion pair is lost with concomitant shift in the pKa of His107 to 5.5. The process of nucleophilic attack on the thiol ester and deacetylation of the thiolate is dependent on the nucleophilic strength of the arylamine substrate. For weak nucleophiles with a pKa <5.5, such as p-aminobenzoic acid (PABA, Figure 7A), a deprotonation step by His107 is required to make the substrates more nucleophilic for the attack on the thiol ester, which is followed by deacetylation of the thiolate. By contrast, those strong nucleophiles with a pKa ≥ 5.5 such as anisidine (ANS, Figure 7A) directly attack on the thiol ester; deacetylation of the thiolate occurs via deprotonation of a tetrahedral intermediate (Wang et al., 2005) (Figure 6).

figure

Figure 7. Substrate-binding site of NAT from M. smegmatis (MSNAT) and the structural determinants of its substrate preference. Panel A: Chemical structures of NAT substrates. Panel B: Diagram representation of MSNAT–isoniazid interactions (PDB code: 1W6F). Interactions with T109 and F130 are important in substrate binding.

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Structure-activity relationships for NAT substrates

  1. Top of page
  2. Abstract
  3. Introduction
  4. NAT Structure
  5. Catalytic mechanism
  6. Structure-activity relationships for NAT substrates
  7. Structure–activity relationships for NAT inhibitors
  8. NAT Polymorphisms
  9. Conclusion
  10. Acknowledgement
  11. Conflict of interest
  12. References

Human NAT1 and NAT2 exhibit an overlapping substrate specificity. Both enzymes display substrate preference for aromatic amines (Kawamura et al., 2005). This is largely explained by the hydrophobic feature of the active site and the presence of aromatic residues in the active site capable of forming ring-stacking interactions (Wu et al., 2007). However, distinct substrate specificity also exists between NAT1 and NAT2 (Kawamura et al., 2005).

Aminobenzyl compounds such as p-aminosalicylic acid (PAS; Figure 7A) were identified as binding preferentially to NAT1. In contrast, the sulfonamide class of compounds such as sulfamethazine (SMZ) bound selectively to NAT2 (Sim et al., 2008b). Wu et al. (2007) explored the molecular basis for this difference in substrate recognition between the two enzymes. The substrate binding pocket in NAT1 is smaller (162 Å3) than that of NAT2 (257 Å3) as a consequence of two key residue substitutions at positions 127 and 129, namely R127 and Y129 in NAT1, whereas in NAT2, serine residues occupy these positions (Figure 5B). The presence of two bulkier groups reduces the volume of the NAT1 pocket by ∼40% compared with that of NAT2. Furthermore, V93 in NAT1 is replaced by F93 in NAT2; this substitution introduces a bump in the van der Waals surface of the pocket in NAT2, thereby significantly altering the shape of the binding pocket (Figure 5B). Molecular docking reveals that PAS and SMZ are closely fitted into the active sites of NAT1 and NAT2 respectively (Wu et al., 2007). The shape and size of the substrate molecules are well matched to those of the binding pockets, which helps to explain the distinct substrate selectivity of the enzymes.

A mutagenesis study showed that F125S substitution virtually abolished NAT1 affinity preference for PAS over SMZ and substrate selectivity of the mutant NAT1 (F125S) resembled that of wild-type NAT2 (Hein, 2002). Structural analysis and molecular docking indicate that F125 plays a key role in binding of PAS to NAT1 as the side chain of F125 forms π-π stacking interactions with the benzene ring of PAS (Wu et al., 2007). Therefore, F125S mutation would reduce the enzyme's affinity for the smaller PAS due to the lack of the π-π stacking interactions (as main forces) orienting the substrate for catalysis. On the other hand, an increase in the pocket space allows accommodation of the bulkier SMZ in the active site with more favourable energy. This is consistent with the fact that the mutant has an increased affinity for SMZ compared with the wild type (Hein, 2002).

Human NAT1 acetylates PABA and 4-aminobiphenyl but not o-toluidine (Grant et al., 1991; Hein et al., 1993; Fretland et al., 1997). The NMR structures of human NAT1 reported by Zhang et al. (2006) provide an explanation of such substrate selectivity. Based on the resolved structures, PABA binds favourably to the active site, consistent with the fact that it is a good NAT1 substrate (Figure 8A). However, binding of o-toluidine to the enzyme is difficult because its methyl group engages in steric clashes with F125 (Zhang et al., 2006). Also o-toluidine lacks functional groups such as the carboxylic acid group of PABA that could provide favourable binding contacts, for instance, hydrogen bonding (Zhang et al., 2006). The important role of F125 in determining the acetylation of o-toluidine is supported by the finding that a mutant human NAT2 in which F125 is replaced by a serine, exhibits 3- to 10-fold higher acetylation capacity for the substrate than NAT1 (Hein et al., 1993; Zhang et al., 2006). Based on this knowledge, Zhang et al. (2006) predicted that 4-amino-3-methylbenzoic acid would be a worse NAT1 substrate than PABA due to the presence of the methyl group. Interestingly, this prediction agreed well with the experimentally determined substrate activities. The authors also explained why 4-aminobiphenyl is acetylated by NAT1 by docking of the substrate with the enzyme (Zhang et al., 2006), as 4-aminobiphenyl was easily accommodated in the active site (Figure 8B). Binding of 4-aminobiphenyl was largely driven by the hydrophobic interactions between residues V93, K100, I106, F125, L209, S215, V216 and F217 and the aromatic ring of the substrate molecule (Figure 8B).

figure

Figure 8. Predicted molecular interactions between p-aminobenzoic acid (PABA) and human NAT1 (Panel A; PDB code: 2DSS) and between 4-aminobiphenyl and human NAT1 (Panel B; PDB code: 2GWZ) based on the NMR model of human NAT1.

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Human NAT2 displays a preference for lipophilic arylamines and the acetylation rate increases as the alkyl chain length of alkoxyanilines increases (Kawamura et al., 2005). On the contrary, human NAT1 displays a decrease in activity with increasing alkyl chain length (Kawamura et al., 2005). It has been proposed that this is due to the smaller overall size of human NAT1 active site compared with that of human NAT2 (Kawamura et al., 2005; Westwood et al., 2006). Substrate profiling demonstrates that none of the arylhydrazine compounds are substrates of human NAT1, whereas these compounds can be acetylated by human NAT2 (Kawamura et al., 2005). However, the reason why human NATs show this difference in substrate selection is unknown.

Brooke et al. (2003a) investigated the metabolism of a series of 4-alkoxyanilines by bacterial NATs (S. typhimurium and M. smegmatis) and found that there was a marked increase in the rate of acetylation with increasing alkyl chain length (Hexyl > Bu > Et > Me). This result indicated that the lipophilicity of the substrate may be a contributory factor to the rate of acetylation, which is further confirmed by a strong correlation between the enzyme activities and the calculated partition coefficients (clogP) of the substrate molecules. The preference for acetylating more lipophilic substrates by the bacterial NATs is probably due to the lipophilic nature of three conserved phenylalanine residues in the active site (Brooke et al., 2003a). The dominant binding forces are contributed by the lipophilic–lipophilic and π-stacking interactions between the phenylalanine residues in the enzyme and the acceptor substrate.

The crystal structure for M. smegmatis NAT (MSNAT)-isoniazid (INH) complex has been used to explain the enzyme substrate selectivity towards hydrazines (HDZ) and arylamines (Sandy et al., 2005b). The arylhydrazines are the best substrates for MSNAT with relatively low Km values of <5 μM for HDZ and 7.3 μM for INH, whereas the arylamines have higher Km values – 586 μM for 5-aminosalicylic acid (5-AS), 1460 μM for ANS, 4500 μM for PAS, and 56 000 μM for PABA. This may be in part due to the formation of hydrogen bonds between the hydrazyl group and residues T109 and C70 (Figure 7B). In addition, HDZ has extra potential hydrogen bonding from the heterocyclic nitrogen to the backbone oxygen of G129 or F130 (Sandy et al., 2005b). The affinity of the arylamine series depends to some extent on the capacity to form interactions with T109 and F130. 5-AS is a good arylamine substrate because it can be well oriented to form hydrogen bonds with both the side chain of T109 and the main chain carbonyl of F130. PAS has a lower apparent affinity compared with 5-AS because it is difficult for T109 to form hydrogen bonds with the substrate molecule, although hydrogen bonds to F130 can be readily achieved (Sandy et al., 2005b). ANS has a lower Km value than PAS despite offering hydrogen bonds to neither T109 nor F130. In this case, the unfavourable ligand binding in the absence of hydrogen bonding might be partly compensated by the generally lipophilic character of the methoxy group. Such groups are preferred in NAT substrates, presumably due to the formation of hydrophobic interactions in the active site. PABA has no additional chemical group on the aromatic ring facing the T109 and has an even higher Km than that of PAS (Sandy et al., 2005b).

Structure–activity relationships for NAT inhibitors

  1. Top of page
  2. Abstract
  3. Introduction
  4. NAT Structure
  5. Catalytic mechanism
  6. Structure-activity relationships for NAT substrates
  7. Structure–activity relationships for NAT inhibitors
  8. NAT Polymorphisms
  9. Conclusion
  10. Acknowledgement
  11. Conflict of interest
  12. References

There is considerable interest in developing small molecule inhibitors for NATs because (i) the inhibitors may be used for therapeutic purposes in breast cancer and tuberculosis; and (ii) the inhibitors can be used to explore the functions of NATs in various organisms (Westwood et al., 2011). Russell et al. (2009) synthesized and evaluated a series of rhodanine (2-sulfanylidene-1,3-thiazolidin-4-one) derivatives as selective inhibitors of human NAT1. The (Z)-5-(20-hydroxybenzylidene)-2-thioxothiazolidin-4-one (Figure 9A) was one of the most potent inhibitors and this compound was docked into the crystal structure of human NAT1 to explore the inhibitor binding mode (Russell et al., 2009). The aryl substituent extends into a hydrophobic pocket surrounded by residues V93, F125, V216 and F287, providing a possible explanation for the preference for lipophilic substituents. Binding of the inhibitor is also stabilized by hydrogen bonds with residues R127 and T289. The additional hydrogen bonding between the 2′-hydroxyl of the inhibitor and the enzyme may provide an explanation for the relatively high potency of 2′-hydroxyl-substituted rhodanine over its 4′-hydroxyl-substituted counterpart (Russell et al., 2009).

figure

Figure 9. Chemical structures of three types of NAT inhibitors, rhodanine derivative (panel A), TZD-benzosultam derivative (panel B) and S-substituted 1,2,4-triazole (panel C). For TZD-benzosultam derivatives, the hydrophobicity of the N-substitution correlates with the binding affinity.

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Brooke et al. (2003a) identified the N-(2-naphthyl)-methyl substituted 1,1-dioxo-2,3-dihydrobenzo[1,2]-thiazine-4-ylidene thiazolidine-2,4-dione [N-(2-naphthyl)-methyl and 2,4-thiazolidinedione (TZD)-substituted benzosultam, Figure 9B] to be a weak inhibitor (IC50 = 78 μM) of MSNAT by high throughput screening of a proprietary small molecule library. Subsequent synthesis and screening of a series of TZD-sultam adducts revealed that the TZD moiety was essential for activity as well as the substitution on the sultam nitrogen (Brooke et al., 2003b). The most effective NAT inhibitors in this series were those incorporating hydrophobic substituents at the sultam nitrogen. Docking of the TZD-sultam adducts with the crystal structure of MSNAT revealed the inhibition mechanism and the detailed molecular interactions between the inhibitor and the enzyme. The sultam moiety was located adjacent to the catalytic residue cysteine, providing a rationale for the observed competitive inhibition. The substituents at the sultam nitrogen occupied a hydrophobic groove in the protein, with a tryptophan residue, W97, appearing to hold the compound in place (Brooke et al., 2003b). The imidazole moiety of H203 participated in a π-stacking interaction with the TZD moiety of the compounds, which is consistent with the observed inactivity of the precursor ketone and ketal derivatives (Brooke et al., 2003b).

A more recent study characterized the inhibitory potency of a series of 1,2,4-triazoles (with long chain aliphatic or planar aromatic substituents on sulfur; Figure 9C) against M. tuberculosis NAT (Westwood et al., 2010). Interestingly, the potency of NAT inhibition increased with extended chain length up to approximately 10 Å, with the S-4-phenylbenzyl and S-octyl derivatives being the most potent in the series. The S-decyl derivative has an approximate chain length of 13 Å and was a less potent inhibitor of NAT than the shorter chain S-alkyl derivatives. The IC50 value against NAT of the S-decyl derivative was 13.1 μM, compared with an IC50 value of 3.8 μM for the S-octyl-1,2,4-triazole. This trend in acetylation difference was explored by molecular docking of the S-substituted 1,2,4-triazoles with NAT from M. tuberculosis. All except the S-decyl derivative were predicted to bind in essentially identical orientations. The triazole ring formed π-stacking interactions with F130 and formed hydrogen-bonds with the backbone carbonyls of F130 and G131. The 4′-methylphenyl group in all molecules (except the S-decyl derivative) pointed towards the active site cysteine residue. The correlation observed between the chain length and NAT inhibitory potency was closely matched by the docking scores for the same series of compounds, indicating that the crystal structures for NATs are powerful tools for structural design of inhibitors and for understanding of the affinity differences of the ligands.

NAT Polymorphisms

  1. Top of page
  2. Abstract
  3. Introduction
  4. NAT Structure
  5. Catalytic mechanism
  6. Structure-activity relationships for NAT substrates
  7. Structure–activity relationships for NAT inhibitors
  8. NAT Polymorphisms
  9. Conclusion
  10. Acknowledgement
  11. Conflict of interest
  12. References

Acetylation was the first enzymic polymorphism to be investigated and was based on the observation that the anti-tubercular drug isoniazid (INH) caused a different level of neural toxicity across different populations (Grant et al., 1997; Walker et al., 2009). NAT polymorphisms have been linked to three types of acetylator phenotypes, namely, fast, intermediate and slow acetylators (Hein et al., 2000; Hein, 2006). It has been suggested that slow acetylators are most at risk from INH hepatotoxicity because the toxic metabolite acetylhydrazine is retained much longer in the liver, compared with fast acetylators (Ellard et al., 1978; Metushi et al., 2011). In NAT1, there are 27 variant alleles that are associated with 11 amino acid alterations, whereas in NAT2, there are 65 variant alleles associated with alterations of 22 amino acids. A complete list of the NAT alleles and their associated phenotypes can be obtained online (http://louisville.edu/medschool/pharmacology). Understanding of why and how the polymorphisms alter the activity of human NATs has been significantly advanced by the availability of the crystal structures for the enzymes (Wu et al., 2007). Based on the location and/or functional role of the original amino acids in the protein, the functional effects of NAT polymorphisms are well elucidated (Wu et al., 2007). Table 2 summarizes the correlations of the structural data and functional implications for the NAT mutations. For more detailed discussion, one is encouraged to read the reviews by Walraven et al. (2008a,b).

Table 2. Mutation sites of human NAT alleles and their location in the NAT proteins as well as the functional effects of the mutations
NATMutation siteNAT AlleleLocation in the proteinFunctional effectReference
NAT1R117NAT1*5On the surfaces of the proteinMutants may be subject to increased ubiquitinylation, leading to reduced protein level and reduction in the enzymic activity. Neither the V149I nor the S214A residue changes alter the structural stability of NAT1. No functional changes occur with M205V and E261K mutations.

Wu et al., 2007

Hein, 2002

Liu et al., 2006

Walraven et al., 2008a

V149NAT1*11A
NAT1*11B
NAT1*30
R166NAT1*5
M205NAT1*21
S214NAT1*11A
NAT1*11B
NAT1*11C
E261NAT1*24
R64NAT1*17On the α4-α5 loopR64 forms H-bonds with the neighbouring residues E38 and N41. The stability of the enzyme is compromised in the absence of these interactions.

Wu et al., 2007

Walraven et al., 2008a

NAT1*19B
E167NAT1*5At the beginning of β10E167 forms H-bonds with the neighbouring residues K185 and D251. The mutant may affect protein stability.Wu et al., 2007
R187NAT1*14AIn the 17-residue insertionR187 forms an H-bond with E182. Substitution of R187 most likely decreases protein stability and lowers protein levels. The mutant may also alter the active site topology.

Wu et al., 2007

Hughes et al., 1998

NAT1*14B
D251NAT1*22On the strand β15D251 forms H-bonds with the neighbouring residues R242 and N245. The mutant may break these interactions and result in destabilization of the protein.

Wu et al., 2007

Hein, 2002

Lin et al., 1998

I263NAT1*25In the α11No change in protein level or catalytic activity for the I263V mutant because the hydrophobic interactions of the residue with others are preserved without introducing steric clashes.Walraven et al., 2008a
NAT2I114NAT2*5On the surfaces of the proteinMutants may be subject to increased ubiquitinylation, leading to reduced protein level and reduction in the enzymic activity.

Wu et al., 2007

Hein, 2002

Liu et al., 2006

NAT2*14C/F
E167NAT2*10
R197NAT2*5E/J
NAT2*6
NAT2*14D
K268NAT2*5
NAT2*6C/F
NAT2*12
NAT2*14C/E-G/I
K282NAT2*18
G286NAT2*6I/J
NAT2*7
R64NAT2*7DOn the α4-α5 loopR64 forms H-bonds with the neighbouring residues E38 and N41. The stability of the enzyme is compromised in the absence of these interactions.

Wu et al., 2007

Walraven et al., 2008b

NAT2*14
NAT2*19
D122NAT2*12DOn the β5-α6 loopD122 is a member of the catalytic triad. Mutations of D122 would adversely affect the activity of the enzyme.

Wu et al., 2007

Walraven et al., 2008b

L137NAT2*5IOn the β6-β7 loopL137 makes contacts with residues L194 and W159 through hydrophobic interactions. The mutant may result in a change in secondary structure that could trigger degradation mechanisms.

Wu et al., 2007

Walraven et al., 2008b

Q145NAT2*17On the β7-β8 loopQ145 forms H-bonds with the neighbouring residues W132 and Q133. The mutant shows lower enzymic activity that may be due to reduced expression levels.

Hein, 2002

Wu et al., 2007

Conclusion

  1. Top of page
  2. Abstract
  3. Introduction
  4. NAT Structure
  5. Catalytic mechanism
  6. Structure-activity relationships for NAT substrates
  7. Structure–activity relationships for NAT inhibitors
  8. NAT Polymorphisms
  9. Conclusion
  10. Acknowledgement
  11. Conflict of interest
  12. References

NAT plays an important role in the biotransformation of many aromatic and heterocyclic amine drugs. In addition, it has been linked to cancer risk because of its roles in the metabolic activation of carcinogens and in cell growth and survival. This review has described the crystal structures for NATs and analysed the structural similarities and differences between prokaryotic and mammalian NATs. Furthermore, breakthroughs in understanding of the catalytic mechanism, substrate/inhibitor binding and polymorphisms were discussed from a structural perspective. Although the crystal structures represent a powerful tool for predicting substrate selectivity, they have also been very useful in rational design of selective inhibitors of NATs with great potential in cancer treatment.

References

  1. Top of page
  2. Abstract
  3. Introduction
  4. NAT Structure
  5. Catalytic mechanism
  6. Structure-activity relationships for NAT substrates
  7. Structure–activity relationships for NAT inhibitors
  8. NAT Polymorphisms
  9. Conclusion
  10. Acknowledgement
  11. Conflict of interest
  12. References
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