Structure-based drug design

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Structure-based Drug Design by Veerapandian, Pandi. New York Marcel Dekker, Inc., 1997.

ISBN: 0824798694 eBook ISBN: 0585157448 Subject: Drugs--Design. Drugs--Structure-activity relationships. Drugs--Conformation. Drug Design. Structure-Activity Relationship. Language: English

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Structure-based Drug Design Table of Contents Structure-Based Drug Design Preface Contents Contributors 1 Inhibitors of HIV-1 Protease 2 Structural Studies of HIV-1 Reverse Transcriptase and Implications for Drug Design 3 Retroviral Integrase: Structure as a Foundation for Drug Design 4 Bradykinin Receptor Antagonists 5 Design of Purine Nucleoside Phosphorylase Inhibitors 6 Structural Implications in the Design of Matrix-Metalloproteinase Inhibitors 7 Structure—Function Relationships in Hydroxysteroid Dehydrogenases 8 Design of ATP Competitive Specific Inhibitors of Protein Kinases Using Template Modeling 9 Structural Studies of Aldose Reductase Inhibition 10 Structure-Based Design of Thrombin Inhibitors 11 Design of Antithrombotic Agents Directed at Factor Xa 12 Polypeptide Modulators of Sodium Channel Function as a Basis for the Development of Novel Cardiac... 13 Rational Design of Renin Inhibitors

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Structure-based Drug Design

14 Structural Aspects in the Inhibitor Design of Catechol OMethyltransferase 15 Antitrypanosomiasis Drug Development Based on Structures of Glycolytic Enzymes 16 Progress in the Design of Immunomodulators Based on the Structure of Interleukin-1 17 Structure and Functional Studies of Interferon: A Solid Foundation for Rational Drug Design 18 The Design of Anti-Influenza Virus Drugs from the X-ray Molecular Structure of Influenza Virus Ne... 19 Rhinoviral Capsid-Binding Inhibitors: Structural Basis for Understanding Rhinoviral Biology and f... 20 The Integration of Structure-Based Design and Directed Combinatorial Chemistry for New Pharmaceut... 21 Structure-Based Combinatorial Ligand Design 22 Peptidomimetic and Nonpeptide Drug Discovery: Impact of StructureBased Drug Design Index

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1 Inhibitors of HIV-1 Protease Krzysztof Appelt Agouron Pharmaceuticals, Inc., San Diego, California I. Introduction Since the discovery of human immunodeficiency virus (HIV) as the causative agent of acquired immunodeficiency syndrome (AIDS), perhaps the largest and most powerful consortium of scientists ever assembled to tackle a single disease has been brought to bear on the problem of AIDS and its treatment. From an unprecedented wealth of information regarding the molecular biology and virology of HIV collected in recent years, it became possible to identify numerous intervention points in the viral life cycle that could be exploited in the development of drugs for AIDS therapy (for reviews see Reference 1, 2, and 3). Among these, the virally-encoded enzymes, in particular reverse transcriptase and protease, have emerged as the most popular targets. A separate chapter of this book is dedicated to the description of reverse transcriptase and its inhibitors [4]. For the purpose of introduction only, it should be noted that nucleoside inhibitors of reverse transcriptase (AZT, ddI, ddC, d4T, and 3TC) have been widely used in clinical practice since 1987. Since then it has become apparent that this class of agents, while slowing progression of disease in HIV-infected patients, is limited in both activity and the duration of the clinical responses produced. Therefore in the search for better anti-HIV agents, the focus of effort was expanded to include the search for clinically useful inhibitors of a second viral enzyme, namely the protease. In contrast to reverse transcriptase, for which activity is required prior to the integration of viral genetic information into the host cell chromosomes, the viral protease plays a key role late in the virus life cycle and inhibitors of this enzyme display equal anti-viral activity in chronic and acute infection models in vitro [5]. The HIV protease (HIV PR) is encoded by the 5' portion of the retroviral pol gene, which encodes all replicative enzymes. Viral structural proteins (p24,

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p17, p9, and p7) and replicative enzymes (protease, reverse transcriptase/ RnaseH, and integrase) are translated as either polyprotein P55-GAG, or a larger frameshift product P160-GAG-POL. In the process of virus assembly these polyproteins are proteolytically cleaved by the protease and this processing step, both in its timing and accuracy, is essential for the formation of infectious particles of HIV [6]. It was also shown early on that the inactivation of HIV PR, either by chemical inhibition or certain mutations, leads to the production of immature, noninfectious viral particles [7,8]. Structurally HIV PR is a 99-amino-acid protein translated initially as a central part of the P160-GAGPOL polyprotein precursor. The autocatalytic processing from the 160 kDa precursor is poorly understood, but most likely occurs during the process of budding of pre-formed viral particles from the host cell [9]. After release from the precursor polyprotein, HIV PR forms a homodimer and acts in trans to correctly process GAG and GAG-POL polyproteins—a process required for formation of the viral capsid and nucleoprotein core. Retroviral proteases such as HIV PR are the latest additions to the wellstudied family of aspartic proteases. This family of enzymes, which includes, among others, proteases such as pepsin, renin, and cathepsins D and E, has been intensely studied in the past, and the knowledge gained from studies of these enzymes allowed early inferences as to the structure and function of the dimeric HIV PR. Moreover, the intensive effort over the past two decades to make inhibitors of human renin provided impetus for the early design of inhibitors of HIV PR. In fact, some of the renin inhibitors have turned out to be effective inhibitors of retroviral aspartic proteases as well and have served as the starting point for drug design. As a result of this many early inhibitors of HIV PR were peptidyl in nature and the best known example of such compounds is Ro31-8959, better known as saquinavir, a hydroxyethylaminecontaining mimetic of a hexapeptide substrate [10]. This potent inhibitor of HIV PR was discovered using a substrate-based rational approach to drug design and displays extremely high in vitro activity against clinical isolates and laboratory strains of HIV. Saquinavir has been recently approved by the FDA for the treatment of AIDS in combination with nucleoside inhibitors of reverse transcriptase, and the discovery of this compound was the first breakthrough and the starting point for many other innovative designs. Determination of the crystal structures of HIV PR gave new impetus to the design of novel inhibitors. One measure of the intensity with which new inhibitors were designed or discovered is the total number of crystal structures of inhibitory complexes, currently exceeding 250, that have been determined over the past 5 years. Very detailed crystallographic analysis combined with extensive biochemical characterization and site-specific mutagenesis studies made HIV PR perhaps the best characterized enzyme to date.

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Based on the avalanche of papers describing the structure-based design of various HIV PR inhibitors, it would be reasonable to assume that, with the exception of saquinavir, all other HIV PR inhibitors that entered the stage of preclinical or clinical development were discovered using the elements of a structurebased approach. From the long list of more than 30 inhibitors considered as clinical candidates [11], currently there are three compounds (saquinavir, ritonavir, and indinavir) already approved by the FDA as anti-HIV drugs. Many factors that are requisite for in vivo activity in AIDS patients can only be predicted a priori in a very general sense. For instance, erratic oral bioavailability in humans, first-pass metabolism, binding to plasma proteins or tissue distribution may disqualify a perfect in vitro inhibitor of HIV replication and such properties can be very poorly predicted by any process of drug design. A potential answer to these problems is the parallel design of several chemically distinct compounds that may have similar in vitro activity but significantly different in vivo properties. The application of protein structure-based design offers such possibilities and in this text the discovery and optimization of different series of potent inhibitors of HIV PR will be discussed. In order to familiarize the reader with the architecture of HIV PR and the properties of its active site, the first paragraphs are devoted to the detailed description of the x-ray structures of the enzyme followed by several examples of inhibitors in a bound conformation. A. Three-Dimensional Structure of HIV PR Retroviral proteases such as HIV PR were tentatively assigned to the aspartic protease family on the basis of putative active-site sequence homology [12]. Mammalian aspartic proteases are bilobal, singlechain enzymes in which each lobe (or domain) contributes an aspartic acid residue to the active site [13]. The active site itself is formed at the interface on the N- and C-terminal domains and exhibits approximate two-fold symmetry. Since the retroviral proteases are only about one-third the size of the two-domain eukaryotic enzymes, they were hypothesized to function as dimers in which each monomer contributes a single aspartic acid to the active site [14]. Obligate homodimeric proteases, in addition to providing a regulatory mechanism to control activation of the enzyme, represent the most efficient use of genetic information which, in retroviruses, is naturally parsimonious. The crystal structures of HIV PR confirmed the predicted dimeric character of the enzyme [15,16] (Figure 1). In all published crystallographic investigations of the unliganded form of the enzyme, the monomers are related to each other by crystallographic two-fold symmetry and are necessarily identical. The general topology of the HIV PR monomer is similar to that of a single-domain

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Figure 1 Stereo view of the α-carbon backbone of HIV PR dimer. (a) The apoenzyme with flaps in the “open” conformation. (b) Inhibited form of HIV PR with flaps in a “closed” conformation. For clarity, the inhibitor is removed from the active site.

pepsin-like aspartic protease and consists of antiparallel β-strands and a short, two-turn α-helix connected by loops of varying length. The dimer interface is formed by an antiparallel β-sheet comprising two strands from each monomer. The hydrophobic residues from those β-strands and two symmetry-related α-helices form the core of the dimer. The dimer is further stabilized by a net of hydrogen bonds involving the residues around the catalytic aspartic acids. The active site is formed by the dimer interface and is composed of equivalent contributions of residues from each monomer. The substrate-binding cleft is bound on one side by the active site aspartic acid (Asp25/25') and on the other side by a pair of two-fold related, antiparallel β-hairpin structures, commonly referred to as “flaps.”

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The conserved active-site residues (Asp25, Thr26, and Gly27 from both monomers) form a symmetrical and highly hydrogen-bonded arrangement virtually identical to that described for pepsin [17]. The two aspartates are nearly coplanar with the “inner” carboxylate oxygens hydrogen bonded to the amide hydrogens of Gly27/27'. This designation (e.g. Gly 27/27') will be used throughout this text to indicate equivalent residues of the dimer. The two threonines are inaccessible to solvent and are hydrogenbonded to the main-chain amide groups of the other monomer, forming a rigid network called a “fireman's grip” [17]. As in the case of the structures of eukaryotic pepsins, there is electron density for a water molecule bound between the two carboxylates of the active-site aspartates. In the structure of the apo-form of HIV PR, the flaps from both monomers are related by crystallographic two-fold symmetry and can be considered as being in an open conformation. In the structures of related proteases from Rous Sarcoma Virus and HIV-2, the flaps are either crystallographically disordered or in a partly closed conformation [18]. This suggests that, in solution, in the absence of ligands, the flaps are rather flexible and that the stable conformation of the flaps observed in the crystal structure of the apo-enzyme of HIV PR could be considered to result from kinetic trapping during the crystallization process. In the apo-form of HIV PR, the active site residues are located at the bottom of a rather shallow groove. Upon binding an inhibitor, the protease undergoes significant structural changes, particularly apparent in the flap region. As a result, a tunnel-like site is formed, which runs diagonally across the dimer interface. The tunnel has a volume of approximately 1140 Å3 and is 23 Å long. Because of the dimeric nature of HIV PR, the active site has approximate two-fold symmetry with the dyad axis intersecting the plane of the catalytic aspartates. Along the active site tunnel, starting from the central aspartates, there are distinct subsites S1, S2, S3, and S4, and corresponding symmetry related subsites S1', S2' S3', and S4' (Figure 2). It should be noted that in this chapter, the convention of Schechter and Burger [19] will be used to describe enzyme specificity subsites (S1, S1', etc.) and the corresponding side chains of inhibitors (P1, P1', etc.). The boundaries of the subsites are formed by residues from both monomers of HIV PR. All subsites, with the exception of S4/S4', which are exposed to solvent, are bounded by mostly aliphatic side chains and have hydrophobic character. The borders of the S1/S1' subsites are formed by the side chains of Ile23/23', Ile50/50', Ile84/84', Pro81/81', the γ carbon of Thr80/80', carboxylates of the active site Asp25/25', and the carbonyl oxygens of Gly27/27'. The S2/S2' subsites are bounded by Val32/32', Ile50/50', Ile47/47', Leu76/76', Ala28/28', and the carboxylates of Asp30/30'. The S3/S3', subsites are partly exposed to solvent and are bordered by the side chains of Leu23/23', Val82/82', Pro81/81', and the guanidinium groups of Arg8/8', which form a salt bridge with the carboxylates of Asp29/29'. Most of

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Figure 2 Schematic representation of the specificity subsites of the HIV PR active site with bound peptidic inhibitor JG-365. Amino acids forming the boundaries of the particular subsites are shown.

the hydrogen bond donor and acceptor functional groups of the active site are located in an approximate plane that lies along the long axis of the tunnel and is somewhat perpendicular to the plane of the subsites. The hydrogen-bonding functionalities include the carboxylates of the catalytic aspartates, the carbonyl oxygens of Gly27 and Gly48, the amide nitrogens of Asp29' and Gly48, the carboxylate of Asp29', and the dimer symmetry-related groups on the other side of the active site. Additional groups capable of forming hydrogen bonds with ligands are located in the outer part of the S2/S2' subsite and include the amide nitrogens and the carboxylates of Asp30/30'. There are five conserved water molecules in the active site of HIV PR. Four of the waters are symmetrically distributed in the S3/S3' subsites and one, hereafter called Wat301, is located near the two-fold axis of the dimer and, in the presence of most inhibitors, is approximately tetrahedrally coordinated by the hydrogen bonds formed between carbonyl oxygens of the ligand(s) and the amide nitrogens of Ile50/50' of the flaps. In the ligand-bound form of HIV PR, Wat301 is completely inaccessible to solvent, and it has been speculated that its functional substitution could be energetically favorable [18] or at least may lead to discovery of novel nonpeptidic inhibitors [20]. Thus, there are 18 hydrogen bond donors or acceptors in

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the active site of HIV PR-16 that could form hydrogen bonds directly and two in which the interaction is mediated by the conserved Wat301. The total solvent accessible surface area of the eight subsites of the HIV PR active site is approximately 1150 Å2. Because of the large number of groups with hydrogen-bondforming potential, 450 Å2 of the surface has a polar character, and the nonpolar area of the subsites is slightly larger, approximately 700 Å2. B. Structural Flexibility of HIV PR In the process of viral assembly, HIV PR specifically cleaves nine cleavage sites on GAG and GAG-POL polypeptides [21]. Examination of the amino acid composition of the recognized substrate sites (Table 1) indicates their hydrophobic character and significant sequence variability. The loose specificity of HIV PR most likely reflects its functions in a world of reduced complexity within the confines of the budding virion. The length of the viral protein precursors (approximately 1500 amino acids) reduces the number of potential sequences the protease must discriminate from in selecting its nine cleavage sites. Therefore, HIV PR and other retroviral proteases are not enzymes that have evolved to carry out a single reaction at a rapid rate, but rather enzymes with minimum specificity required to cleave the viral precursors in a specific and orderly manner. The loose specificity requirements demonstrated by effective binding and catalytic processing of all nine sequences, albeit at different rates [22], was the Table 1 The Sequences of the Proteolytic Processing Sites of HIV-1 HIV-1 PR Cleavage sites

Scissile bond

P17/P24

V

S

Q

N

Y

P

I

V

Q

N

P24/P2

K

A

R

V

L

A

E

A

M

S

P2/P7

S

A

T

I

M

M

Q

R

G

N

P7/P1

E

R

Q

A

N

F

L

G

K

I

P1/P6

R

P

G

N

F

L

Q

S

R

P

TF/PR

V

S

F

S

F

P

Q

I

T

L

PR/RT

C

T

L

N

F

P

I

S

P

I

RT/RN

G

A

E

T

F

Y

V

D

G

A

RN/IN

I

R

K

V

L

F

L

D

G

I

P5

P4

P3

P2

P1

P1'

P2'

P3'

P4'

P5'

Schechter-Berger notation

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TF—transframe, PR—protease, RT—reverse transcriptase, RH—RNAse H, IN—integrase. The location of the processing sites in HIV-1 were determined by protein sequencing of HIV-1 virion proteins.

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first indication that the recognition subsites of the HIV PR can display flexibility upon binding of substrates or inhibitors. Early crystal structures of the HIV PR apo-enzyme and complexes with peptidic inhibitors showed several conformations of the active site forming flaps, which include the residues Met46/46' to Ile54/54' [15,16]. Increased availability of coordinates of HIV PR complexed with various inhibitors and crystallized in different crystallographic space groups allowed for more rigorous examination of domain movements and structural changes in the active site. The alignment of several crystal structures of HIV PR in a common frame of reference, which most commonly includes the region around the symmetryrelated active site triad Asp25/25'-Thr26/26'Gly27/27', will highlight those regions of the backbone where significant displacement occurs upon accommodating the individual inhibitors. Examination of the aligned structures, which included examples of all classes of inhibitors, indicated only small variation of the backbone and limited movements in the two binding loops, comprising residues Leu76-Ile84 from both monomers. The loops form the outer walls of subsites S1/S1' and S3/S3' with inward-facing hydrophobic side chains of isoleucines and valines. The flexibility of these loops, which in some cases can move outward by as much as 2.5 Å, has a significant impact on the volume of the specificity subsites, which in turn can accommodate corresponding P1/P1' and P3/P3' moieties of various sizes. Interestingly, the predominant resistance-causing mutations are located on the same loops and involve changes in residues Val82/82' and Ile84/84' (see below). It should be noted, that while the alignment of several crystal structures provides information about the flexible regions, the extent of flexibility of the residues around the HIV PR active site can be limited by crystal packing forces and may represent a crystallographic artifact. In all characterized crystal forms of HIV PR [23] the loops 76–84 and 46–56 participate in crystal lattice formation and the particular conformation of these loops can be driven by crystallization conditions or interactions with other molecules related by the crystallographic symmetry. C. Inhibitors of HIV PR In general, inhibitors of HIV PR can be divided into three distinct groups. The first group includes peptidic inhibitors that utilize various transition-state dipeptide analogs such as statine, hydroxyethylene, and hydroxyethylamine incorporated into peptidic frameworks of differing lengths. Several crystal structures of this type of inhibitor complexed with HIV PR were solved and the structural information provided a wealth of information as to the minimum size of inhibitors, geometry of hydrogen bonds formed within the active site, and the structural flexibility of the subsites (for reviews see Reference 18 and 23). The second and perhaps largest group of HIV PR inhibitors includes peptidomimetic

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compounds that utilize similar transition-state analogs and retain at least one peptide bond with a side chain corresponding to a naturally occurring amino acid. Several compounds from this group have excellent pharmacokinetic and antiviral properties and, in fact, all three HIV PR inhibitors approved for clinical use (saquinavir, ritonavir, and indinavir) belong to this class of compounds. The last and the smallest group of HIV PR inhibitors has a distinct nonpeptidic character. Compounds from this class were discovered either by screening libraries of existing compounds or by structure-based de novo design. Illustrative examples of inhibitors belonging to all three classes and a brief description of the discovery of selected compounds are presented below. D. Peptidic Inhibitors of HIV PR The concept of peptidic inhibitors of HIV PR can be exemplified by the crystal structure of the statinecontaining peptidic compound AG1002 (Figure 3) [23]. In AG1002, the statine moiety replaces the scissile dipeptide while the flanking amino acids were derived from the natural substrate cleaved by HIV PR. The inhibitor binds to the active site in an extended conformation with the central hydroxyl group of the statine moiety forming hydrogen bonds with the active-site aspartic acids 25/25'. The peptidic backbone and the side chains of the

Figure 3 Stereo view of the peptidic inhibitor AG1002 bound to the active site of HIV PR. The distribution of the specificity subsites S and S' is similar to that shown in Figure 2. The boundaries of the HIV PR active site are indicated by the dotted surface.

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inhibitor form 16 hydrogen bonds and occupy subsites from S4 to S1, S2', and S3'. The carbonyl oxygens of P2 and P1' accept two hydrogen bonds from the flap water Wat301, which in effect is nearly tetrahedrally coordinated. Due to the structural nature of statine, which lacks the P1' side chain, the S1' pocket remains unoccupied. The S1 subsite is only partially filled by the P1 side chain of leucine. The P2 and P2' side chains of asparagine and glutamine form hydrogen bonds with Asp30' and 30, while the aliphatic carbons of both side chains make several hydrophobic contacts in the S2 and S2' pockets respectively. Despite the large number of hydrogen bonds formed within the HIV PR active site, AG1002 has rather low inhibitory potency with a binding constant of 0.55 µM The low binding constant most likely reflects the absence of the P1' group, the free energy required for desolvation of the hydrophilic side chains, and the charged N- and C-termini as well as entropic effects caused by the flexible nature of the heptapeptide. Other interesting examples of peptidic inhibitors are compounds utilizing other transition-state analogs, e.g. reduced amide-containing hexapeptide MVT-101 [24], hydroxyethylene-containing octapeptide U85548e [25], and hydroxyethylamine-containing heptapeptide JG-365 [26]. All these compounds bind to the active site of HIV PR in a similar extended conformation and the small differences in the geometry of hydrogen bonds formed with HIV PR can be attributed to the different character and length of the transition-state analogs. The chemical structures and inhibition constants of these inhibitors are summarized in Table 2. Note that the inhibition constants cited throughout this chapter and in Tables 2, 3, and 6 were determined in different laboratories—often using significantly different assay conditions—and therefore might not be meaningfully comparable. Due to their substantial size and peptidic nature, inhibitors from this class were not suitable for clinical application. Nevertheless, the structural information derived from many crystal structures of peptidic inhibitors bound to the HIV PR active site was critical for subsequent modeling and design of the next generation of peptidomimetic and nonpeptidic inhibitors of HIV PR. E. Peptidomimetic Inhibitors of HIV PR Design and Structure of Ro-31-8959 (Saquinavir) The strategy of designing saquinavir was based on the transition-state mimetic concept, an approach that has been used successfully in the design of potent inhibitors of renin and other aspartic proteases [10]. From the variety of nonscissile transition-state analogs of a dipeptide, the hydroxyethylamine mimetic was selected because it most readily accommodates the amino acid moiety characteristic of the Phe-Pro and Tyr-Pro cleavage sequence of the

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retroviral substrates. In the first step of design, the dipeptide analog consisting of Phe[CH(OH)CH2N]Pro was used to determine the minimum sequence required for potent inhibition. From this study a compound was selected that included benzyloxycarbonyl at the N-terminal side of the inhibitor followed by the P2 asparagine, the hydroxyethylamine isostere with side chains of phenylalanine and proline in the P1 and P1' positions respectively and the NH-t-butyl group at the Cterminal part. In the following design, the side chain of proline was consequently modified to a piperidine and finally to a decahydroisoquino-

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line moiety, and the N-terminal benzyloxycarbonyl group was replaced by the quinoline-2-carbonyl. The resulting compound, Ro-31-8959, was one of the first peptidomimetic inhibitors with very high antiviral potency and became a benchmark for further design of HIV PR inhibitors [10]. The high-resolution crystal structure of saquinavir bound to the active site of HIV PR was solved in many laboratories [23,27]. The incorporation of decahydroisoquinoline moiety, which can be considered as a conformationally restrained mimic of cyclohexylalanine, has some important consequences. First, the length of the C-terminal part of the inhibitor has been restricted to the P2' residue which, in saquinavir, consists of a NH-t-butyl group. Second, it restrained the conformational freedom of the otherwise peptidic backbone, minimizing the entropic penalty to the free energy of binding. In the crystal structure of saquinavir with HIV PR (Figure 4), the decahydroisoquinoline in the preferred chairchair conformation, makes extended hydrophobic contacts in the S1' subsite. The bond between the methylene carbon and the nitrogen of decahydroisoquinoline is in the low-energy equatorial conformation and the nitrogen, even if protonated, is not in a position to form a hydrogen bond with the active-site residues. The central hydroxyl group is in the R(syn) conformation and is within the hydrogen-bond-forming distance with both carboxylates 12640-0012a.gif Figure 4 Stereo view of the peptidomimetic inhibitor Ro 31-8959 (saquinavir) bound to the active site of HIV PR. The distribution of the specificity subsites S and S' is identical to that shown in Figure 2. Note the stacking interaction between the quinoline moiety and the P1 side chain of phenylalanine.

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of the active-site aspartates. Similar to Ag1002 and other peptidic inhibitors, the carbonyl oxygens of the P2 and P1' amides are within hydrogen-bonding distance of the flap water molecule; however, the geometry of the second hydrogen bond is distorted due to the additional spacing between both carbonyl groups. The nitrogen of the t-butylamide is displaced from the normal P2' position by approximately 1.8 Å and, as a result, cannot form a direct hydrogen bond with the carbonyl oxygen of Gly27. Instead the tbutylamide nitrogen interacts via highly ordered water molecules with the amide nitrogen of Asp29 and the carbonyl oxygen of Gly27. The aliphatic t-butyl moiety occupies the S2' subsite and the position of the backbone in this region prohibits any further extension into the S3' pocket. The P1 and P2 side chains of phenylalanine and asparagine, respectively, occupy the corresponding subsites and have a similar conformation to the equivalent groups observed in peptidic inhibitors. In the crystal structure, the N-terminal quinoline-2-carboxylate is moved to the side and, as a result, the carbonyl oxygen forms hydrogen bonds with the ordered water molecule and with the amide nitrogen of Asp29'. The quinoline ring is in a low-energy conformation with respect to the preceding carbonyl oxygen, which places the aromatic nitrogen in unfavorable close contact (3.3 Å) to the carbonyl oxygen of the flap Gly48. Because of the absence of any further contacts with the HIV PR active site residues, the contribution of the quinoline moiety to the free energy of binding remains unclear. Perhaps in solution, a stacking interaction of the P1 phenyl ring and the aromatic quinoline restricts the conformational freedom of Ro31-8959, in effect diminishing the free-energy loss due to the entropic and desolvation effects. Saquinavir, despite its distinct peptidomimetic character is a very potent inhibitor of HIV PR with an inhibition constant of 0.9 nM and an antiviral IC50 in vitro of 0.020 µM [10]. Although it suffers from a low oral bioavailability (5–10% in humans), it became an important starting point for the design of second generation, less-or nonpeptidic inhibitors. Saquinavir became the first HIV PR inhibitor approved by the FDA for treatment of AIDS. Design and Structure of ABT-538 (Ritonavir) An interesting concept for designing specific HIV PR peptidomimetic inhibitors with internal two-fold symmetry was first formulated by John Erickson and his colleagues from Abbott Laboratories [28]. They reasoned that if HIV PR incorporates symmetry into its active site structure, compounds that mimic this symmetry might be novel, more specific, and potent inhibitors and, furthermore, due to the bidirectionality of peptide bonds, might be sufficiently less peptidic in character and pharmacologically superior to the classical peptide-based compounds. The crystal structure of one of the first compounds from this series (A74704) verified the assumption of symmetrical binding conformation in the

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active site of HIV PR. The inhibitor consists of the central diamino alcohol moiety with symmetrically distributed phenylalanine side chains and two flanking, Cbz-blocked, valine residues. Except for the asymmetric hydroxyl group, A74704 binds to the active site in a symmetric mode and the inconsistent distribution of the terminal Cbz groups is most likely caused by crystal lattice contacts and may not reflect the binding mode in solution [28]. The design of symmetrical inhibitors was further extended to include a series of diamino, diol core units, in which the C2 axis bisects the bond connecting the two hydroxy-bearing carbon atoms [29]. Such inhibitors consistently showed greater potency than A74704, but the relative potencies of the diols differed for different diastereomers, and they did not exhibit a uniform dependence on the stereochemistry at the hydroxymethyl position. Surprisingly, high-resolution crystal structures of HIV PR with all possible diol diastereomers, (S,S, R,R and R,S) revealed that most of the inhibitors bind in a clearly asymmetric fashion placing only one of the diol hydroxyl groups on the C2 axis dissecting the active site of HIV PR and the catalytic carboxylates of Asp25/25'. The asymmetric placement of diols causes translation of inhibitors along the long axis of the active site and, as a result, the midpoint of the compounds is displaced by up to 0.9 Å from the two-fold axis of the HIV PR. Nevertheless, the dihedral angles of the symmetry-related bonds are in most cases within 10°, and the inhibitors maintain overall symmetry in the bound conformation [23,29]. The ABT-538 design was a direct consequence of the pioneering work with peptidomimetic compounds with the internal C2 symmetry [30]. Since the high-resolution crystal structures of a family of diolcontaining compounds indicated that in most cases only one of the diol hydroxyls interacts with the catalytic aspartic acids 25/25', in subsequent designs the noninteracting hydroxyl group was replaced by a hydrogen. This substitution reduced the free energy penalty required for desolvation of the hydroxyl group and increased the inhibitory potency without perturbing the binding mode of the compounds [30]. In the further search for related inhibitors with improved oral bioavailability, the focus of effort concentrated on the effect that molecular size, aqueous solubility, and hydrogen-bonding capability has on pharmacokinetic behavior. This study resulted in a smaller compound, A-80987, in which the P2' side chain of valine was eliminated and the terminal 2-pyridinyl group was replaced by 3-pyridinyl moiety that makes van der Waals contacts in the S2' subsite and forms a hydrogen bond with the amide nitrogen of Asp30 [31]. The pharmacokinetic properties of A-80987 were significantly improved over larger, symmetrical compounds from this series and, at the same time, the high antiviral activity typical for these inhibitors was largely unaffected. In subsequent optimization, which focused on the metabolic stability of these inhibitors in vivo, the electron-rich and oxidation-prone pyridinyl groups were replaced by thiazoles.

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Thiazoles are less electron-rich isosteres of pyridines and therefore it was speculated that compounds with such substitution may have improved metabolic stability [30]. The modeling of A-82200 in which the N-terminal pyridinyl group was substituted by a 4-thiazolyl moiety indicated that the 5-membered ring binds in the S3 subsite and can be further derivatized at the 2 position by an isopropyl group. The isopropyl functionality makes van der Waals contacts with Val82 and fills the hydrophobic part of the S3 subsite in nearly optimal fashion.

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The resulting compound, ABT-538 (Table 3), binds to the active site of HIV PR in an extended conformation. The central, asymmetric hydroxyl group is within hydrogen-bonding distance of the catalytic aspartates 25/25', and the P1/P1' phenylalanine side chains are symmetrically distributed in the corresponding subsites. The nitrogens of the symmetric amide bonds on both sides of the central aminoalcohol are barely within the hydrogen-bonding distance of the carbonyl oxygens of Gly27/27' (3.4 Å) and the carbonyl oxygens of these amide bonds participate in the tetrahydral coordination of the flap water molecule Wat301. On the N-terminal side of the compound, the P2 side chain of valine fills the S2 subsite and the terminal 2-isopropyl-4-thiazolyl makes hydrohobic contacts with the residues in the S3 pocket and has a stacking interaction with the P1 phenylalanine. On the C-terminal side, the 5thiazole is positioned to interact within the S2' subsite, and the nitrogen on the 5-membered ring is within hydrogen-bonding distance of the amide of Asp30'. Despite two peptide bonds present in ABT-538, this compound has substantial oral availability in humans and a very high antiviral activity in vivo [30]. Recently, ABT-538, better known as ritonavir, has been approved by the FDA for treatment of AIDS in combination with inhibitors of the reverse transcriptase. Design and Structure of L-735,524 (Indinavir) Indinavir is another example of very potent peptidomimetic compound discovered using the elements the crystal structure-based design [32] and SAR (structure activity relationship). The starting point for the design was a series of compounds containing the hydroxyethylene isostere of a scissile dipeptide [33]. An example of compounds from these series is L-685,434, which consists of a tert-butylcarbamate group forming the P2 moiety, symmetrically distributed phenylalanine side chains in the P1/P1', and the indanol group in the P2' portion of the inhibitor. Although very potent, the optimized molecules from this series lacked aqueous solubility and an acceptable pharmacokinetic profile [32]. The Merck group hypothesized that incorporation of a basic amine-containing functionality, such as the decahydroisoquinoline group of saquinavir, into the backbone of L-685,434 series might improve the solubility and bioavailability of this type of compound. Also the replacement of the P2/P1 functionalities, the tert-butylamide and phenylalanine side chain by the decahydroisoquinoline tertbutylamide, would generate a novel class of hydroxylaminepentanamide isostere with potentially improved metabolic stability in vivo. An additional strong argument for using decahydroisoquinoline as an isostere of P1/P2 moieties was the restricted conformational freedom of the enclosed-into-a-ring basic amine, which should decrease the entropy change upon binding to HIV PR in a similar fashion to that observed in saquinavir. In

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the resulting chimeric inhibitor the central hydroxyl group forms hydrogen bonds with the catalytic aspartic acids 25/25' and the hydrophobic side chains of the P1/P1' decahydroisoquinoline and phenylalanine respectively are separated from the central hydroxyl-bearing carbon by the methylene linkers forming a pseudosymmetrical arrangement. In the subsequent optimization of inhibitors from this novel series, a smaller piperazine group was substituted for the decahydroisoquinoline group, which offered a possibility to expand from the N4 position to the partially lipophilic S3 subsite. One of the first compounds from the piperazine series possessed a benzyloxycarbonyl moiety attached to the piperazine ring and the additional hydrophobic interaction in the S3 subsite was reflected by substantial increase in both intrinsic potency and in the ability to inhibit viral spread in infected cells in vitro. Finally, the replacement of the benzyloxycarbonyl group by the 3-pyridylmethyl moiety (Table 3) provided both lipohilicity for binding to the HIV PR active site and a weakly basic nitrogen that increased aqueous solubility and oral bioavailability. The crystal structure of L-735,524 (indinavir) bound to the active site of HIV PR [34] indicates that the 3-pyridylmethyl group attached to the N4 position of the piperazine ring makes hydrophobic contacts with the residues in the S3 and S1 pockets and the tert-butyl moiety fills the S2 subsite in the fashion previously observed in the structure of saquinavir. The positions of the P2 and P1' carbonyls maintain the proper alignment to form hydrogen bonds with the flap water Wat301. The terminal indanol group of indinavir occupies the S2' subsite with the hydroxyl group within hydrogen-bonding distance of the amide nitrogen of Asp29. The high aqueous solubility and largely nonpeptidic character of indinavir may be responsible for the good oral bioavailability, respectable pharmacokinetic profile, and high antiviral activity observed with this compound. Similar to saquinavir and ritonavir, indinavir has been recently approved by the FDA for treatment of AIDS. F. Nonpeptidic Inhibitors of HIV PR The nonpeptidic inhibitors of HIV PR can be divided into two subclasses. Compounds that belong to the first group maintain the general binding mode of the peptidomimetic inhibitors including formation of the key hydrogen bonds with the active site residues. An example of such nonpeptidic inhibitors of HIV PR is AG1343 (nelfinavir). The second group of nonpeptidic HIV PR inhibitors includes compounds with a binding mode significantly different from that described for the peptidomimetic compounds. Most inhibitors in this latter class were initially discovered by screening natural-products libraries or by structurebased de novo design. The most interesting examples of the nonpeptidic inhibitors from this group are the independently discovered but structurally

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related 4-hydroxypyrans and 4-hydroxycoumarins, the cyclic urea-based DMP323 series, and AG1284. Design and Structure of AG1343 (Nelfinavir) Analysis of the crystal structure of saquinavir with HIV PR indicated that while the nonpeptidic components of the ligand, namely the decahydroisoquinoline and the t-butylamide moieties fill the S1' and S2' subsites nearly optimally, the N-terminal portion offered the possibility for remodeling, aimed at the elimination of the peptidic character. Also, the contribution of the quinoline to the binding affinity to HIV PR was difficult to rationalize. Since the removal of quinoline resulted in a nearly 1000-fold loss in binding constant, it was concluded that the stacking interactions of the P1 phenyl ring and the P3 aromatic moiety of quinoline are necessary for the conformational stability of Ro 31-8959. In an attempt to redesign the N-terminal part of the ligand, the nonpeptidic portions of the P1' and P2' were maintained but for reasons of synthetic accessibility, the decahydroisoquinoline moiety was replaced by an orthosubstituted benzylamide [35]. Crystallographic analysis of both compounds showed that saquinovir and the modified LY289612 bind essentially identically to the active site of HIV PR and their inhibition constants and antiviral activity were very similar (Table 4 and Table 3). In the first attempt to functionally substitute the P2 side chain of asparagine, the isophthalic-acidcontaining compound was modeled and the low-energy conformation of the aromatic ring, required for binding in the S2 subsite, was stabilized by a tertiary carboxamide in the P3 region of the inhibitor [36]. The analysis of the binding mode and interactions of the isophthalic ring in the S2 subsite indicated a lipophilic pocket deep on the border between the S2 and S1' subsites, which could be conveniently filled with a methyl group extending from the 2 position of the ring. The resulting compound II in Table 4 lost most of the peptidomimetic character of LY289612 but retained its inhibitory potency. In an independent line of design, the relationship between the P1 phenylalanine side chain and the P3 quinoline was investigated. In the crystal structure of saquinavir bound to the active site of HIV PR (Figure 4), the aromatic ring of the P1 phenylalanine makes several van der Waals contacts with residues forming the S1 subsite. Computer modeling indicated that an extension of the phenylalanine side chain to phenethyl (homophenylalanine) would lead to prohibitive close contacts of the phenyl ring with the aliphatic side chains of HIV PR. On the other hand, replacement of the γ-carbon of the homophenylalanine by sulphur, which has a more acute C-S-C bond angle, would direct the aromatic ring into the neighbouring S3 subsite without changing the desired lipophilic nature of the P1 side chain. The increased area of hydrophobic interactions in the S1 and S3 subsites by compounds with the Sphenylcysteine and

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S-naphthylcysteine derived side chains in P1 resulted in a substantial increase in the inhibition constants [37]. The increase in the binding affinity to the low picomolar range in enzyme inhibition assay, allowed for subsequent truncation of the P3 quinoline moiety. The final compound from this miniseries (compound III in Table 4) consisted of the ortho-substituted benzamide in the P1' and P2', Snaphthylcysteine in P1 and asparagine in P2. Despite reduced molecular weight, the inhibition constant of this compound for HIV PR was comparable to LY289612. The observation that a larger, nonpeptidic moiety in the P1 could eliminate the need for the P3 side chain led to hybrid molecules that incorporated ring structures as the P2 component and maintained the P1 S-naphthylcysteine side chain of compound III. In this miniseries several bicyclic functionalities were modeled as the P2 substituents and one example, compound IV utilizing a tetrahydroquinoline group, is shown in Table 4 [38]. In subsequent modeling, it was noticed that the P2 bicyclic functionality might be replaced by 2,3-disubstituted phenyl rings. In particular, a methyl substitution in position 2 would increase the area of hydrophobic interaction in a manner previously observed in the isophthal series. Addition of a hydrophilic functionality attached at position 3 could increase the solubility of the compound and contribute to the binding constant by forming a hydrogen bond with the carboxylate oxygen of Asp30. A compound with a 2-methyl-3-hydroxy substitution pattern was synthesized and showed an improved inhibition constant of 3 nM in the HIV PR enzyme assay (Table 4). The crystal structure of compound V with HIV PR was solved and indicated the predicted binding mode with the possibility of a stacking interaction between the P2 phenyl and the P1 thio-naphthyl groups and the expected hydrophobic and hydrogen-bonding interactions of the P2 moiety with the protein side chains in the corresponding specificity pocket [38]. As with the optimized compounds from other series, compound V suffered from low aqueous solubility. The replacement of the P1' aryl group by the basic amine-containing decahydroisoquinoline dramatically increased the solubility and allowed for truncation of the P1 S-naphthylcysteine side chain to S-phenylcysteine without any loss of inhibitory activity. The resulting compound VI, AG1343 or nelfinavir, has an inhibition constant of 1.9 nM in the HIV PR enzyme assay and respectable antiviral activity with an IC90 of 60 nM [39]. The nonpeptidic character, pH-dependent solubility profile, and the small molecular weight of nelfinavir may contribute to its good pharmacokinetic profile in humans [40,41]. Currently, this compound is being tested and is in the advanced phase of clinical trials. The crystal structure of nelfinavir bound to the active site of HIV PR is shown in Figure 5. The general binding mode of this compound, in particular the path of the backbone, is similar to the binding mode of peptidyl inhibitors. Nevertheless, the lack of any peptide bonds utilizing naturally occurring amino

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Figure 5 Stereo view AG1343 (nelfinavir) bound to the active site of HIV PR.

acids qualifies nelfinavir to be a member of the group of nonpeptidic inhibitors of HIV PR. The unique, and perhaps crucial hydrogen-bonding interaction of the P2 hydroxyl group with the carboxylate oxygen of Asp30, combined with the smaller area of hydrophobic contacts in the S1 and S3 specificity subsites are the principal differences from other clinically active HIV PR inhibitors and may contribute to a distinct resistance pattern and point to additional utility of nelfinavir in the treatment of AIDS. Design and Structure of DMP323 A cyclic urea-containing HIV PR inhibitor, DMP323, was discovered using de novo structure-based design principles. Similar to the concept of Erickson and his co-workers from Abbott Laboratories, the group from DuPont-Merck attempted to take advantage of the two-fold symmetry of HIV PR in designing compounds that maintained the interaction of the diol with the catalytic aspartic acids 25/25' and at the same time were able to functionally displace the ubiquitous flap water molecule Wat301. They hypothesized that incorporation of the binding features of this structural water molecule into an inhibitor would be beneficial because of the entropic gain due to its displacement and because the conversion of a flexible linear inhibitor into a rigid, cyclic structure with restricted conformation should provide an additional, positive entropic effect. In the initial design, a cyclohexanone with the ketone oxygen as the structural

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water mimic was used and in subsequent synthetic targets the cyclohexanone ring was enlarged to a 7membered ring to incorporate a diol functionality. This target was further modified to a cyclic urea, which can be symmetrically substituted from both nitrogens without creating unnecessary stereocenters. The crystal structures of about 10 cyclic-urea-based inhibitors with HIV PR were solved [42]. In all cases, the C2 symmetric inhibitors bind to the HIV PR active site with the diad symmetry axes of the protease and the compounds being nearly coincident. The 7-membered ring of the inhibitors is roughly perpendicular to the plane of the catalytic aspartates 25/25' and both hydroxyl groups of the diol are positioned to interact with their carboxylates. The carbonyl oxygen of the inhibitors accepts hydrogen bonds from backbone amides of symmetrically distributed residues Ile50/50' of the flap. In the structure of DMP323, symmetrically substituted moieties of hydroxymethylbenzyls and phenylalanines extend towards the S2/S2' and S1/S1' subsites respectively and are involved in van der Waals interactions with the hydrophobic residues of these pockets [42]. The interaction of DMP323 with the residues of HIV PR are restricted to the central four specificity subsites of the active site. Despite this limited area of hydrophobic interaction and hydrogen bonding restricted to the central cyclic urea functionality, DMP323 is a very potent inhibitor of HIV PR with good antiviral activity in vitro (Table 5). The limited solubility of this compound was perhaps responsible for erratic oral availability in humans, and after short trials, DMP323 was withdrawn from the clinical investigation. Nevertheless, the discovery of this class of compounds represents a very interesting and, by now, classical example of de novo structure-based drug design. Design and Structure of AG1284 Another compound discovered by the application of de novo structure-based design is AG1284 [43]. In the initial design of a lead compound, the nonpeptidic hydroxyethyl-t-butylbenzylamide portion of LY289612 occupying the S1' and S2' subsites was retained as a “starting module.” In attempting to fill the pockets related by the dimer two-fold symmetry it was discovered that, by extending a two-carbon fragment from the central hydroxyl carbon, the S1 subsite could be accessed by an aromatic ring. The ring was oriented orthogonal to the observed P1 phenyl group of the classical inhibitors and this allowed further extension off the ortho position towards the S2 subsite. In order to maintain the critical hydrogen bond to the flap water Wat301, in the initial compounds an acylated amino group was used, replaced in subsequent designs by a benzamide functionality. In this model, the geometry of the hydrogen bonds to the flap water was somewhat perturbed, and the nitrogens of the t-butyl amides on both sides of the compound were in a position to interact favorably with solvent,

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potentially lowering the desolvation penalty. The absence of hydrogen-bonding interactions with the carbonyl oxygens of Gly27/27' was viewed as a positive factor, since the accumulated structural and mechanistic information suggested that formation of these hydrogen bonds may not be energetically favorable [23]. The compound was synthesized as a racemic mixture of two enantiomers of the central hydroxymethyl group and had the inhibition constant of 24 µM. Despite the modest binding constant of compound II (Table 6) and very low water solubility, the co-crystal structure with HIV PR was solved at 2.3 Å resolution, providing a critical starting point for further design. The inhibitor was found to bind largely as anticipated with the two aromatic rings occupying the S1 and S1' subsites and the two benzamide carbonyls forming hydrogen bonds to the flap water Wat301. Both benzamide nitrogens interact via a string of highly ordered water molecules with the amide nitrogens of Asp29/29'. The crystal structure of the complex indicated that the S enantiomer was the more active component of the racemic mixture and this was confirmed by stereoselective synthesis of subsequent compounds [44]. In the subsequent designs, the ortho-substituted benzyl rings were consecutively replaced by larger naphthyl groups that occupied more of the S1–S3 and S1'–S3' subsites. The increased area of hydrophobic interactions with the residues in these subsites was reflected in a substantial improvement in the

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binding constant and also in reduced aqueous solubility. Also, due to the very tight fit of both naphthyl moieties in the S1 and S1' subsites, subsequent design targeting the S3/S3' subsites proved to be difficult and synthetically challenging [44]. In the search for a simpler solution, the di-tertiary amides were designed using the crystal structure of compound II (Table 6) as a starting model. Branching from the amide nitrogens provided an interesting possibility to access S2–S3/S2'–S3' subsites while simultaneously increasing the solubility and stability of the compounds. In the first design, the hydroxyethyl moieties were fused to the amide nitrogens and the hydroxyl groups were intended to form hydrogen bonds with the amide nitrogens of Asp29/29' (compound III in Table 6). The addition of both hydroxyethyl groups resulted in a rather significant increase in the binding constant, and the racemic mixture had the Ki of 1.1µM. When the crystal structure of compound III complexed with HIV PR was solved at 2.2 Å resolution, it was observed that the inhibitor had undergone an inversion in binding mode relative to the secondary amide series. The phenyl groups of compound III occupied the S2/S2' subsites, switching positions with the t-butyl groups, which were in turn occupying the S1/S1' pockets (Figure 6). Due to this change in binding mode, the R enantiomer would be expected to be preferred relative to S. The final position of the hydroxyethyl moieties was less effected by the change, and both hydroxyls were within hydrogen-bonding distance from the amide nitrogens of Asp 29/29'. In the S2/S2' pockets, the phenyl groups occupied only a fraction of subsites, but the interaction was strengthened by highly ordered water molecules involved in electrostatic interaction with the aromatic rings and by forming hydrogen bonds to Asp30/30'. Interestingly the position of the hydrogen bonds with respect to the flap water was significantly disturbed in the new binding mode, and the conserved Wat301 was no longer tetrahydrally coordinated [43,45]. This unanticipated change in binding mode presented a potential for new avenues of design different from those of the secondary amides. The ability to design into neighboring subsites depends to a large extent on the positions of bond vectors suitable for substitution in the bound conformation of a given inhibitor. These vectors in the crystallographically discovered new binding mode of compound III were positioned ideally to access unfilled space in the S3/S3' pockets. The discovery of this new conformation of compound III highlighted the power of crystallographic feedback in the process of inhibitor design and, without this structural information, further design in this series would have been severely impeded. Inspection of the crystal structure of compound III bound to the active site of HIV PR revealed lipophilic cavities extending off the S1/S1' subsites adjacent to the t-butyl groups of the benzamidine moiety. The cavities are bordered by flexible loops around Pro81/81' and previous crystallographic studies indicated that both loops can move back by up to 2.5 Å, extending the size and

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Figure 6 Change of the binding mode of compound III observed during iterative design of AG1284. (a) Crystallographically determined binding mode of compound II. Pseudosymmetrically distributed aryl groups are bound in the S1 and S1' specificity subsites. (b) Crystallographically determined binding mode of compound III. Note the inversion of the binding mode. The ortho-substituted benzyl groups bind in a pseudosymmetric fashion in the S2 and S2' subsites.

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volume of the active site. With this in mind, the dimethylbenzyl group was attached to compound III and the additional phenyl ring was accommodated well in the lipophilic pocket of the S1'/S3' sides. As the S1 pocket was not fully occupied, a Monte Carlo-based De Novo Ligand-Generating program (MCDNLG) [46] was used to identify other amide substituents that might fill this subsite more effectively. From several moieties identified by the MCDNLG program, a larger cyclopentylethyl group showing very good shape complementarity to the S1/S3 subsite was selected for synthesis. In addition, due to the asymmetrical nature of this compound, additional space was identified at the bottom of the S2' pocket that was conveniently filled with either a methyl or a chlorine group on the 5 position of the benzamidine ring. The inhibition constant of the resulting compound (compound IV in the Table 6) was 0.008 µM, which represents approximately a 2500-fold improvement over the first compound from this series. The crystal structure of compound IV or AG1284 complexed with HIV-1 PR was solved, revealing excellent complementarity between the ligand and protein. The ligand forms only 4 hydrogen bonds with either protein functional groups or ordered water molecules, in contrast to the nine hydrogen bonds formed by peptidomimetic LY289612, despite their similar binding affinities. The nonpeptidic character of AG1284 may have contributed to good oral bioavailability and pharmacokinetics in three animal species [43]. Despite very good inhibitory potency on the enzyme level, AG1284 has rather modest antiviral activity in vitro (Table 6). The reason for this discrepancy is unclear but could be related to the low water solubility and higher affinity for membranes, which may effect cell partitioning. A similar lack of correlation between the potency of enzyme inhibition and antiviral activity has been previously observed with other HIV PR inhibitors [11]. Hydroxypyrans and Hydroxycoumarins The lead compounds for the 4-hydroxypyran and 4-hydroxycoumarin series were discovered in biological screens as low potency inhibitors of HIV PR [47–49]. Successful structure solution of both lead compounds with HIV PR enabled rapid optimization of their enzyme inhibitory potencies and antiHIV activities, and one of these compounds, U96988, has already entered Phase I clinical testing [49,50]. The binding mode of this type of inhibitor differs substantially from the classical peptidomimetic compounds and is somewhat similar to de novo-designed compounds from the cyclic urea series. In the case of 4-hydroxycoumarin, the two oxygen atoms of the lactone functionality are positioned within hydrogen-bonding distance of the two NH amides of Ile50/50' on the flap, replacing the ubiquitous water molecule Wat301. The 4-hydroxyl group (Table 5) is located within hydrogenbonding distance of the two catalytic

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aspartic acid residues Asp25/25' and this hydrogen-bonding network of the 4-hydroxycoumarin defines the essential pharmacophore of this new class of inhibitors. In the structure of U96988, this pharmacophore is pseudosymmetrically subsituted by an ethyl and a phenyl group at the C-3a and an ethyl and a benzyl group at the C-6a positions. These four substituents extend into the central core of S2/S2' subsites, where they make van der Waals contacts with the hydrophobic residues of the active site [49]. With a molecular weight of 362 U96988 is the smallest inhibitor of HIV PR in clinical testing. It suffers from rather low antiviral activity (ED90 of ~ 10 µM)but can be considered as the first in a series of this promising class of nonpeptidic HIV PR inhibitors. II. Structural Basis of Resistance of HIV PR Inhibitors The dimeric character and the two-fold symmetry of the active site, in which the monomers contribute equivalent residues to symmetrically distributed specificity subsites, led to early speculations that HIV PR may be less susceptible to resistance than, for example, reverse transcriptase. In the case of retroviral proteases, a single base mutation in the viral genome corresponds to two changes in the threedimensional structure and two structurally identical changes in the active site could result in an enzyme with a drastically modified specificity profile and impaired catalytic activity. Identification of HIV PR variants in cell-culture experiments clearly indicated, however, that this class of drugs is not immune to the challenge of viral resistance. It should be stressed that HIV, unlike other human viruses, is characterized by a dynamic viral turnover in the steady state [51,52]. The rapid replication rate coupled with the lengthy duration of infection will favor the emergence of resistant mutants to targeted antiviral agents [53]. The accumulated data from cell-culture sequential-passage experiments with several structurally different inhibitors and from the resistant variants identified during clinical exposure to four HIV PRtargeting drugs indicate a very complex pattern of mutations in the structure of HIV PR. In contrast to mutations in the reverse transcriptase, which frequently cause multihundredfold resistance [54], single base changes in the HIV PR gene (i.e., two identical substitutions per protease dimer) lead in most cases to 5–10-fold decrease in the antiviral potency of a given drug [11]. It has been shown for the most clinically studied HIV PR inhibitors, such as indinavir and ritonavir, that the clinical manifestation of resistance (increase in the viral load and decrease in the CD4 count) requires the simultaneous appearance of several mutations [55,56]. For example the resistant HIV strain isolated from patients exposed for 40 weeks to indinavir carried mutations at residues 10/10'L > R, 46/46'M > I, 63/63'L > P, 82/82'V > T, and 84/84'I > V [59,60]. However, the combination of these five

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Figure 7 Cartoon representation of the HIV PR dimer. The sites of primary resistance-causing mutations in the active site are indicated. For clarity, the names of the residues are shown for one monomer only.

mutations (ten assuming the dimeric nature of HIV PR) changed the susceptibility of the resistant strain to indinavir by only eight-fold if compared to the wild type HIV. The resistance-causing mutations are localized in a few “hot spots” in the structure of HIV PR and can be divided into two groups. The first group consists of the primary mutations located directly in the active site and includes changes at residues Val82/82', Ile84/84', somewhat less frequently at Gly48/48' and, in the case of nelfinavir, Asp30/30' (Figure 7). Residues 82/82' and 84/84' are located on the flexible loops that form the outer walls of the S3/S3' and S1/S1' subsites, respectively. In the resistant variants, valine 82/82' is most frequently substituted by the smaller side chain of alanine or the larger side chains of phenylalanine or isoleucine [57,58]. The change in position 82/82' is usually accompanied by a substitution of Ile84/84', most commonly to the smaller amino acids alanine or valine [57]. From the clinically tested compounds, ritonavir and indinavir, which were optimized to form strong hydrophobic interactions with the side chain of Val82/82' in the S3 subsite, suffer most significantly from any change at this position. On the other hand, the antiviral activities of saquinavir, and nelfinavir, which do not form any interaction in the S3/S3' subsite are not affected by mutations at Val82/82' and are only marginally cross resistant to changes involving Ile84/84' [57,58,62]. The resistance-causing

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mutation of Asp30/30' to asparagine seems to be specific for nelfinavir and was initially observed in cellculture sequential passage experiments [62]. Recently, the same phenotypic change was confirmed as the predominant mutation in the resistant variants appearing in AIDS patients exposed to low doses of this HIV PR inhibitor [64]. The molecular basis of resistance involving this mutation can be rationalized as follows: in the crystal structure of nelfinavir with the wild type HIV PR, the 3-hydroxyl group of the 2,3-substituted phenyl group is within hydrogen-bonding distance of the carboxylate oxygen of Asp30 in the S2 subsite. Due to the expected coulombic character of this interaction, the hydrogen bond formed with the negatively charged carboxylate of Asp30 would be expected to be a relatively strong one. The change of the negatively charged carboxylate of Asp30/30' to the amide oxygen of the asparagine side chain should reduced the strength of this interaction. Apparently the loss in the enthalpic contribution to the free energy of binding is only partially balanced by the entropic gain caused by the difference in desolvation of a charged vs. neutral side chain of the receptor, leading to decreased binding affinity of nelfinavir and eventually to viral resistance. An additional resistance-causing mutation that qualifies as a primary mutation involves the change of Gly48/48' to valine. This particular mutation seems to be specific for saquinavir and was observed both in cell-culture sequential passage experiments and in AIDS patients exposed to this inhibitor [61,65]. Located on the lower strands of the active-site forming flaps, Gly48/48' can be considered a part of the S4/S4' subsites. The replacement of the glycine hydrogen by the rigid side chain of valine has most likely a dual effect: first it has a direct impact on the interaction of the quinoline moiety of saquinavir with the active site of HIV PR, and second it may change the mobility of the flaps, which in turn will effect the binding kinetics of the natural substrates or inhibitors. Although none of the other clinically tested inhibitors form any interaction with this part of the flap, the HIV variants with mutation of Gly48/48' seem to be cross-resistant to all compounds, which is reflected by a 3–5-fold reduction of their antiviral activity [55,62]. While the effect of primary mutations on reduced binding affinities of inhibitors can be at least partially explained in view of the accumulated structural data, the function of secondary, or compensatory mutations in the resistant HIV PR is difficult to rationalize as yet. The predominant compensatory mutations observed in the resistant variants involve residues Leu63/63', Ala71/71', Met46/46', Asn88/88', Leu10/10', and Leu90/90' (Figure 8) [60,63]. Changes of these residues alone do not confer viral resistance, but their appearance increases the viability of the virus carrying the primary mutations in the active site of protease. All these residues are located far away from the active site of HIV PR do not participate in any apparent way in the inhibitor binding and it seems unlikely that they form a longrange interaction with the natural

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Figure 8 Cartoon representation of the HIV PR dimer. The sites of compensatory mutations are indicated.

substrate. Also, the reported compensatory mutations are conservative in nature and have no effect on the overall distribution of atomic charges on the surface of HIV PR. Sequence polymorphism at the Leu63/63' position, located on the surface at the base of HIV PR, has been observed in clinical isolates of the virus not exposed to any HIV PR inhibitors. Variations of Ala71/71', where the side chains are buried very close to Leu63/63', are less commonly found in clinical isolates. After a prolonged challenge by HIV PR inhibitors, Leu63/63' changes to proline and Ala71/71' to valine. The side chains of Met46/46' are fully exposed to solvent and these residues are located on the βhairpins that form the active side flaps. It has been speculated that the compensatory change of Met46/46' to isoleucine or phenylalanine may affect the dynamics of the flap movement, which in turn could influence the rates of catalytic activity of HIV PR impeded by the primary mutations in the active site [58]. Any changes to Asn88/88' and Leu90/90', buried in the body of HIV PR, most likely affect the structural stability of the enzyme. The side chains of Asn88/88' form buried hydrogen bonds and replacement of this residue by aspartic acid or serine not only eliminates some of these bonds but also introduces unfavorable interactions in the core of HIV PR. Similarly, Leu90/90' is buried in a tight hydrophobic space close to the “fireman's grip” motif, which

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involves the catalytic asparates 25/25'. The structural effect of a mutation of Leu90/90' to the larger methionine is rather difficult to predict since it can either rigidify or destabilize the HIV PR dimer or it may have an effect on the catalytic efficiency of the “resistant” enzyme. The complicated pattern of HIV resistance to protease inhibitors, in particular the appearance of compensatory mutations that alone do not confer any resistance, suggests that the key to understanding the basis of decreased susceptibility of the virus to a given drug is the kinetics of specific processing of the GAG and GAG-POL polyproteins. The reduction in sensitivity of a mutant HIV PR towards any inhibitor can be conveniently reflected by the ratio of Ki mutant/Ki wild type. However, this reduced inhibitor sensitivity is only one component that distinguishes mutant-form from wild-type proteases. For virus encoding of a mutant HIV PR to be viable, the mutant protease must be capable of a minimal (although not yet quantified) level of enzymatic activity towards all substrates required for maturation of the virions. This proteolytic efficiency is reflected in the specificity constants (Kcat/Km) as determined for mutant and wild type HIV PRs. In order to rationalise these potentially conflicting relationships between enzymes, substrates, and inhibitors, Gulnik and his colleagues [66] introduced the term “Vitality Factor,” in which

In order for the “Vitality Factor” to be predictive for the level of resistance expected for a particular drug or combination of drugs for a given resistant strain of HIV, the determination of the specificity constants (Kcat/Km for mutants) must be repeated for all nine known substrates processed by HIV PR. The inhibition constants of a given compound should not depend on the substrate, but the Kcat/Km ratios do and therefore vitality values will differ for different substrates. It will be expected that the mean for all nine “vitality” values will be predictive for the change in antiviral activity for a particular compound. Although those data will be derived from in vitro experiments and are clearly not without some limitations, they may help in understanding the molecular basis of resistance and may contribute some value to possible multidrug strategies for the clinical management of AIDS. III. Perspective HIV PR inhibitors with acceptable oral availability and pharmacokinetic properties offer great promise for the treatment of HIV infection and AIDS. Efficacy studies of indinavir, ritonavir, or nelfinavir using plasma viral RNA as a marker have demonstrated up to three log reductions in RNA copy numbers that are

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sustained in many patients [67–69]. In contrast, nucleoside antiretroviral therapy that targets reverse transcriptase rarely results in more than one log reduction of viral RNA, indicating fairly poor inhibition of viral replication by this class of compounds. One of the reasons for the apparent greater in vivo antiviral activity of HIV PR inhibitors could be the mechanistic difference of the two enzymes and their respective activities in the viral life cycle. However, growing evidence of retroviral resistance to protease inhibitors remains a concern. The availability of several chemically distinct HIV PR inhibitors, including the second generation of compounds currently under preclinical development, offers a possibility of combining two or more drugs that share little cross-resistance. Also, it seems reasonable to evaluate these compounds in combination with various nucleoside and nonnucleoside reverse transcriptase inhibitors. Early clinical data from such combination therapy indicates reduction of retroviral RNA in plasma to levels lower than the currently available limit of detection [70]. This is the first indication that the application of well-chosen combination therapy can place AIDS patients in prolonged virologic and clinical remission. Undoubtedly, protein crystallography and other elements of structure-based drug design were widely applied in the discovery of HIV PR inhibitors. It will be prudent to assume that, in the absence of structural feedback, rapid discovery of several chemically different and potent inhibitors of HIV PR would have been severely impeded if not even impossible. However, structure-based drug design still remains a new and developing technology. Further success of this drug discovery technique largely depends on the development of methods of computational chemistry. Several computational approaches such as ALADDIN [71], DOCK [72], and MCDNLG [46] have been applied with a limited degree of success in a search for novel inhibitors of HIV PR and these methods will be developed further. The most difficult and challenging computational task required for full implementation of structure-based drug design involves assigning a priority to designed compounds before their synthesis, by computation of the absolute free energy of binding or by prediction of the relative difference in the binding constants of chemically related compounds. While the former approach is technically very difficult, due to the size of configurational space that must be sampled and the limited accuracy of the force field that describes atomic interactions in the molecular system [73], the latter approach has had some successes [74,75]. Nevertheless, owing to the various assumptions and approximations that underlie these techniques, such methods are useful only as order-of-magnitude estimates [74]. Further improvements of these methods heavily depend on the availability of structural and thermodynamic data for several closely related compounds that could be used to calculate parameters required for the implementation of such thermodynamic-integration cycles. The large number of high-resolution crystal structures of HIV PR

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complexed with various inhibitors offers a unique opportunity for the development of such computational methods if the structural data can be coupled with thermodynamic measurements of inhibitor-protein binding. These include direct measurements by microcalorimetry of the association constant, K, and in addition the enthalpy, entropy, heat capacity, and stoichiometry of binding. The combination of such thermodynamic and structural data will lead to a more precise understanding of the factors that influence binding and, ultimately, will lead to new general design principles that can be applied to drug discovery in the area of AIDS as well as other challenging diseases. Acknowledgments I wish to thank all my co-workers from Agouron who contributed to these studies, in particular J. Davies, S. Reich, M. Melnick, V. Kalish, A. Patick, L. Musick, and B-W. Wu. Steven Kaldor from Ely Lilly is acknowledged for his contribution in designing AG1343 (nelfinavir). I would like to thank Richard Ogden for critical reading of the manuscript and D. Olson for expert assistance in preparing the manuscript. References 1. Mitsuya H, Yarchoan R Broder S. Molecular Targets for AIDS therapy. Science 1990; 249:1533–1543. 2. DeClerq E. Toward improved anti-HIV chemotherapy: Therapeutic intervention with HIV infections. J. Med. Chem. 1995; 38:2491–2517. 3. Tomaselli AG, Howe JW, Sawyer TK, Wlodawer A, Henrikson RL. HIV-1 protease as a target for drug design. Chimica Oggi 1991; 9:6–14. 4. Ding J, Das K, Yadav PNS, Hsiou Y, Zhang W, Hughes SH, Arnold E. Structural studies of HIV-1 reverse transcriptase and implications for drug design. In: Structure-Based Drug Design. New York: Marcel Dekker 1996, in press. 5. Perno CF, Bergamini A, Pesce CD. Inhibition of the protease of human immunodeficiency virus blocks replication and infectivity of the virus in chronically infected macrophages. J. Infect. Dis. 1993; 168:1148–1156. 6. Kohl NE, Emini EA, Schleif WA, Davis LJ, Heimbach JC, Dixon RAF, Scolnick EM, Sigal, IS. Active human immunodeficiency virus protease is required forviral infectivity. Proc. Natl. Acad. Sci. USA, 1988; 85:4186–4690. 7. McQuade TK, Tomasselli AG, Liu L, Karacostas V, Moss B, Sawyer TK, Heinrikson RL, Tarpley WG. A synthetic HIV-1 protease inhibitor with antiviral activity arrests HIV-like particle maturation. Science 1990; 247:454–456.

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26. Swain AL, Miller M, Green J, Rich DH, Schneider J, Kent, SBH, Wlodawer A. X-ray crystallographic structure of a complex between a synthetic protease of Human Immunodeficiency Virus 1 and a substrate-based hydroxyethylamine inhibitor. Proc. Natl. Acad. Sci. USA. 1990; 87:8805–8809. 27. Krohn A, Redshaw S, Ritchie JC, Graves BJ, Hatada MH. Novel binding mode of highly potent HIV proteinase inhibitors incorporating the (R)-hydroxyethylamine isostere. J. Med. Chem. 1991; 34:3340–3342. 28. Erickson J, Neidhart DJ, VanDrie J, Kempf DJ, Wang XC, Norbeck DW, Plattner JJ, Rittenhouse JW, Turon M, Wideburg N, Kohlbrenner WE, Simmer R, Helfrich R, Paul DA, Knigge, M. Design, activity, and 2.8 Å crystal structure of a C2 symmetric inhibitor complexed to HIV-1 protease. Science. 1990; 249:527–533. 29. Hosur MV, Bhat TN, Kempf DJ, Baldwin ET, Liu B, Gulnik S, Wideburg NE, Norbeck DW, Appelt K, Erickson JW. Influence of stereochemistry on activity and binding modes for C2 symmetry-based diol inhibitors of HIV-1 protease. J. Amer. Chem. Soc. 1994; 116:847–855. 30. Kempf DJ, Marsh KC, Denissen JF, McDonald E, Vasavanonda S, Flentge CA, Green BE, Fino L, Park CH, Kong XP, Wideburg NE, Saldivar A, Ruiz L, Kati WM, Sham HL, Robins T, Stewart KD, Hsu A, Plattner JJ, Leonard JM, Norbeck DW. ABT-538 is a potent inhibitor of human immunodeficiency virus protease and has high oral availability in humans. Proc. Natl. Acad. Sci. USA. 1995; 92:2484–2488. 31. Kempf DJ, Marsh KC, Fino LC, Bryant P, Craig-Kennard A, Sham HL, Zhao C, Vasavanonda S, Kohlbrenner WE, Wideburg NE, Saldivar A, Green BE, Herrin T, Norbeck D.W. Design of orally available, symmetry-based inhibitors of HIV protease. Bioorg. Med. Chem. Lett. 1994; 2:847–858. 32. Holloway MK, Wai JM, Halgren TA, Fitzgerald PMD, Vacca JP, Dorsey BD, Levin RB, Thompson WJ, Chen LJ, deSolms SJ, Gaffin N, Ghosh AK, Giuliani EA, Graham SL, Guare JP, Hungate RW, Lyle TA, Sanders WM, Tucker TJ, Wiggins M, Wiscount CM, Woltersdorf OW, Young SD, Darke PL, Zugay JA. A priori prediction of activity fro HIV-1 protease inhibitors employing energy minimization in the active site. J. Med. Chem. 1995; 38:305–317. 33. Vacca JP, Guare JP, deSolms SJ, Sanders WM, Giuliani EA, Young SD, Darke PL, Zugay J, Sigal IS, Schleif W, Quintero J, Emini E, Anderson P, Huff JR. L-687,908, a potent hydroxyethylenecontaining HIV protease inhibitor. J. Med. Chem. 1991; 34:1225–1228. 34. Dorsey BD, Levin RB, McDaniel SL, Vacca JP, Guare JP, Darke PL, Zugay JA, Emini EA, Schleif WA, Quintero LC, Lin JH, Chen I-W, Holloway MK, Fitzgerald PMD, Axel MG, Ostovic D, Anderson PS, Huff JR. L-735,524: the design of a potent and orally available HIV protease inhibitor. J. Med. Chem. 1994; 37:3443–3451.

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37. Kaldor SW, Appelt K, Fritz JE, Hammond M, Crowell TA, Baxter AJ, Hatch SD, Wiskerchen MA, Muesing MA. A systemic study of P1–P3 spanning sidechains for the inhibition of HIV-1 protease. Bioorg. Med. Chem. Lett. 1995; 5:715–720. 38. Kalish JV, Tatlock JH, Davies JF, Kaldor SW, Dressman BA, Reich S, Pino M, Nyugen D, Appelt K, Musick L, Wu B-W. Structure-based drug design of nonpeptidic P2 substituents for HIV-1 protease inhibitors. Bioorg. Med. Chem. Lett. 1995; 5:727–732. 39. Kaldor SW, Kalish VJ, Davies JF, Appelt K, Tatlock JH, Dressman BA, Campanale KM, Burgess JA, Lubbehusen PL, Muesing MA, Hatch SD, Shetty BV, Patick AK, Kosa MB, Khalil DA. AG1343: a potent orally bioavailable inhibitor of HIV-1 protease. J. Med. Chem. 1996; submitted for publication. 40. Shetty B, Kosa M, Khalil DA, Webber S. Preclinical Pharmacokinetics and Distribution to Tissue of AG1343, an inhibitor of human deficiency virus type 1 protease. Antimicr. Ag. and Chemoth. 1996; 40:110–114. 41. Quart BD, Chapman SK, Peterkin J, Webber S, Oliver S. Phase 1 safety, tolerance, pharmacokinetics and food effect studies of AG1343—a novel HIV protease inhibitor. Abst. LB3. In: Proceedings of the 2nd National Conference on Human Retroviruses and Related Infections. 1995:163. 42. Lam PYS, Jadhaw PK, Eyermann CJ, Hodge CN, Lee YR, Bacheler LT, Meek JL, Otto MJ, Rayner MM, Wong YN, Chang C-H, Weber PC, Jackson DA, Sharpe TR, Erickson-Viitanen S. Rational design of potent, bioavailable, nonpeptide cyclic ureas as HIV protease inhibitors. Science, 1994; 263:380–384. 43. Reich SH, Melnick M, Davies JF, Appelt K, Lewis KK, Fuhry MA, Pino M, Trippe AJ, Nguyen D, Dawson H, Wu B-W, Musick L, Kosa M, Kahil D, Webber S, Gehlhaar DK, Andrada D, Shetty B. Protein structure-based design of potent, orally bioavailable, nonpeptide inhibitors of human immunodeficiency virus protease. Proc. Natl. Acad. Sci. 1995; 92:3298–3302. 44. Reich SH, Melnick M, Pino M, Fuhry MA, Trippe AJ, Appelt K, Davies JF, Wu B-W, Musick L. Structure-based design and synthesis of substituted 2-butanols as nonpeptidic inhibitors of HIV protease: secondary amide series. J. Med. Chem. 1996; 39:2781–2794. 45. Melnick M, Reich SH, Lewis KK, Mitchell L, Ngyen D, Trippe AJ, Dawson H, Davies JF, Appelt K, Wu B-W, Musick L, Gehlhaar DK, Webber S, Shetty B, Kosa M, Kahil D, Andrada D. Bis-tertiary amide inhibitors of the HIV-1 protease generated via protein structure-based iterative design. J. Med. Chem. 1996; 39:2795–2811. 46. Gehlhaar DK, Moerder KE, Zichi D, Sherman CJ, Ogden RC, Freer ST. De novo design of enzyme inhibitors by Monte Carlo ligand generation. J. Med. Chem. 1995; 38:466–472.

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Zipp GL, Dalga RJ, Schwende FJ, Howard GM, Padbury GE, Toth LN, Zhao Z, Koeplinger KA, Kakuk TJ, Cole SL, Zaya RM, Piper RC, Jeffrey P. Structure-based design of HIV protease inhibitors: 4-hydroxycoumarins and 4-hydroxy-2-pyrones as nonpeptidic inhibitors. J. Med. Chem. 1994; 37:3200–3204. 50. Vara Prasad JVN, Para KS, Lunney EA, Ortwine DF, Dunbar Jr. JB, Fergusson D, Tummino PJ, Hupe D, Tait BD, Domagala JM, Humblet C, Bhat TN, Liu B, Guerin DMA, Baldwin ET, Erickson JW, Sawyer TK. Novel series of achiral low molecular weight and potent HIV-1 protease inhibitors. J. Am. Chem. Soc. 1994; 116:6989–6990. 51. Ho DD, Neuman AU, Pereison AS, Chen W, Leonard JM, Markowitz M. Rapid turnover of plasma virions and CD4 lymphocytes in HIV-1 infection. Nature 1995; 373:123–126. 52. Wie X, Ghosh SK, Taylor ME, Johnson VA, Emini EA, Deutsch P, Lifson JD, Bonhoeffer S, Nowak MA, Hahn BH, Saag MS, Shaw GM. Viral dynamics in HIV-1 infection. Nature 1995; 373:117–122. 53. Coffin JM. HIV population dynamics in vivo: implications for genetic variation, pathogenesis and therapy. Science 1995; 267:483–489. 54. De Clercq E. HIV resistance to reverse transcriptase inhibitors. Biochem. Pharmacol. 1994; 47:155–169. 55. Mo H, Markowitz M, Ho DD. Patterns of specific mutations in HIV-1 protease that confer resistance to protease inhibitors in clinical development. Third International Workshop on HIV drug resistance. Kauai, Hawaii, August 1994. Abstract 13. 56. Markowitz M, Mo H, Kempf DJ, Norbeck D. Selection and analysis of human immunodeficiency virus type 1 variants with increased resistance ABT-538, a novel protease inhibitor. J. Virol 1995; 69:701–706. 57. Kaplan AH, Michael SF, Wehbie RS, Knige MF, Paul DA, Everitt L, Kempf DJ, Norbeck DW, Erickson JW, Swanstrom R. Selection of multiple human immunodeficiency virus type 1 variants that encode viral proteases with decreased sensitivity to an inhibitor of the viral protease. Proc. Natl. Acad. Sc. USA 1994; 91:5597–5601. 58. Erickson JW, Baldwin ET, Bhat TN, Gulnik S, Leu B, Yr B. Structural basis of drug resistance to HIV-1 protease inhibitors. Third International Workshop on HIV Drug Resistance. Kuaui, Hawaii, August 1994, Abstract 2. 59. Condra JH, Schleif WM, Blahy OH, Gabryelski LJ, Graham DJ, Quintero JC, Rhodes A, Robbins HL, Roth E, Shivaprakash M, Titus PL, Yang Y, Emini EA. Mutations in HIV protease conferring resistance to inhibitor L735,524. Abstract 187. In: Proceedings of the 2nd National Conference on Human Retroviruses and Related Infections. 1995:88.

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60. Condra JH, Schleif WA, Blahy OM, Gabryelski LJ, Graham DJ, Quintero JC, Rhodes A, Robbins HL, Roth E, Shivaprakash M. In vivo emergence of HIV-1 variants resistant to multiple protease inhibitors. Nature, 1995; 374:569–571. 61. Jackobsen H, Yasargil K, Winslow JC, Craig JC, Krohn A, Duncan IB, Mous J. Characterization of human immunodeficiency virus type 1 mutants with decreased sensitivity to proteinase inhibitor RO 31–8959. Virology, 1995; 206:527–534. 62. Patick AK, Mo H, Markowitz M, Appelt K, Wu B-W, Musick L, Kalish V, Kaldor SW, Reich SH, Ho D, Webber S. Antiviral and resistance studies of AG1343, an

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orally bioavailable inhibitor of human immunodeficiency virus protease. Antimicr. Ag. and Chemoth. 1996; 40:292–297. 63. Korant B, Lu Z, Strack P, Rizzo C. HIV protease mutations leading to reduced inhibitor susceptibility. In: Intracellular Protein Catabolism. New York: Plenum Press, 1996:241–250. 64. Patick AK, Duran M, Cao Y, Pei Z, Keller MR, Peterkin J, Chapman S, Anderson B, Markowitz M. Genotypic and phenotypic characterization of HIV-1 variants isolated from in vitro selection studies and from patients treated with the protease inhibitor, nelfinavir. Fifth International Workshop on HIV Drug Resistance, Whistler, Canada, 1996:29. 65. Jacobsen H, Brun-Vezinet F, Duncan I, Hanggi M, Ott M, Vella S, Weber J, Mous J. Genotypic characterization of HIV-1 from patients after prolonged treatment with protease inhibitor saquinavir. In: Abstracts of the 3rd International Workshop on HIV Drug Resistance. London: MediTech Media, 1994:16. 66. Gulnik SV, Suvorov LI, Liu B, Yu B, Anderson B, Mitsuya H, Erickson JW. Kinetic characterization and cross-resistance patters of HIV-1 protease mutants selected under in vitro drug pressure. Biochemistry 1995; 34:9282–9287. 67. Stein DS, Fish DG, Chodakewitz J. A 24-week open-label phase I evaluation of the HIV protease inhibitor L 735,524. Abstract LB1 Second National Conference on Human Retroviruses and Related Infections, Washington D.C., 1995. 68. Markowitz M, Jalil L, Hurley A. Evaluation of the antiviral activity of orally administered ABT-538, an inhibitor of HIV-1 protease. Abstract 185 Second National Conference on Human Retroviruses and Related Infections, Washington D.C., 1995. 69. Gathe Jr. J, Burkhardt B, Hawley P, Conant M, Peterkin J, Chapman S. A randomized Phase II study of Virocept™, a novel HIV protease inhibitor, used in combination with stavudine (D4T) vs. stavudine (D4T) alone. Abstract Mo.B.413 In: XI International Conference on AIDS, Vancouver, 1996:25. 70. Hammer S. Advances in antiretroviral therapy and viral load monitoring. Abstract Mo.01 In: Abstracts of the XI International Conference on AIDS, Vancouver, 1996:2. 71. VanDrie JH, Weininger D, Martin YC. Aladdin: an integrated tool for computer-assisted molecular design and pharmacophore recognition from geometric, steric, and substrate searching of threedimensional structures. Computer-Aided Mol. Design. 1989; 3:225–234. 72. Kuntz ID, Blanley JM, Oatley SJ, Langridge R, Ferrin TE. A geometry approach to macromoleculeligand interactions. J. Mol. Biol. 1982; 161:269–278. 73. Van Gunsteren WF Berendsen HJC. Computer simulation of molecular dynamics: methodology, application and perspectives in chemistry. Angew. Chem. Int. Ed. Eng. 1990; 29:992–996. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_39.html (1 of 2) [4/5/2004 4:47:09 PM]

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74. Reddy RM, Varney MD, Kalish V, Viswanadhan VN, Appelt K. Calculation of relative differences in the binding free energies of HIV-1 protease: a thermodynamic cycle perturbation approach. J. Med. Chem. 1994; 37:1145–1152. 75. Verkhivker G, Appelt K, Freer ST, Villafranca JE. Empirical free energy calculations of ligandprotein crystallographical complexes—knowledge-based ligand-protein interaction potentials applied to the prediction of human immunodeficiency virus protease binding affinity. Protein Engineering, 1995; 8:677–691.

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2 Structural Studies of HIV-1 Reverse Transcriptase and Implications for Drug Design Jianping Ding, Kalyan Das, Yu Hsiou, Wanyi Zhang, and Edward Arnold Center for Advanced Biotechnology and Medicine, and Rutgers University, Piscataway, New Jersey Prem N. S. Yadav University of Medicine and Dentistry of New Jersey, Piscataway, New Jersey Stephen H. Hughes ABL-Basic Research Program, NCI-Frederick Cancer Research and Development Center, Frederick, Maryland I. Introduction Like all other retroviruses, human immunodeficiency virus type 1 (HIV-1) contains the multifunctional enzyme reverse transcriptase (RT). Retroviral RTs have a DNA polymerase activity that can use either an RNA or a DNA template and an RNase H activity. HIV-1 RT is essential for the conversion of singlestranded viral RNA into a linear double-stranded DNA that is subsequently integrated into the host cell chromosomes [1–4]. In this conversion process HIV-1 RT catalyzes the incorporation of approximately 20,000 nucleotides. Chemotherapeutic agents have been identified that target virtually all stages of the HIV-1 replication cycle (see review [5]). Since both the polymerase and RNase H activities of HIV-1 RT are essential, inhibiting either step blocks viral replication. Therefore, HIV-1 RT is an important target for the treatment of AIDS. Two major classes of antiviral agents that inhibit HIV-1 RT polymerization have been identified; these are nucleoside RT inhibitors (NRTIs) (Figure la) and nonnucleoside RT inhibitors (NNRTIs) (Figure 1b). Nucleoside analogs, such as 3'-azido-2',3'dideoxythymidine (AZT), 2',3'-dideoxyinosine (ddI), 2',3'-dideoxycytidine (ddC), 2',3'-dideoxy-3'thiacyti-

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Figure 1 (a) Chemical structures of representative nucleoside analog inhibitors of HIV-1 RT. AZT: 3'-azido-2',3'-dideoxythymidine; d4T: 2',3'-didehydro-2',3'-dideoxythymidine; ddI: 2',3'-dideoxyinosine; ddC: 2',3'-dideoxycytidine; 3TC: 2',3'-dideoxy-3'-thiacytidine; PMEA: 9-(2-phosphonylmethoxylethyl)adenine.

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(b) Chemical structures of representative nonnucleoside inhibitors of HIV-1 RT. Nevirapine: 11 -cyclopropyl-5,11 -dihydro-4-methyl-6H-dipyrido (3,2-b:2'3'-e)(1,4) diazepin-6-one; α-APA: αanilinophenylacetamine; TIBO: tetrahydroimidazo-(4,5,1-jk) (1,4)-benzo-diazepin-2(1H)-one and thione; pyridinones; HEPT: 1-{(2-hydroxyethoxy) methyl}-6-(phenylthio)thymine; BHAP: bis(heteroaryl)piperazine; TSAO: {2',5'-bis-O-(tert-butyldimethylsilyl)}-3' -spiro-5''-(4''-amino-1",2"-oxathiole)-2", 2"-dioxide; L-737,126: 5-chloro3-(phenylsulfonyl)indole-2-carboxamide; TBA: 1-(2',6'-difluoro-phenyl)-1H,3H-thiozolo -(3,4-a) -benzimidazole; quinoxaline S-2720: 6-chloro-3,3-dimethyl-4-(isopropenyloxycarbonyl) -3-4-dihydroquinoxalin-2(1H)-thione.

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dine (3TC), and 2',3'-didehydro-2',3'-dideoxythymidine (d4T), have been widely used in the treatment of HIV-1 infections [5–7]. However, the effectiveness of these drugs is limited by their cytotoxicity and the rapid emergence of drug-resistant viral strains [5,8–12]. Nonnucleoside inhibitors, e.g., the HEPT derivatives [13], TIBO derivatives [14], nevirapine [15], pyridinones [16], BHAP derivatives [17], TBA derivatives [18,19], TSAO derivatives [20], α-APA [21], and quinoxalines (HBY) [22,23], are potent inhibitors of HIV-1 RT (see reviews [5,11,12]). While these inhibitors differ considerably in chemical structure, all of them are quite specific for HIV-1 RT and inhibit neither HIV-2 RT nor variety of cellular polymerases. Challenging a virus with these drugs

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Page 45 Figure Continued

rapidly selects viral strains containing drug-resistance mutations [9,24,25]. HIV-1 viral variants are known whose RT is resistant to all of the currently available drugs/inhibitors (see reviews [5,9,11,12,26,27]). In some cases, drug-resistant variants can be selected in very short periods of time [9], a consequence of the high viral load and rapid turnover of viral populations in infected individuals [28–30]. A better understanding of how these viral variants confer resistance should provide insight into the limitations of their genetic flexibility. In the past few years, substantial progress has been made in understanding the three-dimensional structure of HIV-1 RT. This paper will discuss the recent biochemical, genetic, and clinical data of HIV-1 RT in the context of the crystal structure of HIV-1 RT and prospects for development of more effective inhibitors of HIV-1 replication. II. Three-Dimensional Structures of HIV-1 RT Three-dimensional crystal structures of HIV-1 RT have been determined both for the unliganded form of the protein and for complexes with either template-primer substrate or nonnucleoside inhibitors (Figure 2 and Table 1). Structures of HIV-1 RT have been determined in complexes with a series of NNRTIs, including nevirapine [31–33], 1051U91 (a nevirapine analog) [33], α-APA R95845 [34], α-APA R90385 [33], HEPT [33], 8-Cl TIBO (R86183) [35], and 9-Cl TIBO (R82913) [36,37]. The structure of HIV-1 RT in a ternary complex with a 19-mer/18-mer double-stranded DNA (dsDNA) template-primer and an antibody Fab fragment has been described [38]. In addition, structures of unliganded HIV-1 RT have also been solved in multiple crystal forms [39–43]. The structure of a polypeptide corresponding to the fingers and palm subdomains of the HIV-1 RT polymerase domain has also been determined [44]. HIV-1 RT is an asymmetric heterodimer consisting of the p66 (66 kDa) and p51 (51 kDa) subunits. The N-terminal 440 residues of the p66 subunit constitute the polymerase domain and the C-terminal 120 residues of p66 form the RNase H domain; the p51 subunit has the same amino acid sequence as the polymerase domain of the p66 subunit [1,2]. The polymerase domain of the p66 subunit has been likened to a human right hand. On this basis the subdomains of both p66 and p51 have been designated as fingers, palm, thumb, and connection (Figure 2) [31,38]. In the p66 subunit, the fingers, palm, and thumb subdomains form a large cleft that can accommodate the DNA substrate. The polymerase active site, which contains three strictly conserved aspartic acid residues (Asp110, Asp185, and Asp186), is located in the DNA-binding cleft and is part of the p66 palm subdomain (Figure 3) [31,38]. In the p51 subunit, however, the thumb is rotated away from the fingers and the connection subdomain is folded over onto the palm subdomain between the fingers and thumb subdomains. As a

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Page 46 Table 1 Crystal Structures of HIV-1 Reverse Transcriptase

PDB entry

HIV-1 RT Structure

Resolution Range (Å)

R-Factor/Free R-Factor (%)

Data Completeness (%)

Temperature (°C)

References

85

-165

41

21.9

89.5

-173

42

1hmv

unliganded RT

6.0-3.0

25.4/29.7

1rtj

unliganded RT

25.0-2.35

1dlo

unliganded RT

8.0-2.7

23.0/33.6

99.5

-165

43

1hmi

RT/DNA/Fab

15.0-3.0

26.0

88.1

-10

38

3hvt

RT/nevirapine

8.0-2.9

26.6

95.6

-165

31,32

1vrt

RT/nevirapine

25.0-2.2

18.6

87.1

16

33

1rth

RT/1051U91

25.0-2.2

21.4

81.4

16

33

1hni

RT/α-APA (R95845)

10.0-2.8

25.5/36

78.5

-15

34

1vru

RT/α-APA (R90385)

25.0-2.4

18.7

86.5

16

33

1hnv

RT/8-Cl TIBO (R86183)

10.0-3.0

24.9/35.6

81

-10

35

1rev

RT/9-Cl TIBO (R82913)

25.0-2.6

22.4

80.7

-173

36

1tvr

RT/9-Cl TIBO (R82913)

10.0-3.0

25.9

72

-165

37

1rti

RT/HEPT

25.0-3.0

23.6

86.3

14

33

1hrh

RT (RNase H domain)

20.0-2.4

20.0

93.1

4

94

1rdh

RT (RNase H domain)

20.0-2.8

21.5

95.6

4

95

1har

RT (fingers and palm subdomains)

7.0-2.2

20.8/27.0

96

4

44

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Figure 2 Ribbon diagrams of the three types of HIV-1 RT crystal structures. (a) Structure of the HIV-1 RT/DNA/Fab ternary complex [38]. The bound nucleic acid is shown as double-stranded helices with template strand in black and primer strand in gray. The Fab fragment is not shown. (b) Structure of the HIV-1 RT complexed with the NNRTI TIBO R86183 [34]. For the sake of clarity, the bound TIBO inhibitor is shown as an atomic model. (c) Structure of unliganded HIV-1 RT [41,43]. (d) A schematic diagram showing the resemblance of the HIV-1 RT p66 subunit to a human right hand. The RNase H domain, which has no counterpart for a human hand, is shown as an oval below the thumb. When a template-primer binds to HIV-1 RT, the fingers, palm, and thumb subdomains of p66 form a large cleft to bind the DNA. The polymerase active site (shown as a small circle) lies at the bottom of the DNA-binding cleft. The NNRTI binds in the highly hydrophobic NNIBP (shown as a large circle), which is located in the vicinity of the polymerase active site. The p66 thumb subdomain in the NNRTI-bound HIV-1 RT structures is in an upright position extended beyond that observed in the structure of RT with bound DNA. In the absence of any bound nucleic acid or NNRTI, however, the p66 thumb folds down into the DNA-binding cleft (shown as dashed drawing).

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Figure Continued

consequence, the p51 submit has no cleft for binding nucleic acid substrates and hence no polymerase activity. There is considerable evidence showing that HIV-1 RT is quite flexible and that this flexibility is essential for DNA polymerization. Comparisons among DNA-bound, inhibitor-bound, and unliganded HIV-1 RT structures provide a demonstration of the enzyme's flexibility. When a DNA template-primer binds to HIV-1 RT, structural elements of the fingers, palm, and thumb subdomains of the p66 subunit form a clamp-like structure that holds the nucleic acid (Figure 2) [38]. The template-primer substrate interacts with amino acid residues of the fingers, palm, and thumb subdomains, especially in the regions denoted as “primer grip” and “template grip,” believed to position the template-primer precisely relative to the polymerase active site [38]. The primary contacts between the template-primer and the protein are along the sugar-phosphate backbone of the DNA and thus are not sequence-specific [38]. In the absence of nucleic acid template-primer or NNRTI, the thumb subdomain of p66 is folded down into the DNAbinding cleft and lies near the fingers subdomain (Figure 2) [40,41,43]. As a consequence, the DNAbinding cleft is closed. However, even in the absence of a bound nucleic acid, binding of an NNRTI induces both short-range and long-range structure distortions, including a hinge-like movement near the base of the p66 thumb that constrains the p66 thumb in a conformation that is extended beyond the upright

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Figure 3 Overall structure of HIV-1 RT p66/p51 heterodimer [38] showing the locations of the major target sites for anti-HIV-1 RT inhibitors. The NRTIs target the dNTP-binding site/the polymerase active site (shown as a small striped circle) that lies at the floor of the DNA-binding cleft. The NNRTIs bind to the NNIBP (shown as a large dotted circle), which is near, but distinct from, the polymerase active site. The RNase H domain is located at the C-terminal of the p66 subunit. The RNase H catalytic site (shown as a medium circle) is an attractive target site for anti-HIV-1 drugs. The HIV-1 RT p66/p51 heterodimer interface is shown as a dashed line. Since HIV-1 RT functions as a heterodimer, any inhibitors that could interfere with the dimerization process might also be potential drugs for treating HIV-1 infection.

position of the thumb observed in the HIV-1 RT/DNA/Fab structure [31,33–37,43]. In the unliganded structure of HIV-1 RT reported by Esnouf et al. [42], the p66 thumb subdomain is in an upright conformation different from that observed in the other unliganded HIV-1 RT structures but similar to that found in the DNA-bound HIV-1 RT and NNRTI-bound HIV-1 RT structures. Esnouf et al. [42] contend that the upright conformation of the p66 thumb subdomain in their unliganded RT structure is appropriate for unliganded HIV-1 RT and that the binding of an NNRTI does not affect the conformation of the p66 thumb. However, we believe that the conformation of the p66 thumb in the Esnouf et al. structure may well be a result of the method used to prepare the

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crystals, which were produced by soaking out a weakly bound NNRTI (HEPT) from pregrown crystals. When the crystals were grown in the presence of HEPT, the p66 thumb was presumably in an upright position similar to that seen in all of the known NNRTI-bound HIV-1 RT structures. As the weakly bound HEPT diffused out, the molecular packing arrangement may have constrained the position of the p66 thumb subdomain. As a consequence, this unliganded HIV-1 RT structure may represent an intermediate between the other unliganded structures and the structure of HIV-1 RT with a bound NNRTI. It is evident that the p66 thumb subdomain has considerable flexibility and can adopt substantially different conformations during the binding of template-primer or inhibitors and, presumably during DNA polymerization as well. III. Polymerase Active Site of HIV-1 RT and the NRTIs Polymerization of DNA by HIV-1 RT involves a sequential stepwise binding of the template-primer and deoxynucleoside triphosphate (dNTP) substrates at the polymerase active site [45,46]. The incoming dNTP is covalently linked via the α-phosphorus to the 3'-oxygen of the primer strand, accompanied by the release of pyrophosphate. An essential requirement for the polymerization reaction is the presence of a 3'-OH group at the end of the primer strand. Nucleoside analogs contain a modified sugar moiety in which the 3'-OH group is replaced by another group (e.g., hydrogen, halogen, or azido) (Figure la). To exert their antiviral activity at the level of RT, the NRTIs must be phosphorylated successively to the 5'monophosphate, 5'-diphosphate, and 5'-triphosphate forms by a series of kinases. Once the NRTI is converted to the triphosphate form and interacts with RT, it can inhibit polymerization in two possible ways. One possibility is that the NRTI binds preferentially to the dNTP-binding site and competitively inhibits the binding of natural dNTPs. Another possibility, which seems to be the predominant mode of inhibition, is that the NRTI is incorporated into the growing chain and acts as a terminator of chain elongation. Once an NRTI is incorporated, no additional nucleotides can be added to the DNA chain since the primer terminal 3'-OH group (the site of phosphodiester bond formation) is absent. Phosphorylation is a crucial step in the intracellular metabolism of the NRTI and often is a limiting factor for the antiviral activity of the NRTI [47]. In an attempt to bypass the first phosphorylation step, several acyclic nucleoside phosphonates have been developed in which the sugar moiety of normal NRTIs is replaced with an acyclic phosphonate group, such as 9-(2-phosphonylmethoxyethyl)adenine (PMEA), (R)-9-(2-phosphonylmethoxypropyl)adenine (PMPA), and (S)-9-(3-fluoro-2phosphonylmethoxyethyl) adenine (FPMPA) (Figure la) (see reviews [5,11]). These acyclic nucleoside phosphonates are dideoxynucleoside

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monophosphate analogs which can easily be converted to the active triphosphate form by adding two additional phosphates [48,49], and can inhibit HIV replication [50,52]. Analysis of the structure of the HIV-1 RT/DNA/Fab complex showed that the dNTP-binding site is composed of both protein and nucleic acid. In addition to the 5'-terminus of the template nucleotide and three carboxylate residues (Asp110, Asp185, and Asp186), the amino acid residues Asp113, Tyr115, Phe116, Gln151, Phe160, and possibly Met184 and Lys219 form part of the putative dNTP-binding site [12,53] (Figure 4 and Table 2). It is important to realize that the precise composition, position, and conformation of the template-primer can influence the recognition and incorporation of incoming nucleotides at the polymerase catalytic site. In the wild-type HIV-1 RT, the dNTP-binding

Figure 4 Stereoview of the polymerase active site of HIV-1 RT [38]. The amino acid residues that compose the putative dNTP-binding site, including the three catalytically essential aspartic acids, are shown with side chains. The double-stranded nucleic acid is shown with the atomic model in the HIV-1 RT/DNA/Fab complex. The dNTP-binding site consists of structural elements from both protein and nucleic acid. The precise composition, position, and conformation of the template-primer can affect the recognition of incoming dNTPs at the polymerase active site.

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Page 52 Table 2 HIV-1 RT Amino Acid Residues Composing the Putative dNTP-Binding Sites and the Locations of NRTIResistance Mutations (see also [12]) dNTP-Binding Site

Nucleoside Drug-Resistance Mutation Site

Residue

Location

Mutation

Location

Possible Effects

Asp110

β6

Met41Leu

αA

template binding

55

Asp113

β6-αC loop

Ile50Thr

β2

unclear

154

Try115

αC

Lys65Arg

β3–β4

template binding

154

Phe116

αC

Asp67Asn

β3–β4

template binding

155

Gln151

β8-αE

Thr69Asp

β3–β4

template binding

156

Phe160

αE

Lys70Arg

β3–β4

template binding

155

Asp185

β9–β10

Leu74Val

β4

template binding

8

Asp186

β9–β10

Val75Thr

β4

template binding

163

Lys219

β11

Glu89Gly

β5

dsDNA binding

157

Tyr115Phe

αC

dNTP binding

170

Ile135Thr

β7–β8

unclear

Gln151Met

β8-αE

dNTP binding

Met184Val, Ile

β9–β10

dNTP-binding/fidelity

Thr215Tyr, Phe, Cys

β11

indirect effect/dNTP binding

Lys219Gln

β11

dNTP binding

References

158,159

154,160,161 155,162

155

site can accommodate both the normal dNTP substrates and dideoxynucleoside analogs. The majority of mutations that confer resistance to NRTIs are not located at the dNTP-binding site; however, they appear to influence the geometry of the dNTP-binding site indirectly in a way that permits RT to discriminate between a normal dNTP and a modified nucleoside triphosphate (see discussion in the next section).

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Analysis of various HIV-1 RT structures has revealed an unusual β-turn geometry for the YMDD motif at the polymerase catalytic site of p66 [33,34,41–43]. The energetically unfavorable main chain conformation of Met184 (torsion angles φ~60° and ϕ~-120°) is stabilized by a hydrogen bond of its carbonyl oxygen to the side chain of either Gln182 [33,41,43] or Gln161 [42]. It has been suggested that this novel β-turn geometry might be required to position the aspartic acids in precisely the correct way for catalysis [41]. IV. Mechanism of NRTI-Resistance Mutations Development of resistance to NRTIs has been a major problem with clinical use of these drugs. Careful analysis of mutations that confer resistance to different

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NRTIs in light of available structural data might provide information that could be used in the development of improved NRTIs that are more effective against the commonly observed NRTI-resistant HIV-1 variants. A viable drug-resistant RT mutant should be able to recognize and incorporate normal nucleoside triphosphate, yet reject a nucleoside analog. The only difference between normal nucleotide substrates and the NRTIs is the modification of the sugar moiety. This alteration may affect sugar puckering and the conformation of the glycosyl bond. Recognition of these differences could render the triphosphate form of the nucleoside analog a good substrate for wild-type RT but a poor substrate for a drug-resistant variant of RT. Structural analysis of HIV-1 RT has shown that most of the NRTI-resistance mutations are not located close to the putative dNTP-binding site and are unlikely to have a direct impact on the binding of dNTP analogs (Figure 5 and

Figure 5 A close-up view showing the relative locations of the commonly identified drug-resistance mutations for NRTIs (in dark-gray) and for NNRTIs (in light-gray) with respect to the bound DNA. Most of the NRTI-resistance mutations are not located at the putative dNTP-binding site, but are at positions to have potential interactions with the nucleic acid template-primer. Conversely, all the NNRTI-resistance mutations are clustered around the NNIBP and have direct contacts with NNRTIs or have direct effect on inhibitor binding.

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Table 2) [12,54]. For example, none of the five mutations, in HIV-1 RT Met41Leu, Asp67Asn, Lys70Arg, Thr215Tyr, and Lys219Gln [55] (Table 2) consistently associated with resistance to AZT, are at locations close to the dNTP-binding site. However, most (but not all, c.f. [56]) biochemical studies have failed to show that recombinant HIV-1 RT enzymes containing these mutations are more resistant to inhibition by AZT triphosphate than the wild-type HIV-1 RT [27,57,58]. Other mutations that confer resistance to NRTIs have been identified at positions 50, 65, 69, 74, 75, 89, 115, 135, 151, and 184 of HIV-1 RT (Figure 5 and Table 2). Most of these mutations do not lie close to the dNTP-binding site (Met184Val/Ile are the exception), but instead are located at positions where they could interact with the nucleic acid template-primer [12,59,60]. Biochemical data have shown that only when the 5'-template extension length is greater than three nucleotides does the wild-type RT begin to incorporate dideoxynucleotides effectively [54]. If the template extension is less than three nucleotides in length, wild-type HIV-1 RT is resistant to dideoxynucleotides. On the other hand, HIV-1 RT variants containing the mutations Leu74Val or Glu89Gly did not readily incorporate dideoxynucleotides either with short or long template extensions [54]. Based on both structural and biochemical data, it was proposed that mutations that cause HIV-1 RT to have reduced sensitivity to NRTIs exert their effects via interactions with the nucleic acid template-primer, which consequently alter the geometry of the polymerase active site [54]. It has been suggested that mutations that confer resistance to foscarnet might use a similar mechanism [61]. One possible exception to this mechanism might be the mutations of Met184Val and Met184Ile (see review [27]). Part of the highly conserved YMDD motif, Met184 is adjacent to residues Asp185 and Asp186, which are two of the three catalytically essential aspartic acid residues at the polymerase active site. In addition, Met184 appears to interact with the ribose moiety of the 3'-terminal nucleotide of the primer strand [12,38,53] (Figure 4). Therefore, mutations at this position could affect interactions with the incoming dNTP directly and/or alter the positioning of the nucleic acid. These mechanisms are not mutually exclusive and which mechanism is responsible for resistance has not yet been resolved [62]. There are two recent reports suggesting that the Met184Val mutant HIV-1 RT has approximately three-fold higher fidelity than the wild-type enzyme [63,64]. Based on these data, it was suggested that this increase in fidelity might reduce the overall rate of generation of viral variants in patients treated with 3TC or other dideoxynucleosides [64]. However, owing to both theoretical and technical problems with these analyses, these conclusions are controversial. Determination of crystal structures of both wild-type and mutant HIV-1 RT complexed with individual NRTIs in the presence of a variety of template-primers and/or dNTP substrates should provide a better understanding of the mechanisms of dNTP selection and drug resistance.ed at Arial for catalysis [41]. IV. Mechanism of NRTI-Resistance Mutations Development of resistance to NRTIs has been

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V. Drug Design Targeting at the Polymerase Active Site All of the existing NRTIs contain a modified sugar moiety that lacks the 3'-OH group that is essential for incorporation of the next nucleotide. Modification can also be made on other functional groups such as the base and the triphosphate moieties. It may be worthwhile to try to alter the base moiety of the NRTIs to produce compounds that will be more specific to HIV-1 RT (i.e., less cytotoxic to normal cellular polymerases) and more effective against both wild-type or drug-resistant viral variants. Structure-activity analysis indicates that the pyrimidine moiety of the NRTIs can be modified at the C5 position. An AZT derivative that has a 3'-azido group on the sugar moiety and a methyl group at the C5 position of the pyrimidine moiety showed potent antiviral activity [65]. Substitution of the methyl group with a chlorine atom at position C5 of AZT results in a compound that has strong anti-HIV-1 activity [66]. Other possible substitutions at the C5 position include other halogen atoms or an ethyl group. Another possible drug-design strategy would be to devise compounds that can interface with the binding of the metal ions (Mg2+ or Mn2+) at the polymerase active site. Metal ions appear to be important in DNA polymerase catalysis. Based on the structural and biochemical data, a two-metal dependent mechanism of polymerization has been postulated [53,67,68] that is similar to that proposed for other DNA polymerases [69–71]. In this model, the metal ions mediate interactions between the three catalytically essential aspartic acid residues (Asp100, Asp185, and Asp186) and the α-, β-, and γphosphates of the incoming dNTP and promote the nucleophilic attack on the α-phosphate by the oxygen atom of the 3'-OH group of the primer strand. In the structure of the fingers and palm subdomains of the RT of Moloney murine leukemia virus (MuLV), a single Mn2+ ion was found bound to the two aspartic acid residues at the polymerase active site [72]. In the structure of the unliganded HIV-1 RT, an electron density peak was located at the polymerase active site with a good coordination geometry to the Oδ1 atoms of both Asp185 and Asp186 [43]. This electron-density peak is in a position similar to that of the Mn2+ ion observed in the MuLV RT structure. It is possible that this position corresponds to a Mg2+ ion-binding site [43]. It might be useful to design inhibitors that would influence the metal-ion coordination using either computer-based calculations (such as DOCK [73–75]) or based directly on an analysis of HIV-1 RT structure. Crystal structures of HIV-1 RT complexed with Mg2+/Mn2+ ion(s) at the polymerase catalytic site in the presence of template-primer and/or dNTP substrates would be helpful in defining the target sites of inhibitors of this type. Further studies on the structure activity relationship of HIV-1 RT complexes with these inhibitors, if active, might ultimately lead to a new type of HIV-1 RT drug that would not compete with the dNTP binding but would affect the DNA polymerization mechanism.

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It is also attractive to consider developing agents that bind to the HIV-1 RT polymerase active site but are not nucleoside analogs. Since the amino acid residues at the dNTP-binding site are highly conserved, viral variants resistant to such inhibitors may be significantly impaired in their polymerase activity. However, there is a good chance that drug resistance could result from mutations in HIV-1 RT that influence the precise positioning of template-primer [54]. Initial attempts to use this approach starting from the crystal structure of the HIV-1 RT/DNA/Fab complex [38] (Ding, et al., in preparation) have uncovered some interesting lead compounds (Kuntz, Kenyon, Arnold, Hughes, et al., unpublished). VI. NNRTIs and the NNIBP Nonnucleoside RT inhibitors (NNRTIs) constitute the other major class of HIV-1 RT inhibitors. Many structurally distinct families of NNRTIs have been identified, including HEPT [13], TIBO [14], nevirapine [15], BHAP [17], TBA [18,19], TSAO [76], α-APA [21], pyridinones [16] and quinoxalines (HBY) [22,23] (Figure 1b). However, development of drug resistance is a major problem when NNRTIs are used to treat AIDS patients. An ideal drug should be able to block replication of all viable strains of HIV-1, but should not inhibit normal cellular enzymes. In this regard, the known NNRTIs may be too specific. While these inhibitors do not inhibit cellular polymerases, they are also inactive against HIV-2 RT (which can be viewed as an extreme variant of HIV-1 RT). In addition, drug-resistant variants of HIV-1 RT emerge rapidly in the presence of most inhibitors. In contrast, the NRTIs inhibit a broad spectrum of polymerases including the host cellular polymerases. Though it appears to be more difficult for the virus to evade NRTIs than NNRTIs (in general, it takes longer for the virus to develop resistance to NRTIs than NNRTIs), NRTI toxicity is a serious problem. Structural and biochemical studies have shown that all NNRTIs bind in a highly hydrophobic pocket in the p66 subunit, located approximately 10 Å away from the polymerase active site (Figures 2 and 3) [31,33–37]. Nevertheless, in all known structures of HIV-1 RT/NNRTI complexes, the bound NNRTIs have not been found to have any direct interactions with residues that compose the putative dNTPbinding site. The nonnucleoside inhibitor binding pocket (NNIBP) contains primarily amino acid residues from the β5–β6 loop (Pro95, Leu100, Lys101, and Lys103), β6 (Val106 and Val108), the β9–β10 hairpin (Val179, Tyr181, Tyr188, and Gly190), and the β12–β13 hairpin (Phe227, Trp229, Leu234, His235, and Pro236) of the p66 palm subdomain, and β15 (Tyr318) of the p66 thumb subdomain, as well as the β7–β8 connecting loop (Glu138) of the p51 fingers subdomain (Figure 6 and Table 3).

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Figure 6 Superposition of the NNRTIs in the HIV-1 RT/TIBO complex [35], HIV-1 RT/α-APA complex [34], and HIV-1 RT/nevirapine complex [32]. The side chains are shown for those amino acid residues that have close contacts with bound inhibitors and the three catalytically essential aspartic acids in the HIV-1 RT/TIBO complex. Most of the amino acid residues that form the NNIBP are hydrophobic. Though the NNRTIs are chemically and structurally diverse, the bound NNRTIs all assume a strikingly common butterfly-like shape.

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Location

Mutation

Possible Effects

References

Pro95

β5

Ala98

β5–β6

Gly

less bulky

9,164

Leu100

β5–β6

Ile

β-branch

9,164-166

Lys101

β5–β6

Glu

charge change

Lys103

β5–β6

Asn, Gln

charge loss, less bulky

9,24,164

Val106

β6

Ala

less bulky

9,58,165

Val108

β6

Ile

bulkier

Glu138

β7–β8(p51)

Lys

charge change

166

Val179

β9

Glu, Asp

charge gain, bulkier

164

Tyr181

β9

Cys, Ile

aromaticity loss, less bulky

24,58,164

Tyr188

β10

His, Cys, Leu

aromaticity loss, less bulky

9,167

Gly190

β10

Glu

charge gain, bulkier

Phe227

β12

Leu228

β12

Phe

aromaticity gain, bulkier

133

Trp229

β12

Glu233

β13

Val

charge loss, less bulky

169

Leu234

β13

Pro236

β13–β14

Leu

increase flexibility, bulkier

133

Lys238

β14

Thr

charge loss, less bulky

133

Tyr318

β15

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164

9,164

9,22,168

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Most of the amino acid residues that form the binding pocket are hydrophobic and five of them are aromatic residues. The hydrophobic interactions of the side chains of these residues, especially Tyr181, Tyr188, and Trp229, with the hydrophobic moieties of the NNRTIs appear to be important for inhibitor binding [32–34,59]. Since most of the NNRTIs also contain polar group(s), they have the potential to form hydrogen bonds with surrounding amino acid residues either directly or via water bridges [33–36]. In the structures of both liganded HIV-1 RT and the HIV-1 RT/DNA/Fab complex, there are two small surface depressions at the base of the NNIBP that are the putative entrances to the pocket [34,43]. One surface depression is located at the p66/p51 heterodimer interface and is composed of amino acid residues Leu100, Lys101, Lys103, Val179, Tyr181, and Tyr188 of p66, and Glu138 of p51 [34]. This putative entrance is narrow compared to the size of the NNRTIs. Another surface depression has been found at the location near the base of the p66 thumb subdomain between two adjacent structural elements: the β5–β6 connecting loop (Lys101 and Lys103) and the β13–β14 hairpin (Pro236 and Leu238) [43]. Since this site is also exposed to solvent, an NNRTI could approach the NNIBP from it. How-

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ever, once an NNRTI is bound to RT, only the first putative entrance remains accessible; the second disappears due to the conformational change and repositioning of the β12-β13-β14 sheet [43]. It is evident, from comparison of the various HIV-1 RT structures that the NNIBP has a highly flexible structure that apparently allows the enzyme to accommodate various types of NNRTIs with different shapes and sizes. Despite apparent differences in the structures of the bound inhibitors, comparison of structures of several HIV-1 RT/NNRTI complexes revealed remarkable similarity in the geometry of both the bound inhibitors and the NNIBP [33,35]. All these chemically diverse NNRTIs assume a strikingly similar butterfly-like shape (Figure 6). The binding of NNRTIs in the NNIBP can be likened to a butterfly sitting on the β6-β10-β9 sheet and facing toward the putative entrance to the pocket. The angle between the two wings of the “butterfly” is approximately 112–115° in the TIBO, α-APA, and nevirapine complexes [35]. This angle might be critical in inhibitor binding and could be a crucial parameter in the design of new NNRTIs. There are many other NNRTIs that are significantly larger or smaller in size than α-APA, TIBO, or nevirapine. It is very likely that the NNIBP can adopt other conformations. For example, BHAP appears to be too large to fit into the NNIBP in any of the reported HIV-1 RT/NNRTI complexes. The NNIBP in the HIV-1 RT/BHAP complex would need to be significantly larger than that observed in the structures of the known HIV-1 RT/NNRTI complexes. It is possible that the BHAP inhibitor may not conform to a butterfly-like shape. This underscores the importance of solving crystal structures for as many HIV-1 RT/NNRTI complexes as possible. Additional structural and biochemical data for other HIV-1 RT/NNRTI complexes should provide the insight needed to define the limits of the flexibility of HIV-1 RT in the NNIBP region. VII. Process of NNRTI Binding In crystal structures of unliganded HIV-1 RT [40,41,43] and of HIV-1 RT/DNA/Fab complex [38], the NNIBP does not exist (although a small cavity is found in the region of the NNIBP proximal to the polymerase active site in the unliganded HIV-1 RT structure described by Esnouf et al. [42]). In these structures, the side chains of Tyr181 and Tyr188 in p66 point away from the polymerase active site and toward the hydrophobic core. However, in the HIV-1 RT/NNRTI complex structures, the side chains of Tyr181 and Tyr188 point toward the polymerase active site, and the side chain of Tyr181 is in a position that prevents Trp229 from occupying the position it has in the unliganded or DNA bound HIV-1 RT structures. Binding an NNRTI also moves the β12-β13-β14 sheet away from the hydrophobic core [34,35,37]. These conformational

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changes create the space in the pocket required to accommodate inhibitors. In other words, significant conformational changes occur during the process of inhibitor binding that lead to the formation of the NNIBP [33–35]. This observation also underscores the importance of determining structures of HIV-1 RT with and without bound inhibitors. An obvious question is, what forces initiate this series of conformational changes during NNRTI binding? One possibility is the contacts between the inhibitor and the protein. Though the NNIBP is hydrophobic, there are three hydrophilic amino acid residues (Lys101 and Lys103 of p66, and Glu138 of p51) at the rim of the putative entrance(s) to the pocket. The flexible and polar side chains of these residues could assist in steering an inhibitor into the pocket and/or could block the bound inhibitor from escaping out of the pocket. Mutagenesis studies have shown that these three residues are important in the binding of NNRTIs. Though the importance could be explained in terms of the interactions between these residues and the bound inhibitor in the final complexes, interactions at the initial stages of inhibitor binding might also be crucial. The flexible and polar side chains of these residues might help in directing the inhibitor toward the entrance to the pocket via electrostatic interactions, in part by replacing the original hydrogen bonds between the drug and the solvent molecules. Any initial energy gains from such polar interactions could potentially be replaced by hydrogen bonds or other types of interactions between the inhibitor and alternative residues as the inhibitor moves deeper into the binding pocket. In addition, significant portions of the aromatic rings of both Tyr181 and Tyr188 are exposed at the bottom of the surface depression and offer the potential for early π-π interactions with the inhibitor. This type of π-π interaction might also play an important role in the initial approach of inhibitors to the binding pocket. This hypothesis may provide a kinetic explanation for the ineffectiveness of NNRTIs against viral strains of HIV-1 that carry nonaromatic amino acids at positions 181 and 188. As the solvated inhibitor approaches the enzyme and proceeds to enter the binding pocket, most of the water molecules of solvation are lost. The few water molecules that remain in the NNRTI-bound complex are typically located at the entrance to the pocket, forming water bridges between the inhibitor and one or two polar residues around the entrance [33,35,36]. Once the inhibitor is in place, the surface residues close down around the drug preventing it from escaping by effectively sealing the entrance to the pocket. VIII. Mechanisms of Inhibition by NNRTIS Based on structural, biochemical, and genetic data several hypotheses have been postulated about the mechanism(s) of inhibition of HIV-1 RT by NNRTIs. It is

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now clear that the binding of NNRTIs provokes substantial conformational changes in both secondary structural elements and in side chains of residues in the NNIBP. These conformational changes in the NNIBP could directly or indirectly affect the precise geometry and/or mobility of the nearby polymerase catalytic site, especially the highly conserved YMDD motif and/or the divalent metal ions [31,33,34,42,68]. The binding of NNRTIs appears to lock the flexible hinge-like structure between the palm and thumb subdomains and restrict mobility of the thumb subdomain, placing constraints on the geometry of the DNA-binding cleft [12,31,34,43]. The “primer grip” (i.e., β12-β13-β14 sheet), which has close interactions with the 3'-terminus of the primer strand [38], forms a part of the NNIBP and is involved in the binding of NNRTIs. It has become apparent that binding of NNRTIs can substantially alter the conformation of the primer grip; this could affect the precise positioning of the primer strand relative to the polymerase active site [34,37]. Displacement of the primer grip by NNRTI binding could lead to repositioning of the primer terminus. This could explain the observation that dNTP binding is largely unaffected by NNRTI binding while the rate of the chemical step of DNA polymerization is reduced [77]. Long-range distortions of the HIV-1 RT structure by NNRTI binding can potentially account for NNRTI inhibition of polymerization [39,41,43] and alteration of RNase H cleavage specificity [43,78]. These possible mechanisms are not mutually exclusive and the binding of inhibitors might have multiple influences on HIV-1 RT polymerization. The exact mechanism(s) of inhibition is still under investigation. IX. NNRTI-Resistance Mutations Analyses of the crystal structures of HIV-1 RT complexed with various NNRTIs have indicated that amino acid residues whose mutations confer high levels of resistance to NNRTIs [9,11,12,26,27] are located close to the bound inhibitors (Figure 5 and Table 2). Subunit-specific mutagenesis studies have confirmed that mutations that confer resistance to the NNRTIs act directly through the change in the NNIBP itself [60,79]. In these studies, recombinant HIV-1 RTs that contained amino acid substitutions only in the p66 subunit were resistant to NNRTIs, while those containing the same amino acid substitutions only in the p51 subunit remained susceptible to the drugs. There is one exception: the amino acid substitution of Glu138 to Lys, which confers resistance to inhibitors only when it is present in the p51 subunit. Amino acid residue 138 is located in the β7–β8 connecting loop of the fingers subdomain. In the p51 subunit this residue forms a part of the NNIBP, while its counterpart in the p66 subunit is far away from the pocket [12,60].

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The mechanism(s) of resistance may depend on the specific amino acid change. It is likely that most NNRTI-resistance mutations exert their effects by altering interactions between protein side chains and the inhibitors [12,34,35]. Drug-resistance mutations that result in a decrease or increase in the size of side chains might lead to loss of favorable contacts or steric conflicts with bound inhibitors. Mutations that alter the local electrostatic potential, i.e., gain, loss, or inversion of charge, may change the affinity of the NNIBP for inhibitor binding. These altered interactions could interfere with the binding of NNRTIs to the hydrophobic pocket or conceivably could even relax the geometric distortion that the binding of an inhibitor causes in the vicinity of the polymerase active site. X. Design of Improved NNRTIs Different NNRTIs, even from the same class of compounds, show remarkable variations in their ability to inhibit HIV-1 replication and can give rise to different spectra of resistance mutations [9,11,26,27]. For example, biochemical studies showed that the 8-chloro TIBO derivative R86183 is quite potent in inhibiting an HIV-1 strain containing the Tyr181Cys mutation, which is one of the frequently occurring HIV-1 RT mutations that gives rise to a high level of resistance to almost all NNRTIs, including other TIBO derivatives [80]. There are several other reports of NNRTIs that are also relatively effective in inhibiting the HIV-1 RT Tyr181Cys variant [81–84]. These results suggest that although all the inhibitors appear to bind in the NNIBP, there are differences in their specific interactions with HIV-1 RT. Structural analyses of HIV-1 RT/NNRTI complexes and computer modeling studies confirmed that the exact conformations of the amino acid residues forming the NNIBP appear to vary in different complexes and that there are specific interactions between individual inhibitor and surrounding residues [33,35,36,85]. However, these differences have not been sufficiently large to allow a successful combination therapy to be developed using two or more of the currently available NNRTIs (discussed in more detailed in a later section) [9,11,26,27,86]. Systematic analysis of wild-type and drug-resistant mutant HIV-1 RT structures in complexes with various NNRTIs should provide additional insights about constraints that could be used to optimize the design of NNRTIs. This knowledge could guide development of more effective inhibitors for AIDs treatment. As discussed earlier, the bound NNRTIs in HIV-1 RT complexes determined so far conform to a common butterfly-like shape (Figure 6). A close inspection of interactions between inhibitors and protein reveals that though

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most of the amino acid residues forming the pocket adjust their side chains to make close contacts with the inhibitor, the inhibitor is not sufficient to fill all of the space in the pocket. There is space for additional nonpolar, polar, or charged groups. Modification of the inhibitor would result in adjustment of the orientation of the side chains and could improve interactions between the inhibitor and surrounding residues such as Leu100, Lys101, Lys103, Val106, or Leu234. Inhibitors designed to have more extensive interactions with essential elements in the pocket should minimize the chances of selecting resistant HIV-1 RT variants. From this point of view, NNRTIs that interact with the relatively conserved residues of the pocket, such as Trp229, Leu234, and Tyr318, may reduce the risk of encountering resistance mutations that do not have significant costs for the enzyme. In addition, compounds could be designed to contain functional groups (for example charged or polar groups) able to fill more of the available space of the NNIBP and also capable of specific hydrophilic interactions with the polar or charged side chains and/or with polypeptide backbone atoms of the NNIBP (for example the main chain amide nitrogens and carbonyl oxygens). The hydrophilic interactions between inhibitors and protein backbone atoms should be advantageous because mutations to any amino acid other than proline would not affect such contacts. In the structures of HIV-1 RT/NNRTI complexes, the bound inhibitors are located very close to the polymerase active site composed of the three catalytically essential aspartic acids Asp110, Asp185, and Asp186. It might be useful to design compounds that have a long and branched aliphatic group or a substituted aromatic group that could not only produce hydrophobic interactions with Tyr181, Tyr188, and Trp229, but could also be able to interact with the three aspartic residues or interfere with the metal ion(s) binding at the polymerase active site. XI. RNase/H-Active Site as a Potential Drug Target Site HIV-1 RT contains RNase H, which is responsible for degradation of viral RNA and removal of RNA primers for minus- and plus-strand DNA synthesis (see reviews [87–89]). The absolute requirement for virus-associated RNase H function [90–93] offers an additional target for antiretroviral drugs. The RNase H domain of HIV-1 RT is located at the C-terminus of the p66 subunit (Figures 2 and 3). In contrast to the polymerase domain of HIV-1 RT, the structure of the RNase H domain is quite similar in all known HIV-1 RT structures and conforms quite well with the structure of the isolated HIV-1 RNase H domain [94–95]. The relative stability of the structure of the RNase H domain suggests that the RNase H active site could be a relatively well-defined target for drug

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design. Mutagenesis studies have demonstrated the interdependence of DNA polymerase and RNase H activities. Mutations that disrupt one of the two enzymatic activities of HIV-1 RT often also impair the second activity [96–99]. Indeed, according to the crystal structure of HIV-1 RT, the polymerase active site and the RNase H active site are separated by approximately 17–18 nucleotides [38] and the RNase H domain has many contacts with the polymerase domain, especially with the connection subdomain of p66 and the thumb and connection subdomains of p51 [31,38,100]. Interactions between the polymerase domain and nucleic acid can modulate RNase H activity. Because the predominant contacts of HIV-1 RT with template-primer occur in the vicinity of the polymerase active site, precise placement of the template strand relative to the RNase H active site may be regulated by the sequence and composition of the template-primer. Mutagenesis experiments showed that mutations located at or near the “template grip” in the polymerase domain of HIV-1 RT can have a greater effect on RNase H than on polymerase activity [99,101,102]. It was also reported that binding of the NNRTI nevirapine alters the cleavage specificity of RNase H [78]. Structural distortions in the position and conformation of template-primer induced by NNRTI-binding may account for alteration of the cleavage specificity of RNase H [43]. Divalent metal ions such as Mg2+ or Mn2+ are essential for the RNase H activity [103–106]. The structure of the isolated RNase H domain crystallized in the presence of MnCl2 revealed two tightly bound Mn2+ ions in close proximity to four catalytically essential acidic residues, Asp443, Glu478, Asp498, and Asp549, that form the active site [94]. Biochemical data have shown that mutations of these conserved residues could either disrupt RNase H activity or lead to a highly unstable enzyme [107–109]. Based on the crystal structures, a two-metal ion-dependent catalytic mechanism for RNase H activity has been postulated [101], which is similar to that proposed for phosphoryl transfer reactions catalyzed by polymerases and their associated nucleases [67,69–71]. In contrast, in the structure E. coli RNase H reported by Katayanagi et al. [111] only one Mg2+ ion was observed, and that led to the proposal of a single metal-ion catalyzed hydrolysis [112]. Interestingly, in the structure of unliganded HIV-1 RT reported by Rodgers et al. [41] and Hsiou et al. [43] only one Mg2+ ion was found at the RNase H active site. The mechanism of RNase H cleavage and the exact role of metal ion(s) in the hydrolysis and formation of phosphodiester bonds are still under investigation (see review [89]). Very few inhibitors specifically target HIV-1 RNase H activity. Illimaquinone, a natural marine product, was shown to preferentially inhibit the HIV-1 RNase H activity [113,114]. However, this compound appears to react with a sulfhydryl group in the polymerase domain and not with RNase H itself. It may be possible to use the available information on structural and biochemi-

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cal properties of the polymerase and RNase H of HIV-1 RT to design compounds that would bind at the RNase H active site or interfere with the metal ion(s) binding and inhibit the RNase H activity of HIV-1 RT. However, for optimal utility, these compounds should selectively inhibit the RNase H activity of HIV-1 and not the RNase H activity of the host cells. XII. Other Possible Target Sites for HIV-1 RT Inhibitors Both polymerase and RNase H activities of HIV-1 RT require that the enzyme be in a dimeric form [115–118] (Figure 3). The exact role(s) of the p51 subunit in the enzymatic activities of HIV-1 RT are not yet known. The three-dimensional structure of HIV-1 RT shows that the interface between p66 and p51 primarily involves interactions between the p66 palm and the p51 fingers subdomains, between the p66 connection and the p51 connection and fingers subdomains, and between the RNase H and the p51 thumb and connection subdomains [34,100,119] (Figure 3). A compound that would interfere with dimerization would be a potential candidate for an anti-AIDS drug. As discussed earlier, the flexibility of HIV-1 RT permits the enzyme to adopt different conformations. In the absence of bound DNA, the thumb and the fingers subdomains come together and close a major portion of the DNA-binding cleft [40,41,43]. Synthetic oligonucleotides that could interact with the specific or conserved regions of the DNA-binding cleft could potentially block binding of templateprimer substrates. An RNA pseudoknot has been reported to bind and specifically inhibit HIV-1 RT [120]. Chemical modification and substitution of specific groups in RNA ligands can change the structure of the pseudoknot, which could result in considerably more effective pseudoknot inhibitors with high binding specificity [121]. Studies employing the phosphorodithioate analogs of the primer sequence recognized by HIV-1 RT showed that these compounds can act as inhibitors and that inhibition is a function of both the sequence and length of these novel single-stranded nucleic acid oligomers [122,123]. A series of natural products, i.e., trihydroxyquinolone compounds isolated from Red Sea marine organisms, were reported to inhibit the DNA polymerase activity of HIV-1 RT [124,125]. This type of inhibitor appears to have a mechanism of inhibition that is different from either the NRTI inhibition mechanism or the NNRTI inhibition mechanism. The inhibition is reversible and noncompetitive with respect to both dNTP and template-primer [125]. This result indicates that there are other potential binding sites for inhibitors of HIV-1 RT.

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XIII. Useful Tools in Structure-Based Drug Design Several computer modeling algorithms have been developed for structure-based drug design. Among them, DOCK [73–75] and 3D SEARCH [126] have been successfully applied in the design of HIV-1 protease inhibitors. These programs search a target protein for invaginations, grooves, and recognition surfaces that could bind a potential receptor molecule. Compounds complementary to the putative receptor binding site in both shape and chemical properties can be identified through searching databases of small molecules, such as the Cambridge Crystallographic Database, the Fine Chemicals Directory, or other commercially available databases. An important issue in analyzing HIV-1 RT is the flexibility of the enzyme. Comparison of structures of unliganded HIV-1 RT and NNRTI-bound HIV-1 RT complexes has shown that the NNIBP is not present in the unliganded form [34,41,43]. This underscores the importance of searching both the unliganded HIV-1 RT and the HIV-1 RT complexes with inhibitors and substrates in order to identify any potential inhibitor-binding sites. Many other approaches have been and are being developed for computeraided design of inhibitors. For example, pharmacophore analysis can identify the spatial arrangement of groups or atoms common to all active inhibitor molecules and then incorporate these elements into a single molecule [127,128]. Detailed analysis of the volumes occupied by different inhibitors bound to the same binding site could also provide new suggestions for inhibitor design. For example, the volume union of all known NNRTIs such as nevirapine, TIBO, α-APA, HEPT, and 1051U91 can be calculated. This type of analysis could be used to screen for new NNRTIs. Since the coordinates for a number of HIV-1 RT/NNRTI complex structures are now available in the Protein Data Bank, these approaches can be applied to the design of new or improved NNRTIs. Given the relatively high flexibility of the NNIBP region and the diversity of NNRTI structures, the NNIBP of HIV-1 RT could be a methodologically challenging yet extremely important target for structure-based drug design. XIV. Enzymatic Efficiency of Drug-Resistant HIV-1 RT Variants Analyses of viral population dynamics indicated that, although drug resistance cannot be seen as a positive outcome of chemotherapy, clinical progress can be made through the development of drugresistant viral variants (see review [30]).alyzed hy Arial H itself. It may be possible to use the available information on structural and biochemi-

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Biochemical data show that HIV-1 replicates extremely rapidly in infected individuals and that the viral load is low in the early stages of the disease because the host immune system is initially successful in limiting viral replication [28,29]. When patients are treated with either RT and/or protease inhibitors, wild-type HIV-1 is rapidly replaced with drug-resistant variants. In fact, even in patients who have not received treatment with any anti-RT drugs, HIV-1 variants that contain residues corresponding to both NRTI- and NNRTI-resistance mutations in RT can be found as minor components of the viral population [129]. Similarly, viral variants that contain residues in protease sequence corresponding to protease inhibitor drug-resistance mutations have also been observed in patients prior to drug therapy (see review [130]). Enzymatic components found in a wild-type virus, such as RT or protease, are optimized for efficient viral replication [30]. In the absence of selective pressure (drug), the wild-type virus has a fitness advantage over drug-resistant viral variants. However, in the presence of drugs, drugresistant variants have a fitness advantage over the wild-type because the drug impairs efficiency of the target enzyme in the wild-type virus [30]. Binding of an NRTI or an NNRTI to wild-type HIV-1 RT interferes with the polymerization reaction. However, the presence of resistant variants in the population allows the virus to escape, and the variants to rapidly replace the wild-type virus. Nevertheless, this escape has a price. When the optimized wildtype virus is replaced by the less fit drug-resistant variants, the relative fitness of the virus decreases. In other words, the enzymatic efficiency of a drug-resistant HIV-1 RT variant is impaired relative to the wild-type enzyme (see review [131]). If the enzymatic efficiency of a drug-resistant viral variant is sufficiently impaired, the replication of the variant virus would be significantly decreased. Thus, an antiviral drug will be useful not because it would completely stop the growth of HIV-1 but because it selects viral variants whose replication is significantly impaired. Positive clinical benefit results from the fact that the viral load is decreased owing to reduced replication of the variant virus. As predicted by this model, some HIV-1 RT and protease inhibitors seem to select for relatively less fit drug-resistant variants. For example, treatment of HIV-1 infection with HBY 097, a quinoxaline inhibitor, induces development of an HIV-1 RT variant containing the Gly190Glu mutation that appears to have substantially decreased polymerase activity and replicates relatively slowly [22,84]. Replacement of the hydrogen atom of Gly190 with an acidic side chain of Glu190 in the hydrophobic NNIBP apparently interferes with the stability of the enzyme as well as the ability of the NNIBP to bind a hydrophobic inhibitor. The relative inefficiency of HIV-1 variant containing the Gly190Glu mutation in RT can be viewed as a positive outcome of the selection pressure provided by this particular inhibitor. However, most of the HIV-1 variants selected by

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currently available antiviral agents are not significantly less fit than the wild-type virus and the clinical benefits are not obvious. In this regard, new drugs should be designed that would be intended to select HIV-1 RT variants that are significantly less fit and do not replicate efficiently. XV. Combination Therapy Using Multiple ANTI-HIV-1 Drugs Monotherapy using either NRTIs or NNRTIs has led to the emergence of drug-resistant viral strains of HIV-1. Though many drug-resistance mutations confer cross-resistance to other inhibitors belonging to the same class, there are indications that some mutations conferring resistance to certain inhibitors are incompatible (see reviews [5,11,131]). A multidrug clinical trial with HIV-1 infected patients has shown that AZT resistance can be reversed by mutations that confer resistance to ddI [8]. The Leu74Val mutation appears to suppress the effects of the Thr215Tyr mutation that confers resistance to AZT [8,27]. The Met184Val mutation, which causes resistance to 3TC or other oxathiolane-cytosine analogs, also appears to reverse the effects of the AZT-resistance mutations [27]. Recent clinical studies have shown that a combination of AZT and 3TC led to a considerable decrease in viral load and a substantial increase of CD4 cells when compared with monotherapy using AZT alone, even after emergence of the Met184Val mutation [132]. Another example is the Pro236Leu mutation that confers resistance to BHAP. The sensitivity of this HIV-1 RT variant to TIBO, nevirapine, and pyridinone is increased ten fold [133]. Although the NRTIs and NNRTIs target two distinct binding sites of HIV-1 RT and lead to different sets of resistance mutations, some of the NRTI- and NNRTI-resistance mutations also appear to be incompatible. For example, the NNRTI-resistance mutations Leu100Ile and Tyr181Cys have been shown to suppress the effects of some AZT-resistance mutations [11,134]. This has led to the suggestion that a combination of anti-HIV-1 drugs would be more effective in inhibiting HIV-1 replication than using individual drugs alone. In fact, both clinical and in vitro studies have shown that combination therapy has considerable advantages over monotherapy. At least in some cases, the effectiveness of the therapy increases with an increase in the number of drugs in the combination [5,135]. Combination therapy may, in addition to increasing antiviral activity, also slow emergence of drug-resistant variants and may have the added benefit that reducing the dosage of individual drugs can reduce toxicity. It is generally believed that synergistic drug interactions arise from the fact that certain combinations of drugresistance mutations are particularly detrimental for the enzyme (and, by extension, the virus). This has focused attention on

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determining the mechanism(s) underlying drug resistance and, from this understanding, to devise ways for identifying combinations of drugs which might provoke drug-resistance mutations incompatible with viral survival. Several protocols have been designed for combination therapy using a variety of anti-HIV-1 drugs (see reviews [5,11]). Combinations of different drugs that interact with the same binding site of the same viral protein but lead to mutually antagonistic or suppressive resistance mutations have been studied extensively, especially for the combined uses of different but structurally related NRTIs (see for example [136–138]) or NNRTIs [139–141]. Combinations of drugs or inhibitors that target different sites of the same viral protein, primarily the combination of NRTIs and NNRTIs of HIV-1 RT, show enhanced inhibition of HIV-1 RT polymerase activity and suppression of the emergence of drugresistance mutations (for example [142–146]). Experiments have also been conducted with combinations of drugs that target different viral proteins, e.g., inhibitors of virus adsorption, virus-cell fusion, and/or uncoating proteins have been tested in combination with protease inhibitors and/or RT inhibitors. Combinations of AZT with the glycosylation inhibitor castanospermine [147], or with the Tat inhibitor Ro 24-7429 [148], or with the protease inhibitor Ro 31-8959 [149] have been shown to potently inhibit HIV-1 viral replication in vitro. Combination therapy can increase the effectiveness of inhibition and significantly impair efficiency of viral replication. However, both NRTI- and NNRTI-resistance mutations can affect the positioning of the nucleic acid and/or the overall structure of HIV-1 RT [23]. These two sets of resistance mutations can communicate with each other and can result in cross resistance. Moreover, new drug-resistance mutations that confer cross-resistance to both NRTIs and NNRTIs can be selected, which reduce the effectiveness of some drug combinations (see reviews [5,11,26]). Biochemical studies showed that both HIV-1 RT mutants [150] and viral variants [151] could be obtained that are resistant to the combination of AZT, ddI, and nevirapine. In clinical trials, treatment with AZT and ddI or AZT and ddC led to a different spectrum of NRTI-resistance mutations [152,153]. The most notable of these new mutations is Gln151Met, which is located at a position close to the dNTP-binding site. Structural analysis of the HIV1 RT/DNA/Fab complex suggests that the side chain of Gln151 in the wild-type enzyme may interact with the first unpaired template nucleotide. The side chain of this residue may play a role in selecting the correct base for the incoming nucleotide [72]. Since the RT mutant containing only the Gln151Met mutation can confer high-level resistance to a number of NRTIs, including AZT, ddI, and ddC, it is not clear why this mutation did not emerge in monotherapy of these NRTIs. However, Gln151 is relatively well conserved and mutations at this position may have an unfavorable impact on HIV-1 RT.

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XVI. Perspective Substantial progress has been made in understanding the structure and function of HIV-1 RT and in the development of anti-HIV-1 inhibitors. However, the genetic flexibility of HIV-1 will continue to make development of a truly effective antiviral therapy for AIDS an exceptionally difficult task. We are beginning to understand how to circumvent drug resistance. The accumulated evidence has shown that the ability of the virus to develop drug resistance is limited and that the drug-resistant viral variants are less efficient than the wild-type virus. If the selection pressure provided by antiviral drugs makes the virus pay a sufficiently high price, then the viral load can be decreased and there will be a measurable clinical benefit. Based on a better understanding of the structure-function relationships of HIV-1 RT, we are now coming to grips with the mechanisms of polymerization, drug inhibition, and drug resistance. This information should make it possible to develop new or improved HIV-1 RT inhibitors that have different properties and provoke different patterns of drug-resistance mutations. Though it is likely that there will be no single drug which would be effective against all HIV-1 variants, we have reasons to believe that new or improved drugs or, more likely, new drug combinations, will be designed that are broadly effective against all of the HIV-1 variants that can grow efficiently. Detailed analysis of the conformational changes among the various HIV-1 RT structures may reveal additional sites (in addition to the currently known NRTI- and NNRTI-binding sites) for binding new inhibitors able to interfere with the polymerization and/or the flexibility of the enzyme required for its activity. The considerable physical and genetic flexibility of HIV-1 RT suggests that more effective anti-RT drugs should be designed to target the conserved portions of HIV-1 RT that the virus cannot easily afford to change. Such conserved elements can be identified by comparing the sequences of RTs from different retroviruses; the functions and relative importance of these conserved elements can be determined by mutagenesis and biochemical and structural analyses. It is our hope that application of structure-based drug design strategies may aid in the development of novel HIV-1 RT inhibitors for a more effective treatment of HIV-1 infection. Acknowledgments We thank the other members of the Arnold and Hughes laboratories and our collaborators for their helpful discussions and assistance, including Koen Andries, Gail Ferstandig Arnold, Paul Boyer, Arthur Clark, Jr., Paul Janssen, Jörg-Peter Kleim, Luc Koymans, Tack Kuntz, Karen Lentz, Chris Michejda, Henri Moereels, Manfred Roesner, Marilyn Kroeger Smith, Rick Smith, Jr., and

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Chris Tantillo. The work in Edward Arnold's laboratory has been supported by Janssen Research Foundation and NIH grants (AI 27690 and AI 36144). Research in Stephen Hughes' laboratory is sponsored in part by the National Cancer Institute, DHHS, under contract with ABL, and by NIGMS. References 1. Goff SP. Retroviral reverse transcriptase: synthesis, structure, and function. J Acquired Immune Deficiency Syndromes 1990; 3:817–831. 2. Jacobo-Molina A, Arnold E. HIV reverse transcriptase structure-function relationships. Biochemistry 1991; 30:6351–6361. 3. Whitcomb JM, Hughes SH. Retroviral reverse transcription and integration: progress and problems. Ann Rev Cell Biol 1992; 8:275–306. 4. Le Grice SFJ. Human immunodeficiency virus reverse transcriptase. In: Skalka AM, Goff SP, eds. Reverse Transcriptase. Plainview, New York: Cold Spring Harbor Laboratory Press, 1993:163–191. 5. De Clercq E. Toward improved anti-HIV chemotherapy: therapeutic strategies for intervention with HIV infections. J Med Chem 1995; 38:2491–2517. 6. De Clercq E. HIV inhibitors targeted at the reverse transcriptase. AIDS Res Human Retroviruses 1992; 8:119–134. 7. Larder BA. Inhibitors of HIV reverse transcriptase as antiviral agents and drug resistance. In: Skalka AM, Goff SP, eds. Reverse Transcriptase. Plainview, New York: Cold Spring Harbor Laboratory Press, 1993:205–222. 8. St. Clair MH, Martin JL, Tudor-Williams G, Bach MC, Vavro CL, King DM, et al. Resistance to ddI and sensitivity to AZT induced by a mutation in HIV-1 reverse transcriptase. Science 1991; 253:1557–1559. 9. Richman DD. Resistance of clinical isolates of human immunodeficiency virus to antiretroviral agents. Antimicrob Agents Chemother 1993; 37:1207–1213. 10. Schinazi RF. Competitive inhibitors of human immunodeficiency virus reverse transcriptase. Perspectives in Drug Discovery and Design 1993; 1:151–180. 11. De Clercq E. HIV resistance to reverse transcriptase inhibitors. Biochem Pharmacol 1994; 47:155–169.

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12. Tantillo C, Ding J, Jacobo-Molina A, Nanni RG, Boyer PL, Hughes SH, et al. Locations of antiAIDS drug binding sites and resistance mutations in the three-dimensional structure of HIV-1 reverse transcriptase: implications for mechanisms of drug inhibition and resistance. J Mol Biol 1994; 243:369–387. 13. Miyasaka T, Tanaka H, Baba M, Hayakawa H, Walker RT, Balzarini J, et al. A novel lead for specific anti-HIV-1 agents: 1-[(2-hydroxyethoxy)methyl]-6-(phenylthio)thymine. J Med Chem 1989; 32:2507–2509. 14. Pauwels R, Andries K, Desmyter J, Schols D, Kukla MJ, Breslin HJ, et al. Potent and selective inhibition of HIV-1 replication in vitro by a novel series of TIBO derivatives. Nature 1990; 343:470–474. 15. Merluzzi VJ, Hargrave KD, Labadia M, Grozinger K, Skoog M, Wu JC, et al. Inhibition of HIV-1 replication by a nonnucleoside reverse transcriptase inhibitor. Science 1990; 250:1411–1413.

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27. Kimberlin DW, Coen, DM, Biron KK, Cohen JI, Lamb RA, McKinlay M, et al. Molecular mechanisms of antiviral resistance. Antiviral Res 1995; 26:369–401. 28. Wei X, Ghosh SK, Taylor ME, Johnson VA, Emini EA, Deutsch P, et al. Viral dynamics in human immunodeficiency virus type 1 infection. Nature 1995; 373:117–122. 29. Ho DD, Neumann AU, Perelson AS, Chen W, Leonard JM, Markowitz M. Rapid turnover of plasma virions and CD4 lymphocytes in HIV-1 infection. Nature 1995; 373:123–126.

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30. Coffin JM. HIV population dynamics in vivo: implications for genetic variation, pathogenesis, and therapy. Science 1995; 267:483–489. 31. Kohlstaedt LA, Wang J, Friedman JM, Rice PA, Steitz TA. Crystal structure at 3.5 Å resolution of HIV-1 reverse transcriptase complexed with an inhibitor. Science 1992; 256:1783–1790. 32. Smerdon SJ, Jager J, Wang J, Kohlstaedt LA, Chirino AJ, Friedman JM, et al. Structure of the binding site for nonnucleoside inhibitors of the reverse transcriptase of human immunodeficiency virus type 1. Proc Natl Acad Sci USA 1994; 91:3911–3915. 33. Ren J, Esnouf R, Garman E, Somers D, Ross C, Kirby I, et al. High resolution structures of HIV-1 RT from four RT-inhibitor complexes. Nature Struct Biol 1995; 2:293–302. 34. Ding J, Das K, Tantillo C, Zhang W, Clark AD Jr., Jessen S, et al. Structure of HIV-1 reverse transcriptase in a complex with the nonnucleoside inhibitor α-APA R 95845 at 2.8 Å resolution. Structure 1995; 3:365–379. 35. Ding J, Das K, Moereels H, Koymans L, Andries K, Janssen PAJ, et al. Structure of HIV-1 RT/TIBO R 86183 reveals similarity in the binding of diverse nonnucleoside inhibitors. Nature Struct Biol 1995; 2:407–415. 36. Ren J, Esnouf R, Hopkins A, Ross C, Jones Y, Stammers D, et al. The structure of HIV-1 reverse transcriptase complexed with 9-chloro-TIBO: lessons for inhibitor design. Structure 1995; 3:915–926. 37. Das K, Ding J, Hsiou Y, Clark AD Jr., Moereels H, Koymans L, et al. Crystal structures of 8-Cl and 9-Cl TIBO complexed with wild-type HIV-1 RT and 8-Cl TIBO complexed with the Tyr181Cys HIV-1 RT drug-resistant mutant. J. Mol Biol 1996; 264:1085–1100. 38. Jacobo-Molina A, Ding J, Nanni RG, Clark AD Jr., Lu X, Tantillo C, et al. Crystal structure of human immunodeficiency virus type 1 reverse transcriptase complexed with double-stranded DNA at 3.0 Å resolution shows bent DNA. Proc Natl Acad Sci USA 1993; 90:6320–6324. 39. Jager J, Smerdon S, Wang J, Boisvert DC, Steitz TA. Comparison of three different crystal forms shows HIV-1 reverse transcriptase displays an internal swivel motion. Structure 1994; 2:869–876. 40. Raag R, Clark AD Jr., Ding J, Jacobo-Molina A, Lu X, Nanni RG, et al. 3.0 Å crystal Structure of HIV-1 reverse transcriptase without dsDNA reveals largescale motion of p66 thumb subdomain. Am Crystallogr Assoc Mtg Abstr, Ser 2 1994; 18:44. 41. Rodgers DW, Gamblin SJ, Harris BA, Ray S, Culp JS, Hellmig B, et al. The structure of unliganded reverse transcriptase from the human immunodeficiency virus type 1. Pro Natl Acad Sci USA 1995; 92:1222–1226.

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42. Esnouf R, Ren J, Ross R, Jones Y, Stammers D, Stuart D. Mechanism of inhibition of HIV-1 reverse transcriptase by non-nucleoside inhibitors. Nature Struct Biol 1995; 2:303–308. 43. Hsiou Y, Ding J, Das K, Clark AD Jr., Hughes SH, Arnold E. Structure of unliganded HIV-1 reverse transcriptase at 2.7 Å resolution: implications of conformational changes for polymerization and inhibition mechanisms. Structure 1996; 4:853–860. 44. Unge T, Knight S, Bhikhabhai R, Lovgren S, Dauter Z, Wilson K, et al. 2.2 Å resolution structure of the amino-terminal half of HIV-1 reverse transcriptase (fingers and palm subdomains). Structure 1994; 2:953–961.

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45. Majumadar C, Abbotts J, Broder S, Wilson SH. Studies on the mechanism of human immunodeficiency virus reverse transcriptase: Steady-state kinetics, processivity and polynucleotide inhibiton. J Biol Chem 1988; 263:15657–15665. 46. Kati WM, Johnson KA, Jerva LF, Anderson KS. Mechanism and fidelity of HIV reverse transcriptase. J Biol Chem 1992; 267:25988–25997. 47. Balzarini J, Herdewijn P, De Clercq E. Differential patterns of intracellular metabolism of 2',3'didehydro-2',3'-dideoxthymidine and 3'-azido-2',3'- dideoxythymidine: two potent anti-human immunodeficiency virus compounds. J Biol Chem 1989; 264:6127–6133. 48. Balzarini J, De Clercq E. 5' -Phosphoribosyl 1-pyrophosphate synthetase converts the acyclic nucleoside phosphonates 9-(3-hydroxy-2-phosphonyl-methoxypropyl) adenine and 9-(2phosphonylmethoxyethyl)adenine directly to their antivirally active diphosphate derivatives. J Biol Chem 1991; 266:8686–8689. 49. Merta AI, Votruba J, Jindrich J, Holy A, Cihlar T, Rosenberg I, et al. Phosphorylation of 9-(2phosphonylmethoxyethyl)adenine and (S)-9-(3-fluoro-2-phosphonylmethoxypropyl)adenine by AMP (dAMP) kinase from L1210 cells. Biochem Pharmacol 1992; 44:2067–2077. 50. De Clercq E. Broad spectrum anti-DNA virus and anti-retrovirus activity of phosphonylmethoxyalkyl-purines and -pyrimidines. Biochem Pharmacol 1991; 42:963–972. 51. Balzarini J, Hao Z, Herdewijn P, Johns DG, De Clercq E. Intracellular metabolism and mechanism of anti-retrovirus action of 9-(2-phosphonyl-methoxyethyl) adenine, a potent anti-human immunodeficiency virus compound. Proc Natl Acad Sci USA 1991; 88:1499–1503. 52. Balzarini J, Holy A, Jindrich J, Dvorakova H, Hao Z, Snoeck R, et al. 9-[(2RS)-3- fluoro-2phosphonylmethoxyethyl] derivatives of purines: a class of highly selective antiretroviral agents in vitro and in vivo. Proc Natl Acad Sci USA 1991; 88:4961–4965. 53. Patel PH, Jacobo-Molina A, Ding J, Tantillo C, Clark AD Jr., Raag R, et al. Insights into DNA polymerization mechanisms from structure and function analysis of HIV-1 reverse transcriptase. Biochemistry 1995; 34:5351–5363. 54. Boyer PL, Tantillo C, Jacobo-Molina A, Nanni RG, Ding J, Arnold E, et al. The sensitivity of wildtype human immunodeficiency virus type 1 reverse transcriptase to dideoxynucleotides depends on template length; the sensitivity of drug-resistant mutants does not. Proc Natl Acad Sci USA 1994; 91:4882–4886. 55. Kellam P, Boucher CA, Larder BA. Fifth mutation in human immunodeficiency virus type 1 reverse transcriptase contributes to the development of high-level resistance to zidovudine. Proc Natl Acad Sci USA 1992; 89:1934–1938.

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56. Martin JL, Wilson JE, Haynes RL, Furman PA. Mechanism of human immunodeficiency virus type 1 resistance to dideoxyinosine. Proc Natl Acad Sci USA 1993; 90:6135–6139. 57. Lacey SF, Reardon JE, Furfine ES, Kunkel TA, Bebenek K, Eckert KA, et al. Biochemical studies on the reverse transcriptase and RNase H activities from human immunodeficiency virus strains resistant to 3'-azido-3'-deoxythymidine. J Biol Chem 1992; 267:15789–15794. 58. Larder BA. 3'-Azido-3'-deoxythymidine resistance suppressed by a mutation conferring human immunodeficiency virus type 1 resistance to nonnucleoside

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3 Retroviral Integrase: Structure as a Foundation for Drug Design Alison B. Hickman and Fred Dyda National Institutes of Health, Bethesda, Maryland I. Introduction A. Retroviral Lifecycle The human immunodeficiency virus (HIV) is one of only a few retroviruses known to infect humans. It is estimated that approximately twenty-two million people are now infected worldwide [1]. With only a tiny number of exceptions, infection ultimately leads to the development of the lethal condition of acquired immunodeficiency syndrome, or AIDS. To date, only a handful of drugs have been shown to have any effect on the course of the disease. These are, in general, relatively ineffective at significantly prolonging life, and drug resistance develops rapidly. Equally discouraging, vaccines have not yet been developed to prevent infection. The retroviral lifecycle presents several steps that can be targeted as possible sites of intervention by inhibitors. As shown in Figure 1, when a retrovirus encounters a host cell, specific recognition between proteins on the surface of the virus and receptors on the host cell surface leads to membrane fusion. The viral core then enters the cell cytoplasm where the process of reverse transcription begins. The requirement of the conversion of viral RNA to double-stranded DNA is a feature unique to retroviruses. With the recent exception of the protease inhibitor saquinavir, ritonavir, and indinavir, the drugs approved to date by the U.S. Food and Drug Administration (FDA) for the treatment of HIV infection have been nucleoside analogs targeted against the viral enzyme that carries out this conversion, reverse transcriptase.

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Figure 1 Retroviral lifecycle as summarized in Reference 67. Reprinted by permission of Springer-Verlag Publishing Co., New York, NY.

Although details of the timing of reverse transcription, nuclear localization, and integration are not yet clear, it is generally recognized that the movement of double-stranded viral DNA across the nuclear membrane is followed by insertion, or integration, of the viral genome into a host-cell chromosome. The viral DNA moves as part of a larger “preintegration complex,” a high-molecular-weight aggregate whose composition has not yet been completely defined. The end result of integration is the incorporation of the viral DNA into the DNA of the host cell. Once there, the provirus can serve as a template for the production of mRNA, allowing for the synthesis of viral proteins. These are assembled at the cell membrane to produce new viral particles, which then bud off to seek out new cells to infect. The integrated viral DNA is also necessarily copied whenever the host cell undergoes cell division. The insidious nature of

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the virus arises because, once integrated, the viral DNA can no longer be distinguished from host cell DNA and has become a permanent fixture of the host cell genome. B. Rationale for Drug Design Against Integrase to Fight HIV and AIDS It has been demonstrated that the chemical steps that comprise DNA integration are carried out by the viral protein, integrase (IN). Integrase is encoded by the 3' end of the viral pol gene, which also codes for two other viral enzymes, the protease and reverse transcriptase. These three enzymes are initially synthesized as part of a larger polyprotein that is subsequently cleaved by the protease to the individual proteins. Why is integrase a good target for drug-design efforts to prevent infection by halting the viral replication cycle? First, integration is required for replication. In the absence of integration, the virus is unable to continue to make copies of itself. Secondly, the enzyme that carries out integration is virally encoded, and when the viral genome is disrupted so that functional integrase is no longer made, sustained viral replication does not occur [2]. This demonstrates that if viral integrase can be effectively inhibited, there is no protein encoded by the host cell that can replace it and carry out viral integration. Finally, since mammalian cells do not have enzymes capable of integrating HIV DNA, there are no vital host cell analogs of integrase carrying out essential reactions whose function would be blocked by integrase inhibitors. Effective inhibition of HIV integrase would add to the number of sites at which the virus replication cycle can be halted. One can imagine treatment protocols in which a mixture of inhibitors, each aimed at a different viral protein, could be administered. This is known as divergent combination therapy. As structural details are a necessary starting point for rational drug design, we present here our recent results on the high-resolution three-dimensional structure of the catalytic core domain of HIV-1 integrase [3]. We also review the current literature discussing integrase inhibitors and present thoughts on ways in which knowledge of the chemical reactions carried out by integrase and its structure might direct the development of effective inhibitors. II. Biochemical Reactions Catalyzed By HIV Integrase A. In Vivo Integration Details of the initial chemical reactions that occur during HIV integration are now well understood (for reviews, see References 4,5). Once linear double-stranded DNA is available for integration, (Figure 2a) integrase then removes

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Figure 2 In vivo reactions carried out by HIV integrase.

two nucleotides from each 3' end of the viral DNA (Figure 2b). The two nucleotides are removed as a dinucleotide rather than in two individual steps. The specificity for this reaction is conferred by the third and fourth nucleotides from each 3' end, a -CA sequence that is absolutely conserved. Once two nucleotides have been removed, leaving recessed 3' hydroxyl groups, the next step is the joining of the 3' ends to target DNA (Figure 2c, d). This process, known as double-ended integration, occurs on opposite strands such that the joining sites on each of the target DNA strands are separated by five base pairs. The final step in integration is the repair of the single-stranded gaps generated by the staggered insertion of the viral 3' ends on opposite strands; this regenerates an intact double-stranded DNA molecule (Figure 2e and f). Gap repair is probably carried out by host cell DNA repair systems. One necessary consequence of retroviral integration is the duplication of five base pairs of host cell DNA on either side of the integrated provirus.

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Another is the loss from the ends of the viral DNA of the original two base pairs that preceded the conserved 3'-CA. B. In Vitro Assays to Monitor Integrase Activity In contrast to the in vivo reaction, concerted integration in vitro of two HIV DNA ends into a target DNA molecule separated by a 5 base-pair stagger occurs very inefficiently. However, in vitro systems have been developed [6,7] using recombinant HIV integrase that have allowed the chemistry of the single-ended integration event to be studied in fine detail. It is possible and routine to use short, doublestranded synthetic oligonucleotides that mimic the viral ends to monitor the removal of two nucleotides from 3' ends (denoted 3' processing or cutting) and the subsequent insertion of one 3' processed DNA molecule into another (known as strand transfer or joining). Typical reactions are depicted in Figure 3. The stereochemical mechanism of 3' processing and strand transfer has been investigated using DNA substrates that incorporate phosphorothioate link-ages [8]. For both reactions, the introduced chiral centers are inverted in the products, implying that the reactions occur via a one-step in-line displacement mechanism rather than via a covalent intermediate. A third assay of integrase activity, termed disintegration, has more recently been developed [9] that monitors the apparent reversal of strand-

Figure 3 Reactions carried out by integrase in vitro, using short oligonucleotide substrates.

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transfer (Figure 3). While disintegration probably has no physiological significance, it has been useful in defining aspects of integrase biochemistry. The three in vitro activities of integrase require divalent metal ions as cofactors. The only two metals that support these activities are Mn2+ and Mg2+. Since quite high metal concentrations must be added to assays (1–10 mM for optimal activity), it has been presumed that Mg2+ is the ion used in vivo. C. Evidence for a Multimer as the Active Unit of Integrase Several lines of evidence demonstrate that the active unit of integrase is a multimer. It is clear, as an isolated protein in solution, that integrase forms dimers [6,10–12], and it has been shown by sedimentation equilibrium studies that Rous sarcoma virus (RSV) integrase exists in reversible equilibrium between monomeric, dimeric, and tetrameric forms [13]. Protein-protein cross-linking studies of HIV-1 [14] and RSV [15] integrases confirm the existence of protein dimers and tetramers in solution, and in vivo, the yeast GAL4 two-hybrid system has demonstrated that HIV-1 integrase can interact with itself [16]. Complementation studies using mutant proteins in vitro provide compelling evidence that the active form of integrase must be at least a dimer [14, 17]. This can be inferred from the result that when certain inactive forms of integrase—generated either by truncation or point mutation—are mixed, robust activity can be reconstituted. This indicates that different monomers in a multimer are capable of providing different essential functions in the context of an active complex. Collectively, these studies suggest that integrase acts as a multimer. This would also seem the most straight-forward model to explain the observation that viral integration requires two coordinated cutting and joining reactions on the target DNA during strand transfer. However, physical studies have not yet addressed what form of integrase actually binds to DNA and carries out the chemical reactions of integration. III. Properties of HIV-1 Integrase A. Domain Structure of Retroviral Integrases A consistent view of the domain structure of retroviral integrases has emerged by combining the results from biochemical studies using deletion and site-specific mutants, limited proteolysis experiments, and sequence comparisons among the family of retroviral integrases. The organization of the domains of integrase is shown schematically in Figure 4. The central domain of HIV-1 integrase, consisting approximately of residues 50 to 200, is largely conserved among retroviral integrases, and forms

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Figure 4 Schematic domain structure of HIV-1 integrase as adapted from Engelman et al. [19]. Structures of two domains, the catalytic core extending from residues 50 to 212 [3] and the nonspecific DNA-binding domain from residues 220 to 270 [28,29], have recently been determined by x-ray crystallography and NMR spectroscopy, respectively.

the protease-resistant core of the protein [18,19]. Within this domain are three invariant residues that comprise the “D,D-35-E motif” (see alignment in Figure 5). These are residues Asp64, Asp116, and Glu152. Even conservative substitution of any of these residues leads to loss of all three in vitro activities of integrase in parallel [19–21]. The D,D-35-E motif is also observed in retrotransposons and some prokaryotic transposases. A truncated form of HIV-1 integrase consisting of residues 50 to 212 is capable of disintegration [22], implying that the catalytic site is contained within this domain. These observations and the absolute requirement for metals for in vitro activity have led to the proposal that the three acidic residues constitute a divalent metal-binding site capable of binding one or two Mg2+ or Mn2+ ions to form a catalytically active enzyme. As will be seen in later sections, the three-dimensional structure of the core domain of HIV-1 integrase is consistent with this hypothesis. The catalytic mechanism may be, therefore, similar to the one proposed by Beese and Steitz for the 3'–5' exonuclease of E. coli DNA polymerase I [23]. It is proposed that for phosphate bond cleavage, one metal ion helps form the attacking hydroxide ion while the other stabilizes a pentacovalent intermediate around the phosphorus. The C-terminus of HIV-1 integrase, consisting approximately of residues 210 to 288, includes the dominant nonspecific DNA binding domain [24, 25], which has been more finely mapped to residues 220–270 [26]. The C-terminus is the least conserved region of retroviral integrases; only one residue, Trp235, is absolutely invariant. However, it has been reported that removal of only five amino acids from the C-terminus of HIV-1 integrase is enough to severely reduce its 3' processing and strand transfer activities [27]. One notable feature of the C-terminus is its high proportion of positively charged residues. As discussed in Section IV.E, the structure of part of this region has recently been determined using NMR spectroscopic methods [28,29].

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Figure 5 Alignment of amino acid sequences of retroviruses. See Engelman et al. [19] for details.

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The role of the N-terminus of integrase, residues 1 to 50, is still unclear. Within this region are four strictly conserved amino acids: two His and two Cys residues. In HIV-1 integrase, the spacing is His-X3His-X23-Cys-X2-Cys. This cluster of His and Cys residues is reminiscent of a zinc-binding motif, and it has been demonstrated that the full-length protein binds Zn2+ [22,30], and that the separately expressed domain consisting only of residues 1 to 55 also binds Zn2+ stoichiometrically [31]. However, it has not been shown that either the structural integrity or the enzymatic activities of integrase require Zn2+. While truncation of residues from the N-terminus of HIV integrase results in loss of 3' processing and strand transfer activities [22,25], in the case of RSV integrase, the N-terminal region can be replaced by unrelated sequences, and the enzyme is still capable of all three in vitro activities [32]. B. Biophysical Properties of Full-Length Recombinant HIV-1 Integrase It has been known for some time that recombinant HIV-1 integrase is a particularly poorly behaved protein in solution. Its solubility in most usual buffers is limited to approximately 1 mg/mL, and even then only in the presence of high concentrations of NaCl. At ~1 mg/mL, HIV-1 integrase slowly precipitates out of solution, revealing one of its characteristic features, a tendency towards aggregation. These properties of the protein are not unreasonable, since in its viral environment integrase is probably never required to be a soluble protein. To maintain the integrity of preintegration complexes, it may even be advantageous for the protein to have the properties of being rather insoluble and sticking to itself, nucleic acid, and perhaps other proteins. C. Properties of Truncated Versions of HIV-1 Integrase It has been our approach to protein structure determination by x-ray crystallography that it is imperative to begin with well-characterized and well-behaved protein. In particular, it is important that the protein be reasonably soluble and monodisperse in solution. Unfortunately, as discussed above, recombinant HIV-1 integrase satisfies neither of these conditions. One approach we and others have taken to circumvent these problems has been to examine truncated versions of HIV-1 integrase to determine if removal of amino acids from either terminus or both affects solubility and aggregation properties. Although we observed that two proteins we constructed, IN213–288 and IN50–288, were more soluble than the full-length HIV-1 integrase, IN1–288 [33, and unpublished observations], our first target protein for crystallization efforts was the core domain of HIV-1 integrase consisting of residues 50 to 212, IN50–212. We reasoned that this protein domain was likely to be compact and

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well folded since it is relatively resistant to proteolysis. As it is also active for disintegration, we concluded that it contained the enzyme active site. We exploited the convenience of histidine-tag (HT) technology to develop methods to purify large quantities of IN50–212 [12]. A 20-amino-acid histidine-containing tag was added to the N-terminus of HIV-1 IN50–212 [22] to allow rapid purification on nickel affinity columns. It was subsequently removed by thrombin cleavage. Biophysical studies showed that although buffer conditions could be identified where the protein was soluble to ~ 4 mg/mL, under these conditions the protein was highly aggregated (unpublished observations). Although the aggregation problem could be largely avoided by the addition of high concentrations of the zwitterionic detergent CHAPS, conditions could not be identified under which protein crystals formed in the presence of CHAPS. D. Systematic Mutation of Hydrophobic Residues to Improve Protein Solubility As it became clear that IN50–212 was crystallographically challenged, a condition readily understood in terms of its aggregation problems and low solubility, a more radical approach was undertaken to try and improve its biophysical properties. Hydrophobic residues in the catalytic core were targeted for sitespecific mutation according to the following criteria: where two or more hydrophobic residues were encountered close together in the primary amino acid sequence, they were each changed to an alanine residue. When a hydrophobic residue stood alone, it was mutated to lysine. In this way, 29 different mutant proteins of IN50–212 were rapidly generated using the overlapping polymerase chain reaction (PCR) and screened for improved solubility properties [34]. Three mutated proteins were identified that were more soluble at lower NaCl concentration than the unmutated core (V165K, F185K, and the double mutation of W131A/W132A). However, one of these in which Phe185 was mutated to Lys had dramatically improved solubility and was ultimately crystallized and its three-dimensional structure determined [3]. The remarkable biophysical properties of this single point mutant of IN50–212 have recently been described [34]. IV. Structure Of The Catalytic Core Domain Of HIV-1 Integrase A. Description of the Structure The Overall Protein Fold The three-dimensional structure of the catalytic core domain of HIV-1 integrase is centered on a mixed five stranded β sheet flanked by several helices forming

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Figure 6 Molscript stereo figure of the three-dimensional structure of the catalytic core of HIV-1 integrase. The two catalytically essential aspartic acid residues (D64 and D116) visible in the x-ray structure are highlighted.

an α-β meander sandwich (see Figure 6) [35]. In the crystal structure, interpretable electron density starts at Cys56, leading to a short loop. The first β strand starts at Gly59 and runs until Val68. The two central residues of a type I' reverse β turn, Glu69 and Gly70, change the polypeptide chain direction to form the second β strand between residues Lys71 and His78, which runs antiparallel with the first strand. A type I'β turn follows, with Val79 and Ala80 changing the chain direction again to form the third β strand between Ser81 and Ile89, which runs antiparallel with the second strand. A short loop between Pro90 and Glu92 leads to the first α helix (helix A) between Thr93 and Trp108. This helix packs against the bottom face of the sheet formed by the first three antiparallel strands by several hydrophobic interactions. A short loop (Pro109 and Val110) leads to the fourth β strand between Lys111 and His114. This strand is parallel with the first. A short loop starting at Thr115 leads to helix B, a one turn helix between Gly118 and Thr122, followed by helix C between Ser123 and Ala133. This helix runs parallel to and packs against helix A. The residues Gly134 and Ile135 form a short loop prior to the fifth and last β strand of the structure between Lys136 and Ala138. This short strand is parallel with the first and the fourth. There is no interpretable electron density due to disorder between Gly140 and Met154. At Met154, the fourth α helix (helix D) starts and runs until Ala169 on the top face of the sheet formed by the first three β strands. The residue Glu170 leads into the next helix (helix E) running between His171 and Lys186, the first residue of a short basic sequence (Lys186, Arg187, and Lys188). Together with

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Gly193 and Tyr194, Lys188 and Gly189 form two short antiparallel β strands separated by a turn of three residues (Gly190, Ile191, and Gly192) not involved in main chain hydrogen bonds. The first residue of the last helix, (helix F), which runs until Asp212, is Ser194. Three Conserved Acidic Residues at the Enzyme Active Site There are four amino acids in the core domain sequence that are absolutely conserved among retroviral integrases: Asp64, Asp116, Glu152, and Lys159. The three acidic residues form the conserved D,D-35E motif and have been shown to be essential for catalysis (see Section III.A). The role of Lys159 in retroviral integrases is not obvious; its replacement with Val does not abolish catalytic activity, although there is a decrease in strand transfer activity [20]. The first essential acidic residue, Asp64, is located in the middle of the first β strand, while Asp116 is in a loop region right after the fourth β strand. These two residues define the active-site area and they are right next to each other three-dimensionally with their α-carbons separated by only 6.7 Å. The closest approach is 3.4 Å between Oδ1 of Asp64 and Cβ of Asp116. These residues are on the surface of the molecule, not part of any obvious substrate-binding cleft. The third catalytically essential acidic residue, Glu152, is in the disordered and hence crystallographically invisible region between Gly140 and Met154. Its location therefore must be inferred from other parts of the structure and from available threedimensional structures of related proteins. The location of Met154, the residue only two positions upstream from Glu152, is known because of interpretable electron density. The distance between the αcarbons of Glu152 and Met154 cannot be larger than about 7.3 Å, which constrains Glu152 to the neighborhood of the two other essential carboxylates, allowing it to contribute to the formation of a divalent metal-binding site. More recently, a crystal structure of the avian sarcoma virus (ASV) integrase core domain was solved [36]. Within this domain, ASV integrase has 24% sequence identity to the HIV-1 integrase core and, as expected, its three-dimensional structure is remarkably similar. The ASV integrase core in its native form has much better solution properties than the HIV-1 integrase core, and did not require any point mutations to render it crystallizable. Due to this fact and perhaps also due to its different crystal packing interactions, the crystal lattice of the ASV integrase core domain is somewhat more ordered than that of HIV-1. The two three-dimensional structures can be aligned quite well, using 74 α-carbons, with an rms deviation of only 1.4 Å in these α-carbon positions. The most remarkable difference between the two structures is that in the ASV structure the electron density is interpretable in all parts of the molecule. This is not to say, however, that serious disorder is not present. For example, in one particular loop, the temperature factors are above 70 Å2 for the α carbons, indicating larger than 1 Å mean displacement value for these atoms. The corresponding

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region in the HIV-1 structure is the uninterpretable stretch from residues 140 to 153, showing that beyond the three-dimensional similarity, the molecules also share a similar disorder pattern despite their different crystal packing interactions. It is clear that in the apoenzyme (metal-free) form of the HIV-1 integrase core, disorder is present in parts of the active site. However, in the holo form, structural stability must be necessary to form a metal-binding site. Position of the Third Essential Carboxylate Does the structure of ASV integrase give us a hint about the likely conformation of the polypeptide chain around Glu152 in the holoenzyme form of the HIV-1 integrase core? The answer is probably yes, considering the overall similarity of the structures. The first residue after the disordered part in the HIV1 integrase core is Met154, which is also the first residue in helix C. The corresponding helix in the ASV core is longer, running between Gln153 and Gly175. The residue analogous to Glu152 is Glu157 in the ASV integrase core structure, located a half-turn upstream from Ala159, which corresponds to Met154 in the HIV-1 integrase structure. It is plausible to assume, therefore, that the polypeptide chain in the holoenzyme form of HIV-1 integrase would also be in a helical conformation around Glu152, and its location would be very close to the one that can be inferred from the location of Glu157 in the ASV integrase core. Secondary structure prediction also supports this assumption, assigning α-helical structure around Glu152. Why does this helical turn show significant disorder in the HIV-1 integrase structure? The answer might be found in the amino acid sequence: Pro145 is a highly conserved residue among retroviral integrases, the only exception being ASV integrase where it is substituted with a Ser. Since the main chain nitrogen of a proline is not capable of participating in hydrogen bonding, it is very rarely found in helices. It is likely that if the polypeptide chain around Glu152 were helical in the holoenzyme form of the HIV-1 integrase core, then this helix would start after Pro145. There is no such restriction in the ASV integrase core, and it is possible that this is why helix C is longer in ASV than in HIV. This may also explain the disorder around Glu152 in the HIV-1 integrase core, since it is closer to the end of the helix and more susceptible to disordering effects. A longer helix and therefore a more ordered active site in the apo form may be a unique feature of the ASV integrase core. B. Similarity to Other Polynucleotidyl Transferases Overall Protein Folds The catalytic core domain of HIV-1 integrase has a topologically identical fold with the RNase H domain of HIV-1 reverse transcriptase [37], the RuvC Holli-

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Figure 7 Structures of the catalytic core of HIV-1 integrase, HIV-1 RNase H, RuvC, and the core domain of MuA transposase demonstrating similarities in folding topology. The catalytically essential residues are highlighted.

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day junction resolving enzyme [38], and also the core domain of the phage MuA transposase [39] (see Figure 7). In the case of HIV-1 integrase, RNase H, and RuvC, definite three-dimensional similarity extends only to the ends of the last β strand; from this point, the structures diverge. In HIV-1 RNase H, there is only one more α helix corresponding to helix D in the HIV-1 integrase core but in a 40° different orientation. In RuvC there are three more helices, with the last one running parallel to helix D of HIV-1 integrase, but in the opposite direction and also 4.6 Å closer to the β sheet. In contrast, the homology between HIV-1 inte-

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grase and the MuA transposase extends until the carboxyl termini of their respective catalytic domains with three very similarly positioned and oriented α helices. Limited three-dimensional alignment between the four molecules can be accomplished by identifying structurally homologous stretches along the polypeptide chains if this search is restricted to between the first well-ordered amino terminal residue and the end of the last β strand. For the core domain of HIV-1 integrase, this corresponds to the region between Ile60 and Gln137. Using the corresponding region in the MuA transposase, the two structures can be aligned with an rms deviation of 1.7 Å over 69 α-carbon positions. The main differences between these structures are two insertions in the transposase core: an 11-residue β-stranded extension replacing the turn between the first and the second strand in the HIV-1 integrase core, and a 15-residue extension before helix B with no secondary structure. Both of these extensions interact with the downstream nonspecific DNA-binding domain of the transposase. For HIV1 RNase H, the alignment results in a rms deviation of 2.0 Å over 48 alignable α-carbon positions. The position of helix A is significantly different in RNase H, as it shifts more than 5 Å toward the β sheet. There is also an additional 2.5 turn helix following the fourth β strand and a 5-residue loop after this helix. For RuvC the alignment yields an rms deviation of 2.0 Å over 50 alignable α-carbon positions. In this case, the differences are mostly the result of longer secondary structure elements in RuvC. Of all the molecules compared, the HIV-1 integrase core is the smallest, with the most compact design in the region where these alignments were performed. For comparison, let us mention again that the homologous ASV integrase core can be aligned with an rms deviation of 1.4 Å over 74 α-carbon positions in this region. Both topological similarity and three-dimensional homology with the MuA transposase was expected based on the similarity of the reactions the enzymes catalyze, but the relationship with RNase H and RuvC was a surprise. This discovery led to the proposal of a new polynucleotidyl transferase superfamily. All the members of the superfamily are divalent metal ion-dependent endonucleases, and they all leave 3'-OH and 5'-phosphate groups at the site of cleavage. All the members of the superfamily display their catalytically essential acidic residues at the same general location. There are three such residues in HIV-1 integrase, RNase H, and the MuA transposase, while there are four in RuvC. Two of these residues are always located on the same three-dimensional structural elements, while the location of the third varies. The Asp64 residue HIV-1 integrase corresponds to Asp443 in HIV-1 RNaseH, Asp7 in RuvC, and Asp269 in the MuA transposase. Based on the three-dimensionally aligned structures, the α-carbon positions of these residues cluster quite well around that of HIV-1 integrase, with an rms deviation of 0.84 Å. All these residues are located in the middle of first β strand. The side chain torsion angle, Chi 1, is

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-62° for HIV-1 integrase, a frequently observed rotamer. For the other three molecules this Chi 1 value varies between -142° and -156°, a common range for rotamer angles. The preference for the first rotamer of the HIV side chain is probably due to the 2.73 Å-long hydrogen bond between Oο°2 of Asp64 and Nε2 of Gln62. There is no such interaction in the other molecules. The different rotamer causes a 2.46 Å rms separation of the Cγ positions around the HIV-1 Cγ, compared to only 1.63 Å around the Cγ of Asp443 in HIV-1 RNase H, indicting the carboxylate of Asp64 in HIV-1 integrase as the outlier. Position of the Second Carboxylate Residue In all the analogous structures, the second essential carboxylate resides just after the end of the fourth β strand. The main-chain atoms are not involved in strand-forming direct hydrogen bonds, therefore the chain diverts from running parallel with the first strand, forming a small cleft. The equivalent residues are Asp116 in HIV-1, Asp498 in HIV-1 RNase H, and Glu66 in RuvC. The clustering is weaker than for Asp64; the rms deviation is 1.77 Å in α-carbon position around the HIV-1 integrase residue. Interestingly, by including the structurally otherwise highly homologous ASV integrase core, the rms deviation increases to 2.15 Å due to the 3 Å distance between the Cα of Asp116 of HIV-1 integrase and that of the corresponding residue, Asp121 of ASV integrase. The rms separation between the ASV position and the rest of the cluster (now excluding HIV-1 integrase) is 2.2 Å, which is rather high, identifying the ASV residue as the outlier. For the Chi 1 torsion angles, all three preferred rotamers are present: Chi 1 is -86° for HIV-1 integrase, 73° for HIV-1 RNase H, and -173° for the MuA transposase. The RuvC Chi 1 value is not included in this comparison because it has a Glu in this position. The different Chi 1 values combined with the variation in α-carbon positions leads to a somewhat more scattered Cγ (or Cδ for Glu66 in RuvC) position with an rms deviation of 2.71 Å around Cγ of Asp116 of HIV-1 integrase. By including the ASV molecule, the scatter increases to 3.82 Å due to the 6 Å distance between Cγ of Asp116 in HIV-1 integrase and Cγ of Asp121 of ASV integrase. The Third Essential Carboxylate The location of the third essential catalytic carboxylate varies between different members of the superfamily. For HIV-1 integrase, Glu152 is in a disordered region with no interpretable electron density. Based on the location of the equivalent residue in the ASV integrase, its position is assumed to be on helix D, as discussed above. For RNase H, Glu478 is located on helix A, with its side chain pointing toward the other two carboxylates to complete the divalentmetal-binding site. Such metal binding has been observed crystallographically

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[37]. For RuvC, four residues have been shown to be essential for catalysis [40]. The third of these, Asp138, is at the amino end of the last helix of the structure. In the three-dimensionally aligned structures, this helix is parallel with the helix D in HIV-1 integrase, although it is running in the opposite direction. The position of helix D in RuvC is also significantly different, mainly due to a 13 Å shift along its axis toward the active site, placing Asp138 close to the other two carboxylates. The fourth essential residue, Asp141, is on the first turn of the same helix, very close in the aligned structures to the essential Glu157 of ASV integrase, their α carbons separated by only 1.6 Å. For the MuA transposase, its third essential carboxylate, Glu392, is in a loop region, just one residue upstream from the amino end of a helix, the topological equivalent of helix D. Unexpectedly, this residue turns away from the region defined by the two other carboxylates to a position where it clearly cannot contribute to the formation of a metal-binding site. It is likely, therefore, that the conformation of the polypeptide chain around Glu392 in the transposase core observed in the crystal structure belongs to an inactive form. In this case, a conformational change upon transposase tetramer assembly or perhaps upon substrate binding is required for activity. Significance of the Disordered Region From the point of view of HIV-1 integrase, it is interesting to note that the apparently flexible part of the MuA transposase structure is topologically equivalent with the disordered and uninterpretable part of the integrase. Similarly, in the crystal structure of the isolated RNase H domain of HIV-1 reverse transcriptase, a five-residue loop in a topologically equivalent location is disordered and therefore uninterpretable. In the ASV integrase core, the corresponding loop is visible but with rather high mobility. It seems that some kind of disorder or flexibility in this region is a common feature of the superfamily. Crystal structures of enzyme-substrate or enzyme-inhibitor complexes will tell us the functional significance of this flexibility as well as the exact configuration of the active site. C. The Dimer Interface HIV-1 integrase is active as a multimer, and the catalytic core domain alone forms dimers in solution, even at low protein concentration (see Section II.C). In the crystal structure, a roughly spherical dimer of about 45 Å diameter was observed, formed by a crystallographic two-fold axis. The dimer has a large solvent-excluded surface of 1300 Å2 per monomer. This area is close to what is expected for dimers in this molecular weight range [41]. Therefore, we are convinced that in the crystal structure the authentic dimer is present. This was subsequently confirmed by the structure of the ASV integrase core. Although

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crystallized under different conditions, forming crystals that are in a different space group with different crystal-packing interactions, the dimer observed for the ASV core is essentially identical with that of HIV-1 integrase, although the solvent-excluded surface is smaller (only 740 Å2). This difference is largely due to the absence of helix F in the ASV structure. The core domain dimer of HIV-1 integrase is held together by several hydrophobic and polar interactions. Hydrophobic interactions dominate the interface between helix E from one monomer and helices A and B from the other. There is a buried salt bridge between Glu87 of the third β strand and Lys103 on helix A. There are also some water-mediated polar interactions between these two secondary structure elements. There are direct hydrogen bonds between residues on helix A and residues on helix E across the interface including one between Lys185 (the substitution responsible for the improved solubility and therefore crystallizability) and the main-chain carbonyl on Ala105. In the ASV core, His198 is in this position, forming a very similar hydrogen bond with the carbonyl oxygen of Ala110. There are about 10 water molecules buried in the interface, all involved in hydrogen bonds. The part of the solvent-accessible surface of the monomer which becomes buried upon dimer formation displays a high degree of shape compatibility with itself: by rotating it 180° around the crystallographic two-fold axis, the resulting surface will fit the original one without forming large pockets. It is possible that the core domain of HIV-1 integrase has evolved to optimize this compatibility in order to increase its stability. It would be interesting to see the effect on protein activity of site-directed mutations aimed at disrupting this interface and hence the dimer (or possibly the higher order multimers in the context of the full-length protein). D. Implications of Crystallographic Dimer for the Chemistry of Catalysis The nearly spherical nature of the dimer formed by two monomers of the integrase catalytic core places active sites on respective monomers on opposite sides of the dimer: approximately 35 Å separates the carboxylate oxygens of Asp64 of each monomer. While we are convinced that the observed dimer is not an artifact or consequence of crystallization, it would seem difficult to reconcile this distance with the observation that, during in vivo strand transfer, cuts on the target DNA occur with a separation of five base pairs, corresponding to 15–20 Å in B-form DNA. How can a single dimer accomplish this? One possibility is that the cuts do not occur simultaneously. One end of the viral DNA could be joined by a reaction at one active site, followed by carefully controlled movement of DNA and protein relative to one another such that the second active site is now positioned five base pairs away from the initial site of strand transfer. It has been proposed, in a variation on this theme, that the first strand-transfer

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reaction is followed by DNA relaxation (unwinding, rotation, etc.), resulting in the site on the target DNA for the second transfer reaction site now being located close to the active site of the second monomer [42]. An alternate possibility is that a multimer larger than a dimer is responsible for the coordinated cutting and strand-transfer reactions. For example, two contacting dimers can be modeled such that two active sites of the resulting tetramer are located 15–20 Å apart. It is also possible that even higher order multimers are involved. There is, as yet, no convincing evidence in support of any one model. The observation that RSV integrase cuts target DNA with a six-base-pair stagger rather than the five observed for HIV correlates intriguingly with the apparently longer distance (~ 38 Å vs. 35 Å) between active sites in the RSV dimer. However, understanding the coordinated cutting and joining reaction awaits three-dimensional information on the arrangement of monomers within an integrase multimer binding to DNA. E. Three-Dimensional Structures of Other Domains of HIV-1 Integrase Three-dimensional structural information has not yet been obtained for a full-length integrase protein. In its absence, attempts have been made to determine the structure of the smaller domains consisting of the separately expressed N- and C-termini that flank the core whose structure is now known. The Amino Terminus of Integrase While the N-terminus of HIV-1 integrase, consisting of residues 1 to 55, has been separately expressed, purified, and biophysically characterized [31], structural data has not yet been obtained. This protein domain binds metal ions such as Zn2+, Co2+, and Cd2+ stoichiometrically, and is monomeric at low protein concentrations. Dramatic changes in helix content (from 14% to 32%) are observed in the circular dichroism (CD) spectrum upon addition of metal. Analysis of CD spectral features led researchers to conclude that it is highly probably that integrase contains a zinc finger that folds in much the same way as the TFIIIA-like DNA binding proteins, with two His residues located on an α helix and two cysteines part of a β sheet [31]. However, confirmation of such a model awaits structure determination by x-ray crystallography or NMR spectroscopy. The Carboxy Terminus of Integrase When the C-terminal domain is expressed as a separate polypeptide, IN213–288 can be purified from the initial soluble fraction from cell lysates [33]. This small protein fragment, therefore, was an attractive target for structure determination.

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Figure 8 Molscript stereo figure of the structure of the nonspecific DNA-binding domain of HIV-1 integrase, IN220–270, determined by heteronuclear NMR spectroscopy [28].

The structure of this domain is of particular interest as it represents the dominant nonspecific DNAbinding region of integrase. Gel filtration and sedimentation equilibrium results indicated that purified IN213–288 partitioned between dimers and highly aggregated material (unpublished observations). However, a smaller domain consisting of residues 220 to 270 maintains the DNA-binding properties of the longer C-terminal domain and is better behaved in solution. Recently, two groups have reported the structure of this smaller fragment, IN220–270, determined using multidimensional heteronuclear NMR spectroscopic methods [28,29]. As shown in Figure 8, the overall structure of IN220–270, is that of a β sandwich formed by two threestranded β sheets. As anticipated by biophysical studies, the polypeptide is a dimer in solution. The interface between monomers is formed by the antiparallel interaction of three β strands from each subunit and is stabilized predominantly by hydrophobic interactions. There is a long loop between strands β1 and β2 which, in the dimer, defines the sides of a cleft that is of the appropriate dimensions (about 24 × 24 × 12 Å) to accommodate double-stranded DNA. The folding topology is very similar to that of SH3 domains that are found in several proteins involved in signal transduction, despite the lack of significant sequence homology. This is rather unusual since SH3 domains are generally involved in protein binding rather than interactions with DNA. V. Prospects for Inhibitors A. Overview of Inhibitor Studies to Date The investigation of HIV integrase inhibitors has been largely restricted to testing available compounds that inhibit other enzymes with similar substrates or

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Page 104 Table 1 Representative Inhibitors of HIV-1 Integrase and IC50 values of compounds that inhibit 3' processing and strand transfer activities of HIV-1 integrase

Compound

IC50 of 3' processing (µM)

IC50 of strand transfer (µM)

Active against IN50–212?

Reference

I aurintricarboxylic acid monomer

10–50

n.d.

n.d.

44

II cosalane

n.d.

25

n.d.

45

III DHNQ

5.7

2.5

yes

47

IV primaquine

15

3.6

n.d.

46

V CAPE

220

19

yes*

46

VI quercetin

24

14

n.d.

47

VII quercetagetin

0.8

0.1

yes

47

VIII AG1717

0.4

0.16

yes

50

IX β-conidendrol

0.5

0.5

n.d.

51

X suramin

0.25

0.11

n.d.

53

XI curcumin

95

40

yes

55

XII (neocuproine)2-Cu+

3

3

yes

56

XIII AZT-monoPi

100–150

100–150

yes

57

XIV GT 17-mer

0.092

0.046

n.d.

59

XV HCKFWW

2

2

yes

43

The third column indicates if these compounds also inhibit disintegration by IN50–212. n.d. = not determined.* = only if pre-incubated.

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proposed mechanisms. For example, as will be seen, many topoisomerase II inhibitors also inhibit HIV integrase. While screening of chemical databases is likely underway at several pharmaceutical companies, results have either not been made available or are discouraging (see below). One foray into integrase inhibitor design—rather than discovery—has recently been described using a peptide combinatorial library approach [43]. We summarize below published reports to date (March 1996) in which compounds have been identified that inhibit integrase in in vitro assays with IC50 values of 100 µM or less (IC50 is the concentration at which the measured activity is inhibited by 50%). In vitro inhibition data is compiled in Table 1; structures of selected compounds are shown in Figure 9. An Effective Pharmacophore: Multiple Hydroxyl Groups on Aromatic Rings The first report of a class of compounds that inhibits HIV integrase appeared in 1992 [44]. Aurintricarboxylic acid (I) and its derivatives, known to inhibit other enzymes that process nucleic acids, were determined to inhibit 3' processing with moderate IC50 values of 10–50 µM. As shown in Figure 9, a recurring structural theme was established early on in which integrase inhibitors often possess aromatic rings with multiple hydroxy substituents that are either located

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Figure 9 Chemical structures of reported inhibitors of HIV-1 integrase. (I) aurintricarboxylic acid monomer; (II) cosalane; (III) dihydronaphthoquinone or DHNQ; (IV) primaquine; (V) caffeic acid phenethyl ester or CAPE; (VI) quercetin; (VII) quercetagetin; (VIII) AG1717; (IX) β-conidendrol; (X) suramin; (XI) curcumin.

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on the same ring or can potentially be positioned close together in three-dimensional space if rings stack on top of each other. More recently it was shown that cosalane (II), a steroid-substituted derivative of (I), was no better in inhibiting integrase in vitro than the parent compound, but showed promise in HIV cytopathicity cell-culture assays [45]. Although cosalane and a number of related analogues inhibit both HIV protease and integrase in vitro, the primary site of action is believed to be inhibition of gp120 binding to CD4 receptors. In 1993, the effects of selected topoisomerase II inhibitors, antimalarial agents, DNA binders, naphthoquinones, and various other agents on integrase activity in vitro were investigated [46]. Although certain effective topoisomerase inhibitors are also good HIV integrase inhibitors, this is not a generalizable correlation. Since many topoisomerase inhibitors also bind DNA, it is difficult to assess whether the observed in vitro effects result from specific interaction with integrase or from the sequestering or distorting of the DNA substrate. However, several compounds were identified that are not known to be DNA binders but that inhibited integrase with reasonable IC50 values. These included dihydroxynaphthoquinone (III), primaquine (IV), caffeic acid phenethyl ester (CAPE, V), and quercetin (VI). Motivated by the structural similarities between compounds III–VI, a more intensive structure-function study of flavones was undertaken in which approximately 50 related compounds were screened for inhibition of in vitro integrase activity [47]. Flavones are planar compounds containing three aromatic rings substituted with various polar groups such as hydroxy substituents. General trends were observed relating structure to inhibitory effectiveness; for example, inhibition required at least three hydroxy groups, the most favorable arrangement being when they were located ortho to one another. The most effective compound, quercetagetin (VII), is a potent topoisomerase II inhibitor and a known DNA intercalator. It was noted by the authors that many flavones are not integrase-specific; rather, they inhibit a broad range of enzymes including DNA polymerases, ATPases, and NAPDH-monooxygenases. They are also, in general, capable of DNA intercalation. It has not been established that their inhibitory effects are due to direct interactions with integrase. A subsequent detailed structure-activity relationship study of CAPE (V) revealed that while the ortho hydroxys were important for in vitro integrase inhibition, both the caffeic acid and phenethyl moieties could be substantially modified [48]. Ortho hydroxyl groups in the context of other classes of compounds also confer anti-integrase potency. For example, several semisynthetic compounds derived from arctigenin, a topoisomerase I inhibitor that itself is not active against integrase, have been shown to inhibit HIV integrase [49]. More compelling evidence of the importance of ortho hydroxyl groups was provided by the tyrphostins, a group of synthetic compounds that are tyrosine kinase inhibitors. Several of these also inhibit integrase in the submicromolar range

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(for example, compound VIII, aka AG1717). In cell-based screening assays, AG1717 demonstrated some antiviral activity [50]. Another study that demonstrated a role for ortho hydroxyl groups in in vitro integrase inhibition identified β-conidendrol (IX) via random screening as an inhibitor with an IC50 value of less than 1 µM [51]. Although β-conidendrol did not inhibit several other nucleic-acid processing enzymes, indicating some specificity for integrase, it was not active in cell-based antiviral assays at concentrations as high as 100 µM. Other Classes of Integrase Inhibitors Several compounds and their derivatives that do not contain adjacent hydroxy groups on a phenyl ring have recently been identified as HIV integrase inhibitors. These include suramin (X), curcumin (XI), phenanthroline-Cu+ complexes (XII), and 3'-azido-3'-deoxythymidylate (AZT) monophosphate (XIII). Suramin (X) is a known inhibitor of DNA and RNA polymerases, retroviral reverse transcriptases, and topoisomerase II. It has also been shown to prevent the infection of T lymphocytes by HIV in vitro [52]. Its six sulfonic acid groups confer a strong negative charge, and it was reasoned that there might be an inhibitory electrostatic interaction with the positive residues of the HIV C-terminus domain. Suramin was shown to be an effective inhibitor of 3' processing and strand transfer, with IC50 values of 0.25 µM and 0.11 µM, respectively [53]. It was not demonstrated, however, that the mechanism of inhibition does involve binding to the C-terminus of integrase, although this could be readily addressed using Cterminal truncated mutants active for disintegration. Curcumin (XI), the coloring dye in the spice turmeric, is structurally related to CAPE (V). It has also been shown to inhibit HIV replication by inhibiting p24 antigen production and tat-mediated transcription [54]. As shown in Table 1, it also has moderate integrase inhibitory properties [55]. Although its twoOH groups are neither adjacent to each other nor on the same phenyl ring, its conformations can be modeled to bring the hydroxy groups into close proximity by stacking the two phenyl rings on each other. Several tetrahedral cuprous phenanthroline complexes, known inhibitors of transcription, were tested against integrase and shown to be reasonably effective inhibitors [56]: IC50 values in the range of 1–10 µM were determined (for example, the neocuproine-Cu+ complex, XII). Analyses of the mode of inhibition demonstrated that these compounds act noncompetitively, and that inhibition does not correlate with inhibition of DNA binding. Thus, it has been proposed that these metal chelates may act at a site distant from the active site, or perhaps in the context of an enzyme-DNA complex. 3'-Azido-3'-deoxythymidine, or AZT, is a nucleoside analog approved for use to treat AIDS. Its metabolites, the mono-, di- and triphosphate forms, accu-

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mulate during treatment; in particular, AZT-monoPi (XIII) accumulates in cells to millimolar levels. These metabolites were investigated as possible integrase inhibitors and it was shown that all three phosphate derivatives inhibit with IC50 values of 100–150 µM, although AZT itself is not inhibitory [57]. These results suggest that despite the weak inhibition by these particular compounds, nucleoside analogs may serve as lead compounds for the development of integrase inhibitors. It was recently observed that oligonucleotides that form guanosine quartet structures inhibit HIV replication [58]. In light of the AZT results that suggested that there may be a nucleotide binding site on integrase—and because integrase binds DNA—these oligonucleotides (for example, 5'GTGGTGGGTGGGTGGGT-3', XIV) were investigated as possible inhibitors [59]. These compounds have the lowest inhibition constants reported to date (see Table 1), and suggest an exciting new avenue for integrase inhibitor development. In a completely different approach to integrase inhibitors, a synthetic peptide combinatorial library was used to select a hexapeptide capable of inhibiting integrase proteins [43]. The first two amino acids were selected using a library of 400 dipeptides, and the remaining amino acids selected one-by-one in an iterative process. The optimal hexapeptide, HCKFWW (XV), inhibits HIV-1 integrase with an IC50 of 2 µM. The peptide does not compete for DNA binding to viral DNA, nor does it represent a sequence present in integrase itself. Although it is not expected that a peptide consisting of L-amino acids would be a suitable drug in itself, the use of D-amino acids or peptidomimetic backbones may be fruitful directions to pursue. Summary Many of the compounds identified to date that inhibit HIV integrase in vitro have common structural features as illustrated in Figure 9. Most notably, these include hydroxy-substitued phenyl rings. However, these compounds may inhibit in vitro activity for reasons unrelated to enzyme binding. For example, it is difficult to know what to make of inhibition studies where the compounds added to the assays are known to bind DNA. Do the compounds affect in vitro activity because they bind and sequester the substrate? It is possible that they distort the DNA or intercalate between base pairs, preventing appropriate binding to the enzyme. While this is itself is a valid basis for the design of inhibitors against HIV infection, particularly if it targets DNA specific to the virus such as the LTR sequences [60], it is not an approach that builds on knowledge of the three-dimensional structure of the protein. To this end, valuable information could be obtained from studies in which direct binding of compounds to integrase is measured. This is also true for those inhibitors that do not contain the hydroxysubstituted phenyl pharmacophore. Binding studies could be carried

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out, for example, using radiolabeled inhibitors or possibly by monitoring any UV-spectral shifts in the case of aromatic compounds. B. What We Need to Know before We can Start Structure-Based Drug Design The review of known integrase inhibitors presented in the previous section demonstrates the paucity of effective inhibitors reported in the literature. A limited number of structural types have been investigated, focused heavily on compounds with aromatic ring systems and hydroxy substituents. Most inhibitors reported to date are only moderately effective in in vitro assays, with IC50 values residing in the low µM range (see Table 1). To build on this knowledge base, and to use known molecules as lead compounds for the development of more effective inhibitors, it would be extremely valuable to obtain a high-resolution crystal structure of an integrase-inhibitor complex. We and others are vigorously pursuing this goal. It is not clear if the lack of success to date is because the inhibitors identified so far manifest their effects in in vitro assays predominantly at the level of the DNA, or if some aspect of the structure of the HIV-1 integrase core itself—for example, its mobility in certain regoins—prevents the formation of a tight complex. It is also possible that binding is weaker to the HIV-1 core domain than to the full-length protein because of missing enzyme-inhibitor contacts. Co-crystallization attempts would benefit from in vitro studies to determine relative binding constants as a guide in selecting the most tightly bound inhibitors. It would also be useful to obtain information on the effect of variables, such as Mn2+ or Mg2+, on binding constants. It may be futile at this stage to attempt to model the binding of known inhibitors to the catalytic core domain of HIV-1 integrase in the absence of more complete information. It cannot a priori be assumed that the site of action of all these inhibitors is the enzyme active site identified by the constellation of conserved acidic residues. For example, certain very effective nonnucleoside inhibitors of HIV reverse transcriptase bind not to the enzyme active site, but rather to a small pocket adjacent to it. There are no obvious structural features of the integrase core—such as the deep trough surrounding active site residues in the case of HIV protease [61,62]—that can be readily identified as a potential inhibitor binding site. Furthermore, since part of the region defining the active site of HIV-1 integrase is disordered in our crystal structure, this prevents a surface rendering of the region around the conserved acidic residues. It also seems unlikely, given the known differences in active-site geometries between the HIV and ASV integrase (see Section IV.B), that the structure of the related ASV integrase core domain by itself would be particularly useful in this regard. As the active form of the enzyme presumably binds divalent metal ions, it will be important to determine how or if the structure of HIV integrase changes

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when metals are bound. Finally, there is no evidence to rule out the possibility that the two termini contribute to part of the active site, occlude some of it, or restrict access to it in as yet undetermined ways. For this and other obvious reasons, three-dimensional structures of larger versions of HIV-1 integrase, such as IN1–212, IN50–288, and the full-length protein, IN1–288, will be required. We also lack a clear picture of how the enzyme substrates, the viral DNA ends and the target DNA, bind to integrase. The DNA must at some point approach the region defined by the three conserved acidic residues so that bond cleavage and joining can occur. However, the dominant DNA binding domain is defined by residues in the C-terminus. It would be extremely valuable to determine the relative orientation of these domains in the context of a larger version of integrase. Even more revealing would be the structure of the full-length protein with bound DNA. Once we possess this information, it should be possible to rapidly progress with structure-based drug design. C. Possible Approaches to the Design of Effective Integrase Inhibitors There are a variety of approaches to the design of integrase inhibitors that are obvious and do not depend on knowing its three-dimensional structure. However, the rational implementation and refinement of these approaches will require high-resolutional structural data, much of which, as indicated above, is not yet available. It still may be useful to discuss here different classes of inhibitors that can be envisioned. Preventing DNA Binding One approach to the inhibition of integrase would be to prevent binding of the DNA substrate. Unfortunately, we do not yet know how or where DNA binds. There are likely to be several sites on the enzyme that contact DNA, including the C-terminus and the region around the active site. In the absence of the structure of an integrase-DNA complex, structures of related enzymes (RNase H, the MuA transposase, and RuvC) with their DNA substrates would be useful guides in suggesting ways in which DNA could interact with integrase. However, there is no guarantee that modes of DNA binding are conserved among members of this polynucleotidyl transferase family. The overall structure of the Cterminus fragment suggests that it should be possible to develop compounds that bind specifically in the cleft formed by dimers of IN220–270 (see Section IV.E) which may prevent DNA binding. Inhibition at the Active Site It may be possible to inhibit integrase by preventing the binding of the required metal cofactor(s) to the active site acidic residues that are presumed to provide

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coordinating ligands to the metal(s). This could be accomplished by sterically blocking access to the active site or by specifically binding the acidic residues themselves. Such a mechanism has been suggested to explain the inhibition of integrase by curcumin [55], which could bind to the active site aspartates or glutamates via its hydroxy groups. Compounds could also be devised that chelate the metal(s) once bound, preventing access of the active site to the substrate or distorting the active site geometry preventing phosphate bond cleavage. An intriguing approach to disrupting metal binding has recently been reported for the Zn2+-binding HIV nucleocapsid (NC) protein [63]. In this case, compounds were developed that specifically eject Zn2+ from the zinc-finger region of NC, interfering with viral replication. Such an approach might be applied to Mn2+ or Mg2+ binding at the active site, although the Zn2+-binding domain at the N terminus of integrase also suggests itself as a target. It is curious that approaches have not been devised for mechanism-based inhibitors, particularly since the stereochemical mechanism of integration has been understood for some time now [8]. Interfering with Multimerization The structures of the domains of HIV-1 integrase determined to date both reveal dimers [3,28,29,36]. It may be possible to develop compounds that bind specifically to the dimer interfaces, preventing interactions between monomers that may be necessary for activity. The success of this approach presumes that during the retroviral lifecycle there is a point where the monomer surfaces are accessible. It is not clear that integrase in the context of preintegration complexes is in equilibrium between the monomeric and higher order forms. This approach need not be restricted to preventing dimer formation if higher order interactions (e.g. formation of a tetramer) are also mechanistically relevant. To this end, the structure of an integrase tetramer, such as that formed by the full-length protein, would be useful in identifying dimer-dimer interface(s). Other Ways to Confound Integrase There are other parts of the retroviral life cycle involving integrase that could be targeted for inhibition. For example, integrase can bind other proteins such as human Ini1 [64], and most likely interacts with the viral proteins that are part of the preintegration complex [65]. Although it is not known if these interactions are essential for viral replication, preventing protein-protein binding would provide another site at which to attack integrase. It is possible that interactions between integrase and other proteins in the preintegration complex are crucial for maintaining the integrity of this complex and its ability to migrate to the cell nucleus. Interfering with these protein-protein or protein-nucleic acid interactions may be another approach to halting viral replication. For example, prein-

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cubation of phosphotyrosine with integrase has been shown to inhibit interaction with the MA protein [65]. Therefore, phosphotyrosine analogs could be a unique approach to antiviral development. D. Concluding Statement It is known from drug design studies with HIV reverse transcriptase and protease that the virus is able to escape from the pressures of inhibitors by mutation of the drug targets [66]. Although integrase is, a priori, a reasonable target for drug-design efforts, it must be anticipated that integrase will also be able to rapidly mutate and thereby avoid inhibition. By analogy with recent approaches to reverse transcriptase inhibitors, it may be possible to design a set of integrase inhibitors that act at slightly different binding sites and from which the virus cannot simultaneously escape. That is, the combination of mutations required to avoid inhibition may be severe enough to prevent integrase from carrying out its required chemistry. (In the absence of direct structural information on the sites of inhibitor binding to integrase, the generation of escape mutants in vitro and their subsequent sequencing may be an indirect way to identify inhibitor binding sites.) It will also be important to determine if the virus will be able to simultaneously mutate reverse transcriptase, integrase, and the protease in response to a combination of inhibitors targeted against all three pol gene products. The answers to these questions will require the development of suitable inhibitors and the beginning of in vitro testing. To this end—while large-scale screening and the development of combinatorial chemistry methods should continue—the structure of the catalytic core domain of HIV-1 integrase is a starting point for the rational design of integrase inhibitors. There is much more structural information that must be obtained for the full-length protein and its complexes with metals, inhibitors, and substrates. We and others are aggressively pursuing results on these fronts. E. Recent Developments Several studies published since March 1996 have expanded the list of in vitro integrase inhibitors effective at IC50 values below 100 µM. These include two dicaffeoylquinic acids obtained from medicinal plants and a synthetic analog, L-chicoric acid [68], the HIV protease inhibitors NSC 117027 and NSC 158393 [69], certain anthraquinone derivatives [70], coumermycin, and pyridoxal phosphate [71]. In addition to exhibiting in vitro inhibition, the dicaffeoylquinic acids effectively inhibited HIV-1 replication in T-lymphoblastoid cell lines [68].

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Follow-up studies were also reported for two previously identified classes of integrase inhibitors. Several nucleotides that were more effective inhibitors than the originally tested AZT nucleotides were identified [71]. For example, the L-enantiomers of 5-fluoro-2',3'-dideoxycytidine monophosphate and triphosphate inhibit 3' processing and strand transfer with IC50 values of ~40 µM. A structure-function study on GT-containing oligonucleotides showed that both the number of quartets formed and the loop sequences between the quartets are important for activity, and that inhibitors of this type may function by interacting with the N-terminus of integrase [72]. A particularly important contribution was the demonstration that preintegration complexes isolated from HIV-infected lymphoid cells can be used in assays to screen for inhibition of integration [70]. Intriguingly, many compounds previously identified as in vitro inhibitors of 3' processing and strand transfer had no effect on integration carried out by either crude or partially purified preintegration complexes. Thus, such an assay may be a valuable method of screening out “false positives” identified using in vitro oligonucleotide assays, or corroborating the evidence that a particular compound is indeed active against integrase. Acknowledgments Our work described here was carried out in the laboratories of R. Craigie and D. R. Davies. We express our gratitude to our co-workers who, over the years, participated in the effort to determine the structure of HIV integrase. In particular, we acknowledge the contributions of F. D.Bushman, M. Carmichael, A. Engelman, S. Hosseini, T. Jenkins, K. Mizuuchi, I. Palmer, P. Rice, P. Sun, and P. Wingfield. We would also like to thank D. R. Davies and T. Jenkins for their comments on the manuscript and A. Mazumder for alerting us to the most recent work on integrase inhibitors and for his contributions to Section V.A. References 1. AIDS WEEKLY Plus. Key KK, ed. Atlanta, Charles Henderson Publisher, 1996: Feb. 5 & 12, p. 14. 2. Cara A, Guarnaccia F, Reitz, Jr. MS, Gallo RC, Lori F. Self-limiting, cell type-dependent replication of an integrase-defective human immunodeficiency virus type 1 in human primary macrophages but not T lymphocytes. Virol 1995, 208:242–248. 3. Dyda F, Hickman AB, Jenkins TM, Engelman A, Craigie R, Davies DR. Crystal structure of the catalytic domain of HIV-1 integrase: Similarity to other polynucleotidyl transferases. Science 1994; 266:1981–1986.

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4. Vink C, Plasterk RHA. The human immunodeficiency virus integrase protein. Trends Genet 1993; 9:433–437. 5. Katz RA, Skalka AM. The retroviral enzymes. Annu Rev Biochem 1994; 63:133–173. 6. Sherman PA, Fyfe JA. Human immunodeficiency virus integration protein expressed in Escherichia coli possesses selective DNA cleaving activity. Proc Natl Acad Sci USA 1990; 87:5119–5123. 7. Bushman FD, Craigie R. Activities of human immunodeficiency virus (HIV) integration protein in vitro: Specific cleavage and integration of HIV DNA. Proc Natl Acad Sci USA 1991; 88:1339–1343. 8. Engelman A, Mizuuchi K, Craigie R. HIV-1 DNA integration: Mechanism of viral DNA cleavage and DNA strand transfer. Cell 1991; 67:1211–1221. 9. Chow SA, Vincent KA, Ellison V, Brown PO. Reversal of integration and DNA slicing mediated by integrase of human immunodeficiency virus. Science 1992; 255:723–726. 10. Grandgenett DP, Vora AC, Schiff RD. A 32,000-dalton nucleic acid-binding protein from avian retravirus cores possesses DNA endonuclease activity. Virol 1978; 89:119–132. 11. Vincent KA, Ellison V, Chow SA, Brown PO. Characterization of human immunodeficiency virus type 1 integrase expressed in Escherichia coli and analysis of variants with amino-terminal mutations. J. Virol 1993; 67:425–437. 12. Hickman AB, Palmer I, Engelman A, Craigie R, Wingfield P. Biophysical and enzymatic properties of the catalytic domain of HIV-1 integrase. J Biol Chem 1994; 269:29279–29287. 13. Jones KS, Coleman J, Merkel GW, Laue TM, Skalka AM. Retroviral integrase functions as a multimer and can turn over catalytically. J Biol Chem 1992; 267:16037–16040. 14. Engelman A, Bushman FD, Craigie R. Identification of discrete functional domains of HIV-1 integrase and their organization within an active multimeric complex. EMBO J 1993; 12:3269–3275. 15. Andrake MD, Skalka AM. Multimerization determinants reside in both the catalytic core and C terminus of avian sarcoma virus integrase. J Biol Chem 1995; 270:29299–29306. 16. Kalpana GV, Goff SP. Genetic analysis of homomeric interactions of human immunodeficiency virus type 1 integrase using the yeast two-hybrid system. Proc Natl Acad Sci USA 1993; 90:10593–10597. 17. Van Gent DC, Vink C, Oude Groeneger AAM, Plasterk RHA. Complementation between HIV integrase proteins mutated in different domains. EMBO J 1993; 12:3261–3267.

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18. Johnson MS, McClure MA, Feng D–F, Gray J, Doolittle RF. Computer analysis of retroviral pol genes: Assignment of enzymatic functions to specific sequences and homologies with nonviral enzymes. Proc Natl Acad Sci USA 1986; 83:7648–7652. 19. Engelman A, Craigie R. Identification of conserved amino acid residues critical for human immunodeficiency virus type 1 integrase function in vitro. J Virol 1992; 66:6361–6369. 20. Van Gent DC, Oude Groeneger AAM, Plasterk RHA. Mutational analysis of the integrase protein of human immunodeficiency virus type 2. Proc Natl Acad Sci USA 1992; 89:9598–9602.

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21. Kulkosky J, Jones KS, Katz RA, Mack JPG, Skalka AM. Residues critical for retroviral integrative recombination in a region that is highly conserved among retroviral/retrotransposon integrases and bacterial insertion sequence transposases. Mol Cell Biol 1992; 12:2331–2338. 22. Bushman FD, Engelman A, Palmer I, Wingfield P, Craigie R. Domains of the integrase protein of human immunodeficiency virus type 1 responsible for polynucleotidyl transfer and zinc binding. Proc Natl Acad Sci USA 1993; 90:3428–3432. 23. Beese LS, Steitz TA. Structural basis for the 3'-5' exonuclease activity of Escherichia coli DNA polymerase I: a two metal ion mechanism. EMBO J 1991; 10:25–33. 24. Woerner AM, Marcus-Sekura CJ. Characterization of a DNA binding domain in the C-terminus of HIV-1 integrase by deletion mutagenesis. Nucl Acids Res 1993; 21:3507–3511. 25. Vink C, Oude Groeneger AAM, Plasterk RHA. Identification of the catalytic and DNA-binding region of the human immunodeficiency virus type 1 integrase protein. Nucl Acids Res 1993; 21:1419–1425. 26. Puras Lutzke RA, Vink C, Plasterk RHA. Characterization of the minimal DNA-binding domain of the HIV integrase protein. Nucl Acids Res 1994; 22:4125–4131. 27. Drelich M, Wilhelm R, Mous J. Identification of amino acid residues critical for endonuclease and integration activities of HIV-1 IN protein in vitro. Virol 1992; 188:459–468. 28. Lodi PJ, Ernst JA, Kuszewski J, Hickman AB, Engelman A, Craigie R, Clore GM, Gronenborn AM. Solution structure of the DNA binding domain of HIV-1 integrase. Biochem 1995; 34:9826–9833. 29. Eijkelenboom APAM, Puras Lutzke RA, Boelens R, Plasterk RHA, Kaptein R, Hard K. The DNAbinding domain of HIV-1 integrase has an SH3-like fold. Nature Struct Biol 1995; 2:807–810. 30. Haugan IR, Nilsen BM, Worland S, Olsen L, Helland DE. Characterization of the DNA-binding activity of HIV-1 integrase using a filter binding assay. Biochem Biophys Res Commun 1995; 217:802–810. 31. Burke CJ, Sanyal G, Bruner MW, Ryan JA, LaFemina RL, Robbins HL, Zeft AS, Middaugh CR, Cordingley MG. Structural implications of spectroscopic characterization of a putative zinc finger peptide from HIV-1 integrase. J Biol Chem 1992; 267:9639–9644. 32. Bushman FD, Wang B. Rous sarcoma virus integrase protein: Mapping functions for catalysis and substrate binding. J Virol 1994; 68:2215–2223. 33. Engelman A, Hickman AB, Craigie R. The core and carboxyl-terminal domains of the integrase protein of human immunodeficiency virus type 1 each contribute to nonspecific DNA binding. J Virol 1994; 68:5911–5917. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_115.html (1 of 2) [4/5/2004 4:52:31 PM]

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34. Jenkins TM, Hickman AB, Dyda F, Ghirlando R, Davies DR, Craigie R. Catalytic domain of human immunodeficiency virus type 1 integrase: Identification of a soluble mutant by systematic replacement of hydrophobic residues. Proc Natl Acad Sci USA 1995; 92:6057–6061. 35. Orengo CA, Thornton JM. Alpha plus beta fold revisited: some favoured motifs. Structure 1993; 1:105–120.

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44. Cushman M, Sherman P. Inhibition of HIV-1 integration protein by aurintricarboxylic acid monomers, monomer analogs, and polymer fractions. Biochem Biophys Res Commun 1992; 185:85–90. 45. Cushman M, Golebiewski WM, Pommier Y, Mazumder A, Reymen D, De Clercq E, Graham L, Rice WG. Cosalane analogues with enhanced potencies as inhibitors of HIV-1 protease and integrase. J Med Chem 1995; 38:443–452. 46. Fesen MR, Kohn KW, Leteurte F, Pommier Y. Inhibitors of human immunodeficiency virus integrase. Proc Natl Acad Sci USA 1993; 90:2399–2403. 47. Fesen MR, Pommier Y, Leteurtre F, Hiroguchi S, Yung J, Kohn KW. Inhibition of HIV-1 integrase by flavones, caffeic acid phenethl ester (CAPE) and related compounds. Biochem Pharmacol 1994; 48:595–608. 48. Burke Jr TR, Fesen MR, Mazumder A, Wang J, Carothers AM, Grunberger D, Driscoll J, Kohn K, Pommier Y. Hydroxylated aromatic inhibitors of HIV-1 integrase. J Med Chem 1995; 38:4171–4178. 49. Eich E, Pertz H, Kaloga M, Schulz J, Fesen MR, Mazumder A, Pommier Y. (-)-Arctigenin as a lead structure for inhibitors of human immunodeficiency virus type-1 integrase. J Med Chem 1996; 39:86–95. 50. Mazumder A, Gazit A, Levitzki A, Nicklaus M, Yung J, Kohlhagen G, Pommier Y. Effects of tyrphostins, protein kinase inhibitors, on human immunodeficiency virus type 1 integrase. Biochem 1995; 34:15111–15122. 51. LaFemina RL, Graham PL, LeGrow K, Hastings JC, Wolfe A, Young SD, Emini EA, Hazuda DJ. Inhibition of human immunodeficiency cirus integrase by bis-catechols. Antimicrob Agents Chemother 1995; 39:320–324. 52. Mitsuya H, Popovic M, Yarchoan R, Matsushita S, Gallo RC, Broder S. Suramin protection of T cells in vitro against infectivity and cytopathic effect of HTLV-III. Science 1984; 226:172–174.

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53. Carteau S, Mouscadet JF, Goulaouic H, Subra F, Auclair C. Inhibitory effect of the polyanionic drug suramin on the in vitro HIV DNA integration reaction. Arch Biochem Biophys 1993; 305:606–610. 54. Li CJ, Zhang LJ, Dezube BJ, Crumpacker CS, Pardee AB. Three inhibitors of type 1 immunodeficiency virus long terminal repeat-directed gene expression and virus replication. Proc Natl Acad Sci USA 1993; 90:1839–1842. 55. Mazumder A, Raghavan K, Weinstein J, Kohn KW, Pommier Y. Inhibition of human immunodeficiency virus type-1 integrase by curcumin. Biochem Pharmacol 1995; 49:1165–1170. 56. Mazumder A, Gupta M, Perrin DM, Sigman DS, Rabinovitz M, Pommier Y. Inhibition of human immunodeficiency virus type 1 integrase by a hydrophobic cation: The phenanthroline-cuprous complex. AIDS Res Human Retro 1995; 11:115–125. 57. Mazumder A, Cooney D, Agbaria R, Gupta M, Pommier Y. Inhibition of human immunodeficiency virus type 1 integrase by 3' -azido-3' -deoxythymidylate. Proc Natl Acad Sci USA 1994; 91:5771–5775. 58. Ojwang J, Elbaggari A, Marshall HB, Jayaraman K, McGrath MS, Rando RF. Inhibition of human immunodeficiency virus type 1 activity in vitro by oligonucleotides composed entirely of guanosine and thymidine. J Acqu Immune Defic Syn 1994; 7:560–570. 59. Ojwang JO, Buckheit RW, Pommier Y, Mazumder A, de Vreese K, Esté JA, Reymen D, Pallansch LA, Lackman-Smith C, Wallace TL, de Clercq E, McGrath MS, Rando RF. T30177, an oligonucleotide stabilized by an intramolecular guanosine octet, is a potent inhibitor of laboratory strains and clinical isolates of human immunodeficiency virus type 1. Antimicrob Agents Chemother 1995; 39:2426–2435. 60. Carteau S, Mouscadet JF, Goulaouic H, Subra F, Auclair C. Inhibition of the in vitro integration of Moloney murine leukemia virus DNA by the DNA minor groove binder netropsin. Biochem Pharmacol 1994; 47:1821–1826. 61. Navia MA, Fitzgerald PMD, McKeever BM, Leu C-T, Heimbach JC, Herber WK, Sigal IS, Darke PL, Springer JP. Three-dimensional structure of aspartyl protease from human immunodeficiency virus HIV-1. Nature 1989; 337:615–620. 62. Wlodawer A, Erickson JW. Structure-based inhibitors of HIV-1 protease. Annu Rev Biochem 1993; 62:543–585. 63. Rice WG, Supko JG, Malspeis L, Buckheit Jr RW, Clanton D, Bu M, Graham L, Schaeffer CA, Turpin JA, Domagala J, Gogliotti R, Bader JP, Halliday SM, Coren L, Sowder II RC, Arthur LO, Henderson LE. Inhibitors of HIV nucleocapsid protein zinc fingers as candidates for the treatment of AIDS. Science 1995; 270:1194–1197. 64. Kalpana GV, Marmon S, Wang W, Crabtree GR, Goff SP. Binding and stimulation of HIV-1 integrase by a human homolog of yeast transcription factor SNF5. Science 1994; 266:2002–2006. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_117.html (1 of 2) [4/5/2004 4:56:01 PM]

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65. Gallay P, Swingler S, Song J, Bushman F, Trono D. HIV nuclear import is governed by the phosphotyrosine-mediated binding of matrix to the core domain of integrase. Cell 1995; 83:569–576. 66. Coffin JM. HIV population dynamics in vivo: Implications for genetic variation, pathogenesis, and therapy. Science 1995; 267:483–489. 67. Williams KJ, Loeb LA. Retroviral reverse transcriptases: Error frequencies and mutagenesis. Curr Top Microbiol Immunol 1992; 176:165–180.

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68. Robinson Jr WE, Reinecke MG, Abdel-Malek S, Jia Q, Chow SA. Inhibitors of HIV-1 replication that inhibit HIV integrase. Proc Natl Acad Sci USA 1996; 93:6326–6331. 69. Mazumder A, Wang S, Neamati N, Nicklaus M, Sunder S, Chen J, Milne GWA, Rice WG, Burke Jr TR, Pommier Y. Antiretroviral agents as inhibitors of both human immunodeficiency virus type 1 integrase and protease. J Med Chem 1996; 39;2472–2481. 70. Farnet CM, Wang B, Lipford JR, Bushman FD. Differential inhibition of HIV-1 preintegration complexes and purified integrase protein by small molecules. Proc Natl Acad Sci USA 1996; 93:9742–9747. 71. Mazumder A, Neamati N, Sommadossi J, Gosselin G, Schinazi RF, Imbach J, Pommier Y. Effects of nucleotide analogues on human immunodeficiency virus type 1 integrase. Mol Pharmacol 1996; 49:621–628. 72. Mazumder A, Neamati N, Ojwang JO, Sunder S, Rando RF, Pommier Y. Inhibition of the human immunodeficiency virus type 1 integrase by guanosine quartet structures. Biochem 1996; 35:13762–13771.

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4 Bradykinin Receptor Antagonists Donald J. Kyle Scios Nova Inc., Sunnyvale, California I. Introduction The term “kinins” is generally made in reference to either the nonapeptide bradykinin (Arg1-Pro2-Pro3Gly4-Phe5-Ser6-Pro7-Phe8-Arg9) or the decapeptide kallidin (Lys1-Arg2-Pro3-Pro4-Gly5-Phe6-Ser7-Pro8Phe9-Arg10). In rats another kinin, Ile1-Ser2-Arg3-Pro4-Pro5-Gly6-Phe7-Ser8-Pro9-Phe10-Arg11 (T-kinin) is produced under certain circumstances and binds to the same receptors as bradykinin [1,2]. A schematic of the human kinin-kallikrein system is shown in Figure 1. The release of kinins from precursor proteins (known as kininogens) is mediated by enzymes called kininogenases [3–5]. The predominant enzymes responsible are kallikreins, but others, which include trypsin, plasmin, and some snake venoms, also release kinins. Kininogens are primarily synthesized in the liver and represent an abundant source of the precursors that are required for kinin generation. These proteins are produced from alternative splicing of a single gene product and there are two forms: high molecular weight kininogen (HMWK) and low molecular weight kininogen (LMWK) [6]. Unlike HMWK, which exists in the circulation as a complex with plasma pre-kallikrein, LMWK circulates freely. During immunological reaction, charged surfaces—which may be derived from bacterial lipopolysaccharide, oligosaccharides, connective tissue proteoglycans, or damaged basement membranes—facilitate the conversion of factor XII to factor XIIa. Once factor XIIa is present, prekallikrein can be cleaved to its active form, known as plasma kallikrein. This enzyme acts upon its preferred substrate, HMWK, to release bradykinin. Plasma kallikrein is further able to convert inactive factor XII to active XIIa, thereby participating in a positive

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Figure 1 Diagram of the human kinin-kallikrein system including the native ligands for B1 and B2 receptor subtypes.

feedback loop. The cleavage of bradykinin from HMWK is highly localized since pre-kallikrein and substrate (HMWK) circulate as a complex. Another kinin, Lys-bradykinin (also known as kallidin), is produced via the action of the tissuekallikrein enzyme on LMWK. This enzyme is found in many tissues, either in the form of a precursor requiring activation or as an active enzyme. In contrast to plasma kallikrein, which preferentially acts upon HMWK, tissue kallikrein can release kallidin from either HMWK or LMWK. Through the action of aminopeptidases, kallidin can subsequently be converted directly into bradykinin. This enzyme is present in both the plasma and on the surface of epithelial cells. Both bradykinin and kallidin can be degraded by a variety of plasma and cell surface enzymes (kininases) [7]. The most widely recognized of these enzymes are kininase I, kininase II (angiotensin converting enzyme, ACE), and carboxypeptidase N. In plasma, kininase I cleaves the C-terminal arginine from both bradykinin and kallidin to form [des-Arg9] kinins. These [des-Arg9] kinins are known to act as agonists of B1 receptors that are present in some species and have been implicated in the pathophysiology associated with prolonged inflammation [8–10].

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Nearly all cells express kinin receptors that mediate