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 the activities of both bradykinin and kallidin. The activation of these G-protein coupled receptors causes relaxation of venular smooth muscle and hypotension, increased vascular permeability, contraction of smooth muscle of the gut and airway leading to increased airway resistance, stimulation of sensory neurons, alteration of ion secretion of epithelial cells, production of nitric oxide, release of cytokines from leukocytes, and the production of eicosanoids from various cell types [11,12]. Because of this broad spectrum of activity, kinins have been implicated as an important mediator in many pathophysiologies including pain, sepsis, asthma, rheumatoid arthritis, pancreatitis, and a wide variety of other inflammatory diseases. Moreover, a recent report demonstrated that bradykinin B2 receptors on the surface of human fibroblasts were upregulated three-fold beyond normal in patients with Alzheimer's disease, implicating bradykinin as a participant in the peripheral inflammatory processes associated with that disease [13]. In contrast to the adverse physiologies associated with bradykinin release, there is a growing body of literature that implicates bradykinin as a protective agent during periods of cardiac or renal stress [14–16]. In this regard there is substantial evidence that the cardioprotective effects afforded by ACEinhibitor treatment are a result of metabolically preserving bradykinin and are therefore mediated by bradykinin B2 (and possibly B1) receptors [17–18]. These results point to a possible therapeutic role for a kinin receptor agonist. Overall, the kinins are an important part of a well-organized physiological system. The various aspects and interdependencies of the kinin system have been, and continue to be, the focus of intensive research efforts in many laboratories. Many pharmaceutical companies have identified this system as an ideal site for therapeutic intervention in many inflammatory diseases. Hence, there have been many diverse approaches taken toward the discovery of antagonists (peptide and nonpeptide) of B2 and B1 receptors. This review focuses on the structure-based design strategies pursued in our laboratories during the past several years. II. Ligand-Based Investigations A. The Solution Conformation of Bradykinin In the late 1980s when we began the pursuit of bradykinin receptor antagonists, information of relevance to medicinal chemists was scarce. For example, not one nonpeptide antagonist of this receptor was known, nor were any series upon which to base a structure-activity relationship. Moreover, all publications described bradykinin as being highly flexible in an aqueous environment, such that no structural mimetics could be rationalized. Of course the receptors had not been cloned at that time so nothing was known about the primary sequence of the receptor or the three-dimensional structure.

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Initially our approach was to complete a detailed examination of bradykinin using two-dimensional NMR methods in combination with empirical energy calculations [19]. Our strategy was derived on the basis of spectral data, biological results from conformationally restricted analogs, as well as the relationship between ordering in bradykinin and the dielectric environment of the solvent. Our guiding hypothesis was that, although in aqueous solution bradykinin is conformationally random, the biologically active form of the peptide is likely ordered and stabilized within the lipid bilayer of the cell membrane prior to binding with its receptor. Alternatively, the receptor binding environment might also be hydrophobic and thereby lead to similar conformational biases in the ligand. We presumed that an appropriate solvent environment should be able to stimulate, at least in terms of hydrophobicity and dielectric constant, the nature of a cell membrane, and a 90:10 d8-dioxane-H2O mixture was selected for NMR experiments. It was anticipated that under these nonsolvating conditions the conformational diversity of bradykinin might be severely restricted. The ultimate analysis of the two-dimensional NMR data collected at 500 MHz supported a single major conformational species. There were five HN-CαH connectivities, one for each amide. This was confirmed in the 13C NMR spectrum where only nine carbonyl resonances, one for each amino acid, were present. In order to provide a starting point for subsequent molecular dynamics simulations the assumption was made, based on multiple observed long-range amide-to-amide nuclear overhauser effects (NOEs), that it was indeed a single major conformational species. Although bradykinin contains three proline residues, the absence of any strong CαHi-CαHj+, cross peaks in the nuclear overhauser enhancement spectroscopy (NOESY) spectrum was taken as proof that all peptide bonds were trans. In total, 35 interproton distances were extracted from the NOESY spectrum and, whenever possible, stereospecific assignments for pro-R and pro-S hydrogens were made explicitly. A temperature-dependent study of the chemical shifts of the amide protons resulted in a near-linear dependence suggesting no major conformational changes were coinciding with the temperature change and thereby allowing a comparison of slopes (∆δ/∆t). The lowest values obtained for these slopes corresponded to Phe8 and Arg9 suggesting solvent sequestering for these amides. Given the high Chou and Fasman probability of β-turns in the sequences Pro2-Pro3-Gly4-Phe5 and Ser6Pro7-Phe8-Arg9 (3.79 × 10-4 and 1.99 × 10-4, respectively), the computational strategy employed was to begin from two initial structures: (a) an extended β strand, and (b) a structure containing these two predicted β turns. Utilizing custom routines written using the program CHARMm, version 21 [21], the interproton distances were incorporated into the potential-energy expression in the form of an additional potential-energy term. During the 3-ps heating step of the molecular dynamics, the temperature was raised from 0K to 300K in steps of 20K every 0.2 ps. Since the target distances

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were poorly satisfied in the starting structures, the potential-energy term corresponding to the imposed NOE data (ENOE) was applied gradually by increasing the scale factor in a nonlinear fashion such that it was 0.0 after 0.2 ps, 0.1 after 1.2 ps, and 1.0 after the full 3.0 ps. Following 15 ps of equilibration, 7 ps of incremental production dynamics was completed. During this stage the NOE scale factor was raised from 1.0 to 4.5. By slowly raising the force constants for the NOE restraints as the target distances became better satisfied, no dramatic increase in temperature was observed. Finally the NOE scale factor was set to 5.0, 10 ps of production dynamics were completed, and an average structure was extracted from the last 5 ps of the coordinate trajectory. Analysis of the two average structures obtained from the two unique starting points demonstrated convergence to a similar conformational species. In each, the sum of the NOE restraint energy was less than 4.7 kcal/mol and the RMS deviation from the target distances was below 0.25 Å. Similar results were obtained for each simulation when they were repeated without the electrostatic term being included in the total potential-energy function. This important data lends credence to the hypothesis that the final structures are derived from the NOE restraints and not by poorly represented electrostatic interactions. The average dynamic structures are characterized as having all trans peptide bonds and hydrophobic side chain groups oriented outward into solution, perhaps ready to interact with the receptor. There is a possible 1–3 hydrogen-bonded γ turn bridging Phe5, although it is not explicitly defined by the NOE data set. If present, then the preferred overall geometry would be U shaped and, if absent, an S shaped geometry is possible based on coincident conformational analyses. According to the dihedral angle values for Pro7 and Phe8, a type-II β turn extending from Ser6 to Arg9 also exists. A variety of reports have subsequently appeared that are in agreement with the conformation we described in this work. A similar C-terminal turn structure was observed in an analogous NMR study of a first-generation kinin antagonist, NPC 567 (DArg0-Arg1-Pro2-Hyp3-Gly4-Phe5-Ser6-DPhe7-Phe8-Arg9), although the type of turn was not the same. Our initial speculation was that this slight structural difference might partially account for the functional differences of bradykinin and NPC 567. These solution conformations, one of an agonist and the other of an antagonist, were subsequently used to focus the design and synthesis of conformationally constrained peptide analogues of NPC 567. B. Conformationally Constrained Bradykinin Antagonist Peptides The ligand-based approach of conformationally constrained peptides has been widely used. The process involves the incorporation of conformational constraints into known peptides, either agonist or antagonist, which enforce a

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predictable geometry. A series of peptides containing these types of constraints can be useful for extrapolating the steric and electronic environment of a given binding site. This structural information can be derived regardless of whether or not the constrained peptide binds to the target receptor. Since peptides can be prepared rapidly, it is typical to establish a structure-activity relationship using them and then at some later time transpose that information onto a nonpeptide lead molecule in an attempt to improve its potency. As part of an expansion upon the hypothesis that a C-terminal β turn was a structural prerequisite to highaffinity antagonist binding, a novel series of constrained decapeptides was prepared [22–24]. These peptides are of the sequence DArg0-Arg1-Pro2-Hyp3-Gly4-Phe5-Ser6-DHype7-Y8-Arg9, where Y is either tetrahydroisoquinoline-3-carboxylic acid (Tic), or octahydroindole-2-carboxylic acid (Oic). DHype denotes an organic ether of D-4-hydroxyproline in either the cis or trans geometric form. The C-terminal portion of a representative member of this class of peptides was shown—first by empirical calculation [22], then by NMR at 600 MHz—to adopt a β turn nearly unambiguously (figure 2) [25,26]. Moreover, it was shown by calculation that the turn was adopted regardless of the nature of the ether group (alkyl, aryl, etc.) or its geometry (cis or trans). Hence, a diverse series of these peptides was initially used as a tool to probe the steric and electrostatic topology of an antagonist

Figure 2 Lowest 5 kcal of the calculated overall potential energy surface for a model peptide of Ser-DHype(trans propyl)-Oic-Arg. The contour interval is 0.5 Kcalmol-1 and the highest (outermost) and lowest contour energy values are labeled. Superimposed on the contour plots are values for ψi+1 and ψi+2 from each of the thirty structures generated from the NMR data corresponding to the tetrapeptide Ser-DHype(trans propyl)-Oic-Arg. mol-1

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binding site on the bradykinin B2 receptor in the guinea pig ileum. The cis ethers, in all cases, bound to the receptor with significantly lower affinity than did the trans. A more complete listing of the peptides used in the study is shown in Table 1. These results support the hypothesis that the domain of the receptor that binds these antagonist ligands is partly made up of a hydrophobic cavity about one side of the C-terminal turn. However, adjacent to the other side of the

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Figure 3 Receptor binding curves for the binding of NPC 17410 and NPC 17643 to B2 receptors from the guinea pig ileum and cloned rat and human B2 receptors. Legends are noted on the figure.

turn, there appears to be some type of steric interference (or lack of a pocket) that might otherwise accommodate the ethers of the cis configuration. More recently, bradykinin B2 receptors have been cloned from both rat and human sources [27,28]. In receptor-binding experiments using these new receptors, selected members of the DHype-containing decapeptides were used to probe these receptors [24], a representative sample of the data is shown in Figure 3. Specifically, NPC 17643 (a trans propyl ether of D-4-hydroxyproline at position 7) and NPC 17410 (a cis propyl ether of D-4-hydroxyproline at position 7) were used. Although the trans ethercontaining decapeptide behaved similarly in binding assays directed toward the bradykinin B2 receptors in guinea pig, rat, and human, the cis ether-containing decapeptide, NPC 17410, displayed an interesting pharmacology. In particular, NPC 17410 bound with similar affinity to both the guinea pig and rat bradykinin B2 receptors, but had an appreciably higher affinity for the human B2 receptor. This result strongly suggests that there are slight structural differences in the antagonist binding sites of the rat and human B2 receptors. With clones available for the rat and human bradykinin B2 receptors, this prompted a systematic search using NPC 17410 binding to rat/human bradykinin B2 receptor chimeras and point mutations in an attempt to discover residues on the receptor that comprise this antagonist site. The details and results of the subsequent application of these novel receptor-probing ligands is fully described later in this chapter.

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The des-Arg9 forms of these peptides have also been shown to have high affinity for the recently cloned human B1 receptor [24]. An extension of the work described herein would be to use a more complete series of des-Arg9 DHype-containing nonapeptides to probe the binding site of this new receptor where other interesting pharmacological differences are likely to exist since the B1 receptor is only 33% homologous to the human B2 [29]. In summary, the DHype-containing decapeptides have been useful in many regards. First, they incorporate a novel β-turn mimetic that was alternatively functionalized and used to probe the unknown topology of the guinea pig, rat, and human bradykinin B2 receptors. In this role, one of these tools showed differential pharmacology between rat and human forms of the receptor. This tool was used in a synergistic fashion with subsequent molecular biological and computational procedures in the elucidation of an antagonist binding site. Second, these peptides, together with another potent decapeptide antagonist with similar conformational constraints [30,31], provide the first strong experimental evidence that high-affinity decapeptide bradykinin receptor antagonists adopt a C-terminal β turn in the receptor-bound conformation. Third, certain members of this series of decapeptides contain alkyl ethers of D-4-hydroxyproline at position seven. In this regard, they are the very first examples of decapeptide bradykinin receptor antagonists that do not contain a D-aromatic amino acid at the seventh position as had been previously deemed to be essential. Commercially, this renders the series patentably distinct from all other known bradykinin receptor antagonists. Finally, several members of the series (i.e., NPC 17731, NPC 17761, NPC 17974) are among the most potent antagonists for this receptor yet reported. Hence, there may be applications for these compounds as human therapeutics. Several “second generation” decapeptide antagonists have been reported, but the prototype from the class, which was first to be reported, is HOE 140 (DArg0-Arg1-Pro2-Hyp3-Gly4-Thi5-Ser6-DTic7-Oic8Arg9) [30,31]. This decapeptide has also been shown to preferentially adopt a C-terminal β turn, consistent with the previous discussion [26,32,33]. The side chain of DTic at position seven is, however, flexible. While side-chain rotational movement is not allowed, the saturated six-membered ring easily undergoes an endo/exo ring-flipping motion. Hence, the β turn predominates about the backbone dihedral angles, but the side chain of DTic could be either endo or exo ring flipped in the receptor-bound conformation. In the absence of an appropriate x-ray crystallographic structure, there is no definitive means of establishing which possibility is correct. This type of ring-flipping conformational change serves to orient the bulky hydrophobic side chain of the DTic residue to either one side of the β turn, or the other. The data collected from the decapeptides containing either cis or trans ethers of D-4hydroxyproline at the analogous position in the

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sequence (discussed above) support a hypothesis that the DTic is exo ring flipped in the receptor-bound state. There are two factors that must be considered when applying structure-activity-relationship (SAR) information from a series of peptides toward the design of nonpeptide mimetics and putative library scaffolds. One is in regard to the backbone conformation that primarily serves as a structural scaffold upon which the various functionalities (side chains) are attached. The other factor is the side chains themselves whose spatial positions are primarily dictated by the backbone structure. Usually, the threedimensional arrangement of these differing chemical groups are responsible for affinity and triggering of the receptor. Knowledge of the relative importance of the individual side chains and amide bonds for receptor affinity is therefore a critical aspect of small molecule design from a peptidic structure-activity relationship. Conformationally constrained derivatives of HOE 140 have been prepared in continuing efforts to elucidate the ideal backbone conformation peptide antagonists must adopt for bradykinin B2 receptor interaction. One such series made use of Cα- or N-methyl substituted amino acids, incorporated at position(s) Gly4, Phe5, or both, in the peptide D-Arg0-Arg1-Pro2-Hyp3-Gly4-Phe5-Ser6-D-Tic7-Oic8-Arg9 (NPC 18545) [34]. An N-methyl substitution in the backbone of an L-amino acid is known to disfavor helical, or twisted, backbone conformations while favoring an extended backbone. The contrasting Cαmethyl modification tends to favor a helical (twisted), rather than extended, conformation [35,36]. These conformational preferences apply only to the backbone φ, ψ angles (where φi and ψi correspond to backbone dihedral angles for residue i, defined by the four adjacent amino acid backbone atoms Ci-1-NiCαi-Ci band Ni-Cαi-Ci-Ni+1, respectively) of the amino-acid residues bearing the modification. Receptor binding assays were performed in membrane preparations of the guinea pig ileum, a source of B2 receptors, wherein these constrained peptides were evaluated for their abilities to compete with bradykinin binding. With the exception of the Cα-methyl-Phe5-containing peptide (NPC 18540), each conformational constraint caused a significant, at least 1000-fold, loss in binding affinity with respect to the unconstrained parent peptide, NPC 18545. There are several factors that could contribute to the poor receptor affinities measured for these peptides. In addition to the possible induction of an adverse conformation via the N-methyl substitution, this modification also eliminates an amide proton that might be an important hydrogen-bond donor during ligand-receptor interaction. Furthermore, the N-methyl substitution enhances the likelihood of trans-cis amide bond isomerization, which could also disrupt an optimal ligand-receptor interaction by altering the spatial display of the local side chains. The Cα-methylPhe5 substitution of NPC 18540 is well

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tolerated by the receptor as evidenced by only a seven-fold loss in receptor affinity (Ki=0.54 nM) with respect to the parent of the series, NPC 18545. This implies that the φ, ψ backbone dihedral angles about Phe5 are on the order of -60°, -60° in the biologically active conformation. This combination of dihedral angles represents a helical twist or “kink” in the midsection of the peptide. Since the original submission of the manuscript describing these constrained linear and cyclic peptides, bradykinin B2 receptors have been cloned from other species including human [27,28]. Toward the goal of designing a nonpeptide antagonist as a human therapeutic agent, it would be interesting to evaluate these analogues against these newly reported receptor homologues. This would likely be fruitful and valuable given that there is evidence, including that presented in this chapter, showing that guinea pig and rat B2 receptors differ structurally from the human B2 receptor at the antagonist binding site. Hence, a structure-activity relationship established against the guinea pig or rat receptors could be misleading in the context of potential human therapeutics. A systematic study of the relative importances of amides and side chains in a prototypical second generation antagonist, NPC 18545 (DArg0-Arg1-Pro2-Hyp3-Gly4-Phe5-Ser6-DTic7-Oic8-Arg9) has recently been described [37,38]. The D-Arg0 and Ser6-DTic7-Oic8-Arg9 segments were left intact in all peptides on the assumption that N-terminal positive charge(s) and a hydrophobic C-terminal β turn are minimally required for binding. In a systematic fashion, the amino acids in the core of the peptide (Arg1Pro2-Hyp3-Gly4-Phe5) were substituted with glycine, an amino acid bearing no chirality or side chain. Binding assays, either in membranes from the guinea pig ileum or in membranes from a stable cell line expressing the human B2 receptor, were performed on each peptide and the results compared with the parent, NPC 18545, which has a Ki against [3H]-bradykinin of 0.08 nM. The elimination of all chirality and sidechain moieties in the segment Arg1-Pro2-Hyp3-Gly4-Phe5 via replacement by Gly1-Gly2-Gly3Gly4-Gly5 (NPC 18152), led to a peptide that no longer binds the receptor. This demonstrated that one or more of the side chains in this segment are critical during ligand-receptor interaction. Incorporation of either the Arg1 or Phe5 side chains led to improved potency (285 nM and 483 nM, respectively). Constructing a peptide with side chains of both Arg1 and Phe5 in place (NPC 18149) yielded a peptide with good affinity, Ki of 13.7 nM. Overall, this study shows that to maintain potency in the low picomolar range, peptides in this series require Arg1, Phe5, and either Pro2 or Hyp3 (but not both). The Nterminal charged moieties and the hydrophobic C-terminal β turn are also required. Potencies in the low nanomolar range are attainable without including the side chains or chirality associated with Pro2 and Pro3. These data have

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subsequently been successfully applied toward the design and synthesis of several nopeptide scaffolds and mimetics. Ultimately these mimetics were assembled in a combinatorial fashion as discussed in Section IV. In a related study, wherein NPC 18149 (DArg0-Arg1-Gly2-Gly3-Gly4-Phe5-Ser6-DTic7-Oic8-Arg9; Ki = 13.7 nM; Guinea pig ileum) was taken as the lead peptide, the relative contributions to binding affinity from each amide bond in the segment Arg1-Gly2-Gly3-Gly4-Phe5 were examined. Aminovaleric acid was used in a systematic fashion as a surrogate for any pair of adjacent Gly-Gly residues in the peptide. Aminovaleric acid is atomically identical to Gly-Gly with the exception that the amide bond linking the two glycines is replaced by two methylenes. The synthesis of Gly4-Phe5 required a special Gly-Phe mimic that has since been reported [39]. Since this substitution introduces flexibility into the peptide, it is a means of probing the structural role a given amide bond plays during receptor interaction. Potential hydrogen-bond donor and acceptor groups in the amide bond are removed via this substitution, which yields additional insights into potential electrostatic interactions that may also be important during ligand-receptor interactions. The conclusions drawn from the data are that in terms of structural or electrostatic interactions with this antagonist site on the receptor, the amide bond linking residues two and three may not be as critical as those linking residues three to four and four to five. Each of these investigations was aimed toward an understanding of either the backbone conformation of this prototypical decapeptide or the relative importance of the functional groups in the side chains that make significant contributions to receptor affinity. From the former, nonpeptide frameworks and scaffolds can be imagined. From the latter, insights into which functionality is required for high-affinity binding is derived. The remaining challenge is to reassemble these fragments onto synthetically feasible nonpeptide frameworks as potential new lead compounds. Our approach toward addressing this challenging problem is described later in this chapter. III. Receptor Structure-Based Investigations A. Elucidation of an Agonist Binding Site on the B2 Receptor In addition to the deductions one might make about a receptor binding site on the basis of receptor binding data from conformationally constrained ligands as previously described, models of bradykinin and bradykinin antagonists bound to their respective sites on the receptor as complimentary aspects of the overall strategy are also valuable. Unfortunately, due to the nature of the bradykinin

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receptor, it has not yet been obtained in crystalline form, nor is it likely to be in the near future. The bradykinin receptor is a member of a family of receptors for which an intracellular interaction with a G-protein is a critical part of the signal transduction pathway following agonist binding. Structurally, these G-protein-coupled receptors extend from beyond the extracellular boundary of the cell membrane into the cytoplasm. The tertiary structure is such that the protein crosses the bilayer of the cell membrane seven times, thus forming three intracellular loops, three extracellular loops, and giving rise to cytoplasmic C-terminal and extra-cellular N-terminal strands. It is generally presumed that the transmembrane domains of these receptors exist as a bundle of helical strands. This assumption is derived primarily from the known structure of the trans-membrane portions of a structurally related protein, bacteriorhodopsin [40]. G-Protein-coupled receptors do not lend themselves to analysis by either NMR or x-ray crystallography due to their structural dependence on an intact cell membrane. In our laboratories we pursued this valuable structural information by utilizing a combination of structural homology modeling, molecular dynamics, systematic conformational searching methods, and mutagenesis experiments. The combination of these techniques led to a proposed model of bradykinin bound to the agonist site on its receptor [41]. A hydrophobicity (Kyte-Doolittle) calculation [42] on the amino acid sequence of the rat bradykinin receptor yielded seven segments, each of which were 21 to 25 contiguous residues with predominantly hydrophobic side chains. These were presumed to be the seven transmembrane portions of the receptor. Cartesian coordinates of the backbone atoms within each of these seven segments were built by structural homology from the cryomicroscopic structure of the analogous segments of bacteriorhodopsin. Subsequently, side chains were added to these seven segments as appropriate for the rat bradykinin receptor, and the resulting geometry was optimized via constrained energy minimization to alleviate bad contacts. Extracellular and intracellular loops were extracted from the Protein Data Bank library, following a geometric search based upon a vector defined by terminal alpha carbons in adjacent helices. The model was subsequently subjected to a series of constrained and unconstrained energy minimizations as well as molecular dynamics simulations. The resulting structure of the receptor was used in a novel two-step docking procedure. Following our hypothesis that bradykinin adopts a C-terminal β turn upon complexation with the receptor, the φ, ψ backbone dihedral angles in the tetrapeptide corresponding to the C-terminus of bradykinin (Ser-Pro-Phe-Agr) were constrained in a harmonic fashion (force constant = 15 Kcal Å-1 mol1) to values that define a type II' β-turn [43]. This tetrapeptide probe was then systematically translated about the interior of a theoretical box inscribing the rat

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receptor model. The translations were such that the tetrapeptide probe molecule was incrementally repositioned within the receptor by following a 3 Å × 3 Å × 3 Å grid pattern. At each new position, both the probe and receptor were reset to their initial conformations, then the geometry of the complex was optimized using 200 steps of steepest descent followed by 500 steps of Adopted-Basis Newton-Raphson energy minimization. Subsequently, the sum of the steric and electrostatic contributions to the overall potential energy (interaction energy)—as measured only between the tetrapeptide probe molecule and the atoms of the receptor—were calculated. Slices through the receptor illustrating the energy of interaction as grayscale contour lines (darker gray = lower interaction energy) for that portion of receptor that was sampled by the tetrapeptide probe molecule is shown as an edge-on, frontal view in Figure 4. In this Figure it is qualitatively clear where the transmembrane domains are located (white), as well as where the most favorable sites of probe interaction are located (black).

Figure 4 Complete group of contour plots showing energy of interaction between probe and receptor. Each contour plot corresponds to a different horizontal slice as part of the first stage in the conformational search. Darker gray indicates most favorable interaction and the light shades represent least favorable interactions.

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This initial stage of the docking process was used to reduce the computational difficulties that would be inherent in “tumbling” a complete bradykinin molecule (which has great flexibility) about the receptor in a similar fashion. However, following this initial stage, insight into those regions of the receptor capable of accommodating the C-terminal portion of the bradykinin molecule was obtained. On the basis of energetics, and as qualitatively shown in Figure 4, those particular regions are clustered in the central part of the receptor near to the extracellular domain. Using this information as a steering device to limit the size of the problem, an exhaustive conformational search was performed using the entire nineresidue sequence of bradykinin as a probe molecule, again enforcing a C-terminal β turn via dihedral angle constraints. Specifically, 24 unique geometric orientations (eight on each of three axes) of the bradykinin molecule were sampled at each of 100 grid points identified during the initial stage as likely zones to bind the tetrapeptide probe molecule. Bradykinin-receptor complexes within the lowest 150 kcal mol-1 interaction energy with respect to the lowest found (17 complexes out of 2400) were grouped into sets of related conformational families, of which there were five. Computationally, each of the five complexes were presumed to be equally likely. All of these simulations were accomplished using custom routines written using the program CHARMm [21]. To guide the selection of which of the five bradykinin-receptor complexes to consider a “lead” model, supporting experimental evidence was sought from site-directed mutagenesis experiments. This support was taken primarily from work describing bradykinin binding assays performed of mutant rat B2 receptors [44,45]. The underlying strategy of the mutation studies was based on the hypothesis that, since bradykinin has positive charges at either end of its sequence (Arg1 and Arg9), separated by a group of rather hydrophobic amino acids (Pro2-Pro3-Gly4-Phe5-Ser6-Pro7-Phe8), it was likely that some acidic residues in the receptor participated during ligand binding. Several mutant receptors were made such that each contained either a point mutation or a small cluster of point mutations, wherein native residues, having negatively charged side chains (Asp, Glu), were replaced by alanine(s). Table 2 lists the initial cluster mutations (rat) that were prepared as well as the follow-up single point mutations (rat). Figure 5 shows a stereoview of the selected ligand-receptor complex chosen on the basis of best agreement with the results of these mutagenesis studies. None of the other four putative complexes were in agreement with this experimental data and were not considered further. Of particular significance in this work was that the trans-membrane residue Glu49, when mutated to alanine, showed no adverse effect on bradykinin receptor affinity with respect to rat wild type. A similar result was reported for the Glu196 rarrow.gif Ala196 mutation. These residues are remotely situated with respect to the proposed site of bradykinin

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binding and are colored light gray in Figure 5. In contrast, the [Asp175, Glu178,179] rarrow.gif Ala175,178,179 cluster mutation showed a 12-fold loss in bradykinin binding affinity, and the [Glu282, Asp286] rarrow.gif Ala282,286 cluster mutation lost 17-fold with respect to the wild type receptor. The Asp268 rarrow.gif Ala268 and Asp286 rarrow.gif Ala286 point mutations caused 19-and 28-fold respective losses in affinity for bradykinin. Close inspection of the bradykinin Arg1 side chain location and surrounding receptor interactions led to the suspicion that Asp286 and Asp268

Figure 5 Proposed model of bradykinin bound to the rat B2 receptor at the agonist binding site. Only the upper portion of the receptor is shown as gray helical ribbons. Bradykinin backbone and side chain atoms are shown as thick white licorice. Positions of point mutations having no significant adverse effects on bradykinin binding are shown as light gray spheres. Positions of mutations affecting bradykinin binding are shown as dark gray spheres.

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might be jointly interacting either with the guanidino group in the side chain of Arg1 or the N-terminal amino group in bradykinin. Therefore a receptor containing a double mutation (Asp268,286 rarrow.gif Ala268,286) would be expected to show a much more dramatic loss in affinity for bradykinin than would receptors containing the individual point mutations. The appropriate double mutation experiment confirmed this by causing a 500-fold loss in affinity for bradykinin, as predicted (Table 2). The mutagenized residues of this double mutant B2 receptor are colored dark gray in Figure 5. This type of an ionic interaction is also precedented by the body of literature that exists supporting the requirement of an N-terminal arginine residue and a free N-terminal amino group in both bradykinin peptide agonists and antagonists for high affinity binding. All of these mutant receptors were demonstrated to be functional receptors on the basis of bradykinin-induced membrane depolarization in a Xenopus oocyte expression system [44,45]. The selected agonist site model is characterized by an overall twisted S-shape ligand, similar to the conformation of bradykinin determined previously in a hydrophobic environment by NMR [19,46]. Overall, the model suggested that the N-terminal amino and guanidine groups of Arg1 interact directly with negatively charged amino acids in extracellular loop three, and the C-terminal end is in a β-turn conformation buried just below the extracellular boundary of the trans-membrane domain of the receptor. Noteworthy is the presence of a hydrophobic cavity in our receptor model located adjacent to Pro7 of the bradykinin ligand. This cavity is made up, in part, by the residues Phe261, Leu104, Val108, and Ile112. Given the historical significance of position seven in peptide bradykinin-like ligands, these residues represent interesting targets for further mutagenesis experiments. One such result, the mutation of Phe261 to Ala261, has already been described, and the results were supportive of this proposed model [47]. Antibodies to the extracellular loops two and three have also been shown to compete with bradykinin binding, lending further experimental support for an extracellular domain on this agonist binding site. More recently, chemical crosslinking combined with site-directed mutagenesis was used to analyze the bradykinin binding site in the human B2 bradykinin receptor [48]. Previous studies using the bovine B2 receptor showed that heterobifiunctional reagents reactive to amines and free sulfhydryls crosslink the bound bradykinin N-terminus to a sulfhydryl(s) in the receptor [49]. To identify this sulfhydryl(s), two conserved candidate residues in the human B2 receptor—Cys20 in the N-terminal domain and Cys277 in extracellular loop 3—were mutated to serine residues. Single and double mutants were expressed in Cos 7 cells. All mutants bound [3H]bradykinin with typical B2 receptor specificity. The heterobifunctional reagent m-maleimidobenzoyl-N-hydroxysuccinimide ester crosslinked bradykinin to wild-type and mutants with maximum efficiencies of 35% (wild type), 40% (Ser20), 20% (Ser277), and 0%

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(Ser20, Ser277). This clearly demonstrated that Cys20 and Cys277 are the only sulfhydryls available for crosslinking receptor-bound bradykinin. These results provided direct biochemical evidence that the Nterminus of bradykinin, when bound to the B2 receptor, is adjacent to extracellular loop 3 and the Nterminal domain in the receptor. Further consideration of the model led to the hypothesis that agonist peptides may minimally require an intact C-terminal β-turn structure with appropriate side chains in place and N-terminal amino and guanidine groups for primary electrostatic interaction(s) with Asp286 and Asp268 in extracellular loop 3. As a test of this hypothesis, the prototypical second generation antagonist, NPC 18545, (DArg0-Arg1Pro2-Hyp3-Gly4-Phe5-Ser6-DTic7-Oic8-Arg9) was modified such that residues 2–5 were replaced by a simple twelve carbon chain spacer (12-aminotridecanoic acid). The resulting compound, NPC 18325, contains only the appropriately charged moieties at the N-terminus, separated by a simple organic spacer moiety from a known β turn forming tetrapeptide [25,26]. This pseudopeptide was tested in the human bradykinin B2 receptor binding assay and found to have a Ki of 44 nM against [3H]bradykinin binding [50]. Functionally, this pseudopeptide was an agonist as measured by its ability to stimulate IP production in a stable CHO cell line expressing the human B2 receptor and in WI-38 cells. Since it was designed on the basis of an agonist site on the receptor, this result was not completely surprising despite the incorporation of the DTic-Oic pair at the C-terminus that previously had been shown to be critical in high affinity antagonists. Subsequently the length of the linear carbon chain was varied to further explore the hypothesis that the agonist site on the receptor had two domains, a hydrophilic site in the extracellular loop area and a hydrophobic domain in the transmembrane area [50]. Presumably, the two terminal portions of NPC 18325 can only simultaneously interact with each putative domain of the receptor binding site when the carbon chain is 12–13 methylenes long. But if the carbon chain length is shortened too far, this ligand might be unable to simultaneously interact with both domains, resulting in an affinity loss. This series of pseudopeptides and their respective human bradykinin B2 receptor affinities are presented in Table 3. The data are consistent with the hypotyhesis since the receptor affinity decreases as the carbon chain length is shortened. An alternative explanation of the data is that a certain hydrophobicity profile is required of the compounds in this series for good receptor affinity. These results indicated that there may be additional hydrophobic or flexibility prerequisites to binding in this series of pseudopeptides. One noteworthy observation was that NPC 18325 showed divergent behavior when evaluated against different species homologues of the bradykinin B2 receptor. Notably, in the guinea pig ileal membrane preparation assay, the affinity for the receptor was approximately 10-fold less than what had been

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observed for the human B2 [37]. Furthermore, in contrast to the functional activity of NPC 18325 at the human B2 receptor, the compound is a functional antagonist as measured against bradykinin-induced contraction of the isolated guinea pig ileum (pA2 = 5.5). These findings are in agreement with the concept that as a ligand is made smaller (i.e., fewer contact points possible with the receptor), the subtle structural differences in the binding sites on species variants of the same receptor become amplified. This observation further supports a cautionary posture toward developing nonpeptide antagonists for use in human diseases on the basis of results obtained in some animals including the guinea pig. Taking this new molecule as a lead structure, together with the receptor model and structure-activity relationship associated with related peptides including cyclic antagonists, the pursuit of several related pseudopeptides was undertaken. B. Elucidation of an Antagonist Site on the B2 Receptor There have been a variety of single alanine point mutations experimentally introduced into both rat and human bradykinin B2 receptors. Several of these have been shown to decrease the affinity of bradykinin to the receptor and have been implicated structurally near the agonist binding site. In contrast, at the time of this manuscript, there have been no mutations reported that adversely affect the ability of any peptide antagonists to bind to the receptor. Furthermore, antibodies raised against the certain extracellular domains of the kinin receptor compete with bradykinin for binding to the receptor but have no inhibitory

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action on the binding of antagonist peptides. In addition, it has been shown that bradykinin can be covalently crosslinked to the B2 receptor while antagonists cannot. These observations have fostered the belief that the agonist and antagonist binding sites of the receptor are not the same. At best, they may be partially overlapping, although there is no direct evidence for this. The ultimate identification of the amino acid residues that make up the antagonist site would be another important step toward the goal of structure-based design of novel nonpeptide antagonists. As described in a previous section of this chapter, characterizations of the bradykinin B2 receptors from rat and human using NPC 17410 (Figure 3) revealed different pharmacologies. Specifically, it showed a higher affinity for the human B2 receptor than it did for the rat B2 (human IC50 = 0.95 nM, rat IC50 = 48.0). This ligand “tool” provided a means for evaluating a series of bradykinin rat/human B2 receptor chimeras [51–53]. Several different chimeras were prepared in a systematic fashion and the affinity of NPC 17410 was determined for each. The chimeras are depicted schematically in Figure 6 together with the IC50 values determined for NPC 17410. Chimeras I through III sample the N-and C-terminal sections of the receptor for any contributions to an antagonist binding site. The remaining chimeras sample the core transmembrane domains of the receptor. Each chimera was shown to induce a membrane depolarization similar to wild type receptor in response to bradykinin when expressed in Xenopus oocytes. For each NPC 17410 assay, [3H]-NPC 17731 was used as the radioligand. From this systematic approach, specific groups of contiguous residues within the receptor were identified as possible contributors to an antagonist binding site. The NPC 17410 binding to chimeras III, IV, and VIII showed rat-like pharmacology (low NPC 17410 affinity). The NPC 17410 binding to chimeras I, II, VI, and VII showed human-like NPC 17410 pharmacology (high receptor affinity). Binding to chimeras V and VIII, however, was similar to rat-like NPC 17410 pharmacology, but the affinity of the compound was slightly shifted back toward human-like results. Comparisons of rat and human receptor sequences in the regions sampled by the chimeras reveals that only two clusters of residues differ between rat and human B2 receptors. Specifically, TM2 has the same sequence in rat and human receptors so it is unlikely that the differential pharmacology associated with NPC 17410 binding can be attributed to residues there. However, TM3 has a cluster of 3 residues that differ (rat rarrow.gif human: Thr110 rarrow.gif Ala108, Met111 rarrow.gif Ile109, Tyr113 rarrow.gif Ser111) and TM6 has a cluster of 5 residues that differ (rat rarrow.gif human: Phe259 rarrow.gif Leu257, Leu256 rarrow.gif Ile254, Val255 rarrow.gif Ile253, Gly252 rarrow.gif Leu250, Ala249 rarrow.gif Val247) in rat and human receptors. These differences represent important targets for follow-up point (and cluster) mutation experiments. Our current thinking is that the largest effects on NPC 17410 pharmacology, if any, might be derived from the TM3 mutants since

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Figure 6 Rat and human B2 receptor chimera constructs and affinity data for binding NPC 17410. Also shown is the affinity NPC 17410 to rat and human wild type receptors.

between rat and human, these are quite diverse. However, it is also possible that the cluster of residues identified in TM6, while not radically dissimilar, may as a group create different hydrophobic environments between these species homologues. The most significant individual difference within the TM6 zone is the Phe259 (rat) rarrow.gif Leu257 (human) swap and might therefore be most significant

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Figure 7 Schematic of the primary amino sequence of the human B2 receptor. Shown in black are residues experimentally identified as contributing to an agonist binding site. The dark gray residues are suspect positions for contributing to an antagonist site. The residues colored light gray have been mutagenized only in the rat B2 receptor, but they are conserved in the human. The Thr263 rarrow.gif Ala mutation interferes with agonist binding only, while Gln260 partially interferes with agonist and first generation antagonist binding.

within the context of these TM6 residues. Currently, we have prepared these mutant receptors, but at this time binding to NPC 17410 remains unfinished. A summary of the amino acids in the human B2 receptor implicated in comprising either agonist or antagonist sites are highlighted in Figure 7. Marked in dark black are the residues of extracellular loop 3, TM 6, and the extracellular N-terminal segment that have been shown to participate in agonist binding. Marked in dark gray in TM 6 and TM 3 are residues likely to partially comprise an antagonist binding site based primarily on the chimeric receptor studies described previously, although there is no explicit experimental evidence as yet. Shown in light gray are two residues that are conserved between rat and human B2 receptors. Mutagenesis experiments have been done on this pair in the rat B2

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receptor with interesting results [54]. Mutations in Thr263 only affect agonist binding, not antagonist. Mutations in Gln260 affect binding of bradykinin and first generation antagonist peptides. As depicted in the figure, it is possible that the agonist and antagonist binding sites have domains on opposite sides of the helix that makes up TM 6, with Gln260 being situated partly in both. IV. Design and Combinatorial Synthesis of Nonpeptidic Antagonists As was previously described, a significant body of information was generated that provides insights into the key structural features of bradykinin receptor binding sites and the residues that participate in ligand binding. In addition, from the ligand-based studies, knowledge about relevant structure-activity relationships was acquired. Our modular synthetic strategy was based primarily upon the recognition that high-affinity ligands appear to be comprised of three domains. These domains are (1) a positively charged N-terminal segment, (2) a midsection containing a bend or twist with some hydrophobic substituent attached and, (3) a C-terminal segment of appropriate hydrophobicity and structurally simulating a type II' β turn. Models of potent cyclic and linear peptide bradykinin receptor antagonists (described previously) were used in a comparative fashion to select nonpeptide ring systems from a database of chemical structures fine chemicals database. For each, some degree of chemical diversity was achieved by altering one of several parameters including, o, m, or p substitution of an aromatic ring or nature of alkyl substituent(s) as well as point(s) of synthetic attachment [55,56]. Each nonpeptide fragment was designed within the framework of several criterion. First, a given scaffold must closely match the known SAR and be compatible with the putative ligand binding site structure. Second, each scaffold must be a relatively simple synthetic target, having readily available starting material, no chiral centers and having a total synthesis of not more than 4–5 steps. Finally, each template must have a “C-terminal” carboxylate and an “N-terminal” amino group with no interfering functionality such that it could be readily used in a solid phase synthetic strategy. Given that each nonpeptide we identified was a viable surrogate for either the second or third domain of high-affinity ligands (as described above) our goal was to rapidly explore the receptor affinities of all possible combinations of these nonpeptide templates at position X and Y of the sequence DArg-Arg-X-Y-Arg, hence a combinatorial synthetic approach was taken. In this study, there were four linear aminoalkanoic acids [50], four different cinnamic acids, three different carbolines, three different phenanthridinones, and five different spirocyclics. The variability in the phenanthridinione series

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was that the central ring could be opened or cleaved and the amino group could be meta or para substituted on the latter. In the carboline series, the cyclic amino group was either at the β or γ position of the cyclohexenyl ring and the methylene chain bearing the “C-terminal” carboxylate could be of variable length. The spirocyclic series was varied by alkyl, cycloalkyl, and aryl substitution on the fivemembered ring amine nitrogen. The cinnamic acids had two carbon chains that could be of varying length, one of which had the further possibility of containing a double bond(s). Rather than perform individual syntheses of all possible combinations of these nonpeptide units, members of each ring type or scaffold family were pooled in equimolar amounts prior to incorporation into the sequence DArg-Arg-X-Y-Arg. Since each individual member of a given pool was constructed on a similar carbocyclic scaffold, the chemical environment of the N-terminal amino group and Cterminal carboxylate groups were expected to follow similar kinetic and thermodynamic controls during the attachment of the nonpeptide residue to the growing peptide chain. The use of these smaller, directed libraries made it readily practical to obtain HPLC and mass spectral data for each and therefore confirm the composition of the library. Ultimately, 10 libraries of novel nonpeptidic structures were synthesized following typical solid-phase methodologies. Each library contained from nine to thirty-six different compounds in approximately equimolar amounts. Unpurified libraries were tested in a receptor binding assay utilizing membrane preparations from a stable CHO cell line expressing the human B2 receptor. Each library was tested at concentrations between 10 nM and 1µM. The ability of each library to inhibit[3H]-bradykinin binding was assessed and the results are presented in Figure 8a. Although this type of screening is highly qualitative, certain libraries appear in Figure 8a that show higher affinity to the receptor than other libraries. Library one (of the series DArg-Arg-PH-CN-Arg) was ultimately selected for further deconvolution. This library was further broken down (decoded) in order to determine which compound(s) were responsible for the apparent activity. It is important to note that breaking these libraries down to elucidate the structure of the hit(s) was feasible due to the inherently small size of each library. Library one contained 12 different structures (recall that there were originally three different phenanthridinones and four different cinnamic acids). The first deconvolution step of the approach is shown in Figure 9. Here, only the CN position was randomized, and the PH moieties were specific. This led to the preparation of three new libraries of 4 compounds each. Receptor binding was again performed as before and only one of these three new sublibraries showed affinity for the receptor at 10 µM (Figure 8b). The final step in the process required to elucidate the active component(s) was to synthesize and purify each of the 4 members of this library as shown in Figure 9. Receptor binding on these

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Figure 8 (a) Binding assay results for 10 original nonpeptidic libraries of the sequence DArg-Arg-X-Y-Arg, where X and Y are defined as PH = phenanthridinone, CB = carboline, SP = spirocycle, SC = straight chain, CN = cinnamic acid. Each library was tested at 1 µnM and 10 nM. Results were compared to cold bradykinin binding, which was tested at two lower concentrations, 0.1 nM and 1 nM. Panels (b) and (c) correspond to the receptor binding results obtained using the two breakdown steps from original library number 1, as shown in Figure 9.

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Figure 9 Composition of ten original nonpeptidic libraries of the sequence DArg-Arg-X-Y-Arg. X and Y were selected from the set of scaffolds shown in Table 1. Also shown are the subsequent breakdown libraries from original library number 1. Two-letter codes used in the figure correspond to the different nonpeptide moieties described in Table 1. Specifically, PH = phenanthridinone, CB = carboline, SP = spirocycle, SC = Straight chain, CN = cinnamic acid.

four novel nonpeptidic structures showed that only one of the four had affinity to the receptor (Figure 8c). This new compound, I, was subsequently shown to be an antagonist in a cellular assay measuring bradykinin-stimulated IP turnover [18]. Overall, there were 285 possible structures to survey due to the number of structure-based scaffolds that were prepared. This was rapidly accomplished via 19 synthetic couplings, 19 assays, and 4 purifications. Not surprisingly, compound I showed divergent potency when assayed in different species homologues of the bradykinin B2 receptor. In particular, in a model of bradykinin-induced hypotension in rats and rabbits, it showed no activity. Likewise, it did not block bradykinin-induced contraction of the isolated guinea pig ileum. Since compound I is considerably smaller than previously reported decapeptide antagonists, subtle structural differences (which are http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_144.html (1 of 2) [4/5/2004 5:00:24 PM]

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known to exist in species homologs of the bradykinin B2 receptor) are likely amplified. A more comprehensive pharmacological analysis of compound I is currently underway. A. Lead Optimization We have previously reported that the C-terminal guanidinyl moiety of Arg [9] in prototypical peptide bradykinin antagonists is likely to behave more as an aromatic functional group rather than a hydrogenbond donor/acceptor. This speculation was based on proposed models of the agonist and antagonist binding sites of this receptor that have been elucidated using molecular biological and computational procedures. On this premise, the newly discovered lead compound, I, was altered such that the Cterminal arginine was replaced by 3',5'-dimethylpyrimidylornithine in an attempt to increase potency. This known mimetic of arginine contains an aromatic 3',5'-dimethylpyrimidyl ring in the side chain rather than the guanidino group on naturally occurring arginine. The results of the receptor binding assay performed using this compound, IA, are shown in Table 4 where it is clear that affinity to the human B2 receptor is improved with respect to compound I. This data is supportive of the notion that the C-terminal residue(s) in this new series of bradykinin antagonist compounds interact with a hydrophobic environment, perhaps within the transmembrane domain of the receptor as previously suggested. The discovery of I and IA is significant in many regards. First, they are highly nonpeptidic lead compounds that could be further modified to improve potency and/or reduce molecular weight. Such improvements might lead to novel therapeutic agents for the treatment of inflammatory diseases. Thus far in the kinin antagonist literature there is significant evidence showing that, for compounds containing a C-terminal arginine residue, removal of that arginine generally yields compounds that are antagonists of the B1 subtype of the bradykinin receptor. Following a similar strategy with compound I could lead to the discovery of a novel series of nonpeptidic B1 receptor antagonists, although this remains to be demonstrated. V. Conclusions There has been a significant effort invested toward the discovery of novel bradykinin receptor antagonists during the past decade. In that time, several generations of peptide antagonists have been developed and a few are in human clinical trials. The pursuit of nonpeptide antagonists of the human bradykinin B2 receptor continues and incorporates a wide range of strategic approaches. The approach described herein is an early and very good example of a combinatorial synthesis of nonpeptide building blocks that mimic peptide structure,

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ultimately tested in a nontagged, solution-phase form. Perhaps more significant is that the success described here demonstrates a possible synergy between structure-based design and combinatorial methodology. This approach has many merits, but the most significant is the application of structurally directed libraries toward target binding-site structures which, for one reason or another, may not be fully characterized. This method serves to aim the combinatorial syntheses in a logical direction, rather than attempt to prepare libraries of vast diversify (and numbers). Finally, since the libraries of compounds that were prepared contained few members, it was possible to analytically characterize each of the pools to assess integrity of their composition. Overall, there are many important advantages in the paradigm we have adopted that make the strategy generally viable in the context of structure-based lead compound discovery. References 1. Greenbaum LM. Adv Exp Med Biol 1986; 198A:55. 2. Okamoto H, Greenbaum LM. Biochem Biophys Res Commun 1983; 112:701. 3. Muller-Esterl W. Thromb Haemost 1989; 61:2. 4. Bhoola KD, Figueroa CD, Worthy K. Pharmacol Rev 1992; 44:1. 5. Proud D, Perkins M, Pierce JV, Yates KN, Highet PF, Mangkornkanok/Mark M, Bahu R, Carone F, Pisano JJ. J Biol Chem 1981; 256:10634. 6. Kitamura N, Takagaki Y, Furoto S. Nature 1983; 305:545. 7. Ward PE. In: Burch RM, ed. Bradykinin Antagonists: Basic and Clinical Research. New York: Marcel Dekker, 1991:147–170. 8. Perkins MN, Campbell EA, Davis A, Dray A. Br J Pharmacol 1992; 107:237P. 9. Perkins MN, Campbell EA, Dray A. Pain 1993; 53:191. 10. Perkins MN, Kelly D. Br J Pharmacol 1993; 110:1441. 11. Kyle DJ, Burch RM. Curr Opin Invest Drugs 1993; 2:5. 12. Kyle DJ, Burch RM. Drugs of the Future 1992; 17(4):305. 13. Huang HM, Lin TA, Sun GY, Gibson GE. J Neurochem 1995; 64:761. 14. Rubin LE, Levi R. Circ Res 1995; 76:430. 15. Bao G, Gohlke P, Unger T. J Cardiovasc Pharmacol 1992; 20(Supplement 9):S96. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_147.html (1 of 2) [4/5/2004 5:00:59 PM]

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16. Martorana PA, Kettenbach B, Breipol G, Linz W, Scholkens BA. Eur J Pharmacol 1990; 182:395. 17. McDonald KM, Mock J, D' Ajoia M, Parrish T, Hauer K, Francis G, Stillman A, Cohn JN. Circulation 1995; 91(7):2043. 18. Schwieler JH, Kahan T, Nussberger J, Hjemdahl P. Am J Physiol 1993; 264:E631. 19. Kyle DJ, Blake PR, Hicks RP. In: Burch RM, ed. Bradykinin Antagonists: Basic and Clinical Research. New York: Marcel Dekker, 1991:131–146. 20. Chou PY, Fasman GD. Biophys J 1979; 26:367. 21. (a) Brooks BR, Bruccoleri RE, Olafson BD, States DJ, Swaminathon S, Karplus MJ. CHARMM: a program for macromolecular energy minimization, and dynamics calculations. J Comp Chem 1983; 4:187–217.

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(b) Molecular Simulations, Inc., 16 New England Executive Park, Burlington, MA 01803-5297. 22. Kyle DJ, Martin JA, Burch RM, Carter JP, Lu S, Meeker S, Prosser JC, Sullivan JP, Togo J, Noronha-Blob L, Sinsko JA, Walters RF, Whaley LW, Hiner RN. J Med Chem 1991; 34:2649. 23. Kyle DJ, Burch RM. Drugs of the Future 1992; 17(4):305. 24. Hiner RA, Chakravarty S, Lu S, et al. J Med Chem 1996; manuscript in preparation. 25. Kyle DJ, Green LM, Blake PR, Smithwick D, Summers MF. Pep Res 1992; 5:206. 26. Kyle DJ, Blake PR, Smithwick D, Green LM, Martin JA, Sinsko JA, Summers MF. J Med Chem 1993; 36:1450–1460. 27. McEachern AE, Shelton ER, Shakta S, Obernolte R, Bach C, Zuppan P, Fujisaki J, Aldrich RW, Jarnagin K. Proc Natl Acad Sci USA 1991; 88:7724. 28. Hess JF, Borkowski JA, Young GS, Strader CD, Ransom RW. Biochem Biophys Res Commun 1992; 184(1):260. 29. Menke JG, Borowski JA, Bierilo KK, et al. J Biol Chem 1994; 269:21583–21586. 30. Hock FJ, Wirth K, Albus U, Linz W, Gerhards HJ, Wiemer G, Henke S, Breipohl G, Knoig W, Knolle J, Schölkens BA. Br J Pharmacol 1991; 102:769. 31. Wirth K, Hock FJ, Albus U, Linz W, Alpermann HG, Anagnostopoulos H, Henke S, Breipohl G, Knoig W, Knolle J, Schölkens BA. Br J Pharmacol 1991; 102:774. 32. Sawutz DG, Salvino JM, Seoane PR, Douty BD, Houck WT, Bobko MA, Doleman MS, Dolle RE, Wolfe HR. Biochem 1994; 33:2373. 33. HOE SDS MICELLES 34. Chakravarty S, Wilkens D, Kyle DJ. J Med Chem 1993; 36:2569. 35. Momany FA. In: Metzger, RM, ed. Topics in Current Physics. Vol. 26; New York: Springer Verlag, 1981:41–79. 36. Momany FA, Chuman H. Meth Enz 1986; 124:3–17. 37. Kyle DJ. Brazil J Med Bio Res 1994; 27:1757. 38. Chakravarty S, Mavunkel BJ, Lu S, Goehring R, Wu JP, Connolly M, Valentine H, Liu YW, Tam C, Andy R, Kyle, DJ. 23rd European Peptide Symposium, Braga: Sept 4–10, 1994.

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39. Mavunkel BJ, Lu Z, Kyle DJ. Tett Lett 1993; 34:2255. 40. Henderson R, Baldwin JM, Ceska TA, Zemlin F, Beckmann E, Downing KH. J Mol Biol 1990; 213:899. 41. Kyle DJ, Chakravarty S, Sinsko JA, Stormann TM. J Med Chem 1994; 37:1347. 42. Kyte J, Doolittle RF. J Mol Biol 1982; 157:105. 43. Rose GD, Gierash LM, Smith JA. Adv Prot Chem 1985; 37:1. 44. Novotny E, Bednar D, Connolly M, Connor J, Stormann T. In: Burch RM, ed. Molecular Biology and Pharmacology of Bradykinin Receptors. Austin, TX: R. G. Landes Company, 1993:19–30. 45. Novotny EA, Bednar DL, Connolly MA, Connor JR, Stormann TM. BBRC 1994; 201:523. 46. Lee SC, Russell AF, Laidig WD. Int. J Pept Prot Res 1990; 35(5):367. 47. Freedman R, Jarnagin K. Cloning of a B2 Bradykinin Receptor: Recent Progress on Kinins. Basel: Birkhauser Verlag, 1992:487–496. 48. Herzig MCS, Leeb-Lundberg F, Nash N, Connolly M, Kyle DJ. Kinin '95. International conference on kallikreins and kinins, Denver, CO, Sept, 1995. 49. Herzig MCS, Leeb-Lundberg F. J Biol Chem 1995; in press. 50. Chakravarty S, Connolly MA, Kyle DJ. Peptide Research 1995; 8:16.

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51. Nash N, Connolly MA, Stormann TM, Kyle DJ. 14th Am Pep Sym, June 18, 1995. 52. Nash N, Connolly MA, Stormann TM, Kyle DJ. Mol Pharm 1996; manuscript in preparation. 53. Burch RM, Kyle DJ, Stormann TM. In: Molecular Biology and Pharmacology of Bradykinin Receptors. Austin, Texas: R. G. Landes Company, 1993:19–32. 54. Nardone J, Hogan PG. PNAS 1994; 91:4417. 55. Chakravarty S, Mavunkel B, Andy R, Kyle DJ. Network Science 1995; 1:1. 56. Chakravarty S, Mavunkel BJ, Andy R, Kyle DJ. 14th American Peptide Symposium, Columbus, Ohio, June, 1995.

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5 Design of Purine Nucleoside Phosphorylase Inhibitors Y. Sudhakara Babu, John A. Montgomery, and Charles E. Bugg BioCryst Pharmaceuticals, Inc., Birmingham, Alabama W. Michael Carson, Sthanam V. L. Narayana, and William J. Cook The University of Alabama at Birmingham, Birmingham, Alabama Steven E. Ealick Cornell University, Ithaca, New York Wayne C. Guida and Mark D. Erion* Ciba-Geigy Corporation, Summit, New Jersey John A. Secrist, III Southern Research Institute, Birmingham, Alabama I. Introduction A. Enzymology Purine nucleoside phosphorylase (PNP, E.C. 2.4.2.1) catalyzes the reversible phosphorylysis of ribonucleosides and 2'-deoxyribonucleosides of guanine, hypoxanthine, and related nucleoside analogs [1]. It normally acts in the phosphorolytic direction in intact cells, although the isolated enzyme catalyzes the nucleoside synthesis under equilibrium conditions. Figure 1 shows the chemical reaction. The enzyme has been isolated from both eukaryotic and prokaryotic organisms [2] and functions in the purine salvage pathway [1,3]. Purine nucleoside phosphorylase isolated from human erythrocytes is specific for the 6-oxypurines and many of their analogs [4] while PNPs from other organisms vary in their specificity [5]. The human enzyme is a trimer with identical subunits and a total molecular mass of about 97,000 daltons [6,7]. Each subunit contains 289 amino acid residues. *Current

affiliation: Gensia, Inc., San Diego, California.

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Figure 1 The reaction catalyzed by PNP.

B. Pharmacology Interest in PNP as a drug target arises from its ability to rapidly metabolize purine nucleosides and from its role in the T-cell branch of the immune system. Unfortunately, PNP can also cleave certain anticancer and antiviral agents that are synthetic mimics of natural purine nucleosides, thus interfering with therapy. One such substance is ddI (2'3'-dideoxyinosine), which the Food and Drug Administration approved as a treatment for AIDS in 1991. Another is the potential anticancer agent 2'-deoxy-6thioguanosine [8]. Our goal was to develop a compound that when administered with the nucleoside analogs would inhibit PNP while the anticancer and antiviral agents accomplished their therapeutic missions. The combination of purine nucleoside analogs and a PNP inhibitor might prove to be a more effective treatment. The PNP inhibitors alone have potential therapeutic value based on the importance of PNP to the immune system. Patients lacking PNP activity exhibit severe T-cell immunodeficiency while maintaining normal or exaggerated B-cell function [9]. We, like other researchers, quickly recognized that PNP inhibitors might selectively suppress the T-cell proliferation associated with an array of autoimmune disorders such as rheumatoid arthritis, psoriasis, systemic lupus erythematosus, multiple sclerosis, and insulin-dependent (juvenile-onset) diabetes [10]. This profile also suggests that PNP inhibitors might be useful in the treatment of T-cell proliferative diseases—such as T-cell leukemia or Tcell lymphoma—and in the prevention of organ transplant rejection. C. Drug Design Strategy Recent advances in biotechnology, macromolecular crystallography, computer graphics, and related fields have led to a new approach in drug discovery called structure-based drug design. Structure-based drug design requires a detailed structural knowledge of the target (enzyme or receptor) and the interaction of small molecules with it.

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Figure 2 Structure-based drug design strategy.

A tight fit is necessary for potency and specificity. A drug that binds to its target and inactivates it for a long time can be administered in lower doses than one that rapidly separates from its target. A substance designed to mesh perfectly with a particular binding site of one target is unlikely to interact well with any other molecule, minimizing unwanted interactions and side effects. Having chosen PNP as the target, we followed a systematic strategy for designing inhibitory compounds. Figure 2 outlines the overall strategy of this approach. To serve as a drug, an inhibitor has to readily cross cell membranes to the interior of cells, where PNP is located.

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We determined the structure of human PNP by x-ray crystallography and used these results in combination with computer-assisted molecular modeling to design inhibitor candidates. We examined how well the shape and chemical

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structure of a candidate would complement the active site of PNP. We used computational chemistry to estimate the strength of the attractive and repulsive forces between a candidate and the enzyme. We synthesized only those candidates suggested by chemical intuition and computer simulation to have high affinity for the target. Then we measured the inhibition of PNP and compared the proposed with the actual fit. Because modeling programs and expert opinion are imperfect, certain compounds did not meet expectations. After exploring the reasons for the successes and the failures, we returned to interactive computer graphics to propose modifications that might increase the effectiveness of drug candidates. The resulting compounds were evaluated by determination of their IC50 values (the inhibitor concentration causing 50% inhibition of PNP) and by x-ray diffraction analysis using difference Fourier maps. This iterative strategy—modeling, synthesis, and structural analysis—led us to a number of highly potent compounds that tested well in whole cells and in animals. D. Previously Known Inhibitors At the beginning of our studies several PNP inhibitors had been reported with Ki values in the 10-6 to 107 range, including 8-aminoguanine [11], 9-benzyl-8-aminoguanine [12], and 5'-iodo-9-deazainosine [13]. Acyclovir diphosphate had been shown to have a Ki near 10-8 if assayed at 1 mM phosphate rather than the more frequently used value of 50 mM phosphate [14]. During our studies, the synthesis of 8-amino9(2-thienylmethyl)guanine was reported with a Ki of 6.7 × 10-8 M [15]. Figure 3 illustrates some of these structures. Despite the potential benefits of PNP inhibitors and the large number of PNP inhibitors that had been synthesized, no compound had reached clinical trials. None of these compounds were potent enough to be useful for therapy and also capable of crossing the cell membrane intact. Although potencies for the best compounds had affinities 10–100 fold higher than the natural substrate (Km = 20 µM), it is expected that T-cell immunotoxicity will only occur with very tight binding inhibitors (Ki < 10 nM) due to the high level of in vivo PNP activity and competition with substrate. II. Crystallography At the present time, x-ray crystallography is the preferred technique for obtaining the required atomic resolution structural data. In the late 1970s when this project was first conceived, determining the structure of a protein was far from routine. The x-ray structural determination occupied a team of crystallographers led by Steven E. Ealick, then at the University of Alabama at Birmingham through most of the 1980s.

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Figure 3 Previously known inhibitors.

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The stumbling block did not lie with obtaining pure PNP or converting the protein into crystals. Robert E. Parks, Jr. and Johanna D. Stoeckler of Brown University had already isolated the enzyme from human cells. They supplied quantities of protein to William J. Cook, who succeeded in preparing the well-ordered crystals required for x-ray studies [16]. We established that PNP crystals function normally as a catalyst. Thus crystalline PNP is essentially identical to PNP in the body. If it were profoundly different, one would have no justification for basing drug design on the crystal structure. In the early years we had to depend on a relatively low-intensity x-ray source. High-resolution data was obtained through collaboration with John R. Helliwell and his group at the Daresbury Laboratory Synchrotron Radiation Source in England. Today greatly improved equipment and more synchrotron facilities are available for protein crystallography. The three-dimensional structure was determined by multiple isomorphous replacement techniques using synchrotron radiation [17]. The native and guanine-PNP complex structures have been refined to 2.8 Å resolution [18,19]. A. Structure of the Enzyme Crystals of human PNP are grown from ammonium sulfate solution and stored in artificial mother liquor solution made of 60% ammonium sulfate in 0.05 M citrate buffer at pH 5.4. The space group is R32 with hexagonal cell parameters a=142.9(1) Å and c=165.2(1) Å. The PNP crystals contain about 76% solvent and diffract to around 2.8 Å resolution. The x-ray data established that PNP crystals contain a high percentage of water. This feature proved very useful; proposed drugs could easily be soaked into the active site without disrupting the crystal packing. Figure 4A shows the

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Figure 4 Structure of PNP as stereo drawings. (A) Crystal packing. The low-resolution surfaces of six trimers are shown. The top level is related to the bottom level by a 2-fold axis along X. The distance between the 3-fold axes is 143 Å. A drug molecule is shown in the solvent channel near the entrance to an active site. (B) Ribbon drawing. The lowermost trimer of Figure 4A is shown. This is the native structure; the guanine and phosphate are shown to mark the active site. (C) The swinging gate. The trimer of Figure 4B is rotated about 30° counterclockwise in the plane, followed by a roughly 90° rotation about X to view the entrance to the active site. A model of the transition state is shown as a line drawing. Conformational changes of the protein on binding guanine are shown. Arrows are drawn from the Cα positions in the native structure to their positions in the complex. (D) Active site of PNP. The orientation is approximately that of Figure 4C, but enlarged and clipped to focus on the substrate. Key side-chain residues are labeled. Residue 159'F, in the center of figure toward the viewer, is the only residue from the adjacent subunit. The guanosine and phosphate are shown with thicker bonds. Oxygen and sulfur atoms are shown as white spheres, nitrogen and phosphorus as black spheres. (E) Purine binding pocket. The style is the same as Figure 4D, but the figure is rotated slightly and enlarged. The key group interacting with bound guanine are highlighted. (F) The best inhibitor. The style is the same as Figure 4D, but the figure has been enlarged and rotated to place the phosphate binding site far from the viewer in the upper left.

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Figure 4 (Continued) Only one nitrogen atom from Arg 84 is visible, which—along with Ser 220 and His 86—interact with the acetyl group branching from the benzylic carbon. The chlorinated phenyl group is in the center of the figure, interacting with the aromatic groups in the sugar binding site. The guanine group interactions are the same as seen in Figure 4E. Figure prepared with ribbons (http://www.cmc.uab.edu/ribbons).

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large solvent channels and the position of the active site. The x-ray analysis confirmed the trimeric nature of the enzyme, as the subunits are related by the crystallographic three-fold axis. A ribbon diagram of the trimer is shown in Figure 4B. Each monomer contains an eight-stranded β sheet and a five-stranded β sheet that join to form a distorted β barrel. Seven α helices surround this β sheet structure. The active site is an irregular indentation on the surface of the enzyme, located from the position of a tightly bound sulfate ion and various substrate analogs. These investigations revealed the identity of the exact amino acids constituting the active site region; such detail was a prerequisite to drug design. Information of greater import emerged from analyses of the complexes formed when synthetic nucleosides, including previously discovered inhibitors, were diffused into the active site. B. The Active Site The structural determinations also yielded a surprise. The shape of the enzyme changes when a purine is bound. The famous lock-and-key analogy [20] has a fallacy; the shape of the lock is not static, but flexible. Awareness of these conformational changes critically aided our modeling efforts, allowing prediction of which parts of PNP could change shape to interact with a proposed inhibitor. A “swinging gate” consisting of residues 241–260 controls access to the active site (Figure 4C). These residues in the native structure had poorly defined electron density with high thermal motion. The gate opens in the native enzyme to accommodate the substrate or inhibitor. The maximum movement caused by substrate or inhibitor binding occurs at His 257, which is displaced outwards by several angstroms. After binding, the electron density becomes well defined. The gate is anchored near the central β sheet at one end and near the C-terminal helix at the other end. The gate movement is complex and appears to involve a helical transformation near residues 257–261. Consequently, initial inhibitor modeling attempts using the native PNP structure were far less successful than subsequent analyses in which coordinates for the guanine-PNP complex were used. Because of the magnitude of the changes that occur during substrate binding, it is unlikely that modeling studies based on the native structure alone would have accurately predicted the structure of PNP/inhibitor complexes. The active site is located near the subunit-subunit boundary within the trimer and involves seven polypeptide segments from one subunit and a short loop from the adjacent subunit (Figure 4D). The purine binding site employs residues Glu 201, Lys 244, and Asn 243 to form hydrogen bonds with N1, O6, and N7 of purine. The remainder of the purine binding pocket is largely

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hydrophobic, composed of residues Ala 116, Phe 200, and Val 217. The phosphate binding site uses residues Ser 33, Arg 84, His 86, and Ser 220 with the phosphate positioned for nucleophilic attack at C1' of the nucleoside. The sugar binding site is mostly hydrophobic consisting of residues Tyr 88, Phe 200, His 257 from one subunit and Phe 159 of the adjacent subunit. This hydrophobic pocket orients the sugar to facilitate nucleophilic attack by phosphate and subsequent inversion of C1'. C. Initial Inhibitor Complexes In order to understand the interaction of inhibitors with the active site residues, the previously known inhibitors were obtained and crystallographic analyses were carried out. The most important findings were (1) 8-amino substituents enhance binding of guanines by forming hydrogen bonds with Thr 242 and possibly the carbonyl oxygen atom of Ala 116; (2) substitution by hydrophobic groups at the 9position of a purine enhances binding through interaction with the hydrophobic region of the ribose binding site; and (3) acyclovir diphosphate is a multisubstrate inhibitor with the acyclic spacer between the purine N9 and the phosphate of near optimal length to accommodate these two binding sites. Based on these results, a number of starting compounds were proposed that incorporated these and other features predicted to enhance inhibitor binding. III. Molecular Modeling Structural information in combination with graphical methods for displaying accessible volume, electrostatic potential, and hydrophobicity of the active site of the target macromolecule greatly facilitates the drug design process. Accurate prediction of binding affinities and protein conformational changes are currently not routinely possible, although significant advances are being made. Proposed compounds were screened by modeling the enzyme-inhibitor complex using interactive computer graphics. Macromodel [21] and AMBER [22] based molecular energetics were used along with Monte Carlo/energy minimization techniques [23] to sample the conformational space available to potential inhibitors docked into the PNP active site. Methods based on the work of Goodford [24] employing custom software were also used. Qualitative evaluation of the enzyme-inhibitor complexes by molecular graphics and semiquantitative evaluation of the interaction energies between the inhibitors and the enzyme aided in the prioritization of compounds for chemical synthesis.

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IV. Drug Design Progression We focused initially on filling the purine binding region of the active site. That done, we planned to fill the sugar binding region and, finally, the phosphate binding site. We expected that each successive step, moving the compound closer toward fully occupying the active site, would enhance the affinity of the drug candidate for the enzyme. A. The Purine Site From our crystallographic examinations, we knew that three amino acids in the purine binding pocket of PNP formed hydrogen bonds with purines and their mimics. Such linkages are among the strongest reversible chemical bonds that exist. In proposing inhibitor candidates, we concentrated on compounds that would at least form hydrogen bonds with the same three amino acids. Figure 4e shows a close-up of the purine site. We favored exchanging a carbon atom for the nitrogen atom that normally occupies position nine, since there was no interaction of this nitrogen with the active site and earlier studies showed such a change promotes binding to PNP. Guanine modified in this way is called 9-deazaguanine. The first structures selected for synthesis were 9-deazaguanines substituted by an arylmethyl group at the 9 position. These compounds were prepared by adaption of a literature procedure [25]. We further expected that attaching an amino group to the carbon atom in position eight on 9-deazaguanine would enhance affinity, since 8aminoguanine was the first significant inhibitor of PNP. Both 8-aminoguanine analogs and 9-deazaguanine analogs are good inhibitors of PNP. However, introduction of an 8-amino group into the 9-deazaguanine derivatives resulted in decreased potency. To understand this poor binding, we undertook crystallographic analysis of PNP complexes with four compounds having the 9-thienyl substituent attached to guanine (G), 8-aminoguanine (8AG), 9deazaguanine (DG), and 8-amino-9-deazaguanine (8ADG). The results of this analysis are summarized in Figure 5. These data show one mode of binding for compounds that accept a hydrogen bond from Asn 243 at N7 (G and 8AG) and another for compounds that donate hydrogen to Asn 243 from N7 (DG and 8ADG). The 8AG analogs make use of the Thr 242 side chain to form an additional hydrogen bond, which improves binding affinity. In the 9-deazaguanine series, where N7 has an attached hydrogen atom, Asn 243 undergoes a shift that is clearly seen in difference Fourier maps. This shift is caused by the formation of the N7-H…OD(243) hydrogen bond. A concomitant shift by Thr 242 prevents it from hydrogen bonding to the 8-amino group of 8ADG. Furthermore, the shift moves the methyl group of Thr 242 towards the 8-amino

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Figure 5 Comparison of 8-amino and 9-deaza substitutions on guanine. Details are extensively discussed in the text. Cross-hatching schematically indicates the enzyme. Ball and stick diagrams show the inhibitor and key side-chain residues. Nitrogens are dark gray spheres, oxygens are light gray. Arrows indicate hydrogen bonding, with the arrow size showing relative strength.

amino group, generating a hydrophobic environment for the group and decreasing binding affinity. The carbon-for-nitrogen switch in the 9-deaza variant favors association with PNP by substituting a strong hydrogen bond for the relatively weak one occurring between Asn 243 and guanine. Formation of a simple 8-aminoguanine variant leads to tight binding by giving rise to an extra hydrogen bond between the purine derivative and Thr 242. The combination of the two “improvements”—the carbon-for-nitrogen substitution and the addition of the amino group to position eight—was counterproductive because the carbon in position nine prevented the amino group at

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position eight from forming the extra bond with Thr 242. In fact, it set up an unfavorable, repulsive clash between the threonine and the added amino group. In the absence of detailed structural information, it would have been extremely difficult to explain why affixing the amino group to the carbon in position eight proved unhelpful. But crystallography quickly provided the explanation. 9-Deazaguanine itself would be a better choice for the purine component of an inhibitor. This experience underscores the wonderful economy of the structure-based approach. Without crystallographic data, we might have pursued a logical but unproductive avenue of research much longer than we did. B. Ribose Site The next task was to fill the sugar binding site. The sugar in a nucleoside does not attach to PNP primarily by forming hydrogen bonds, but through hydrophobic attractions. The sugar binding pocket of PNP consists of three hydrophobic amino acids: Phe 200 and Tyr 88 from the same monomer that binds guanine and Phe 159 from the adjacent monomer. Several known inhibitors carried a benzene group attached to position 9 of the purine in place of the sugar in the nucleoside. An initial series of compounds was synthesized to exploit the hydrophobic region in the ribose binding site. A number of 9-substituted 9-deazapurine analogs were prepared with various aromatic, heteroaromatic, and cycloaliphatic substituents. The first 9-deazaguanine derivatives synthesized, such as 9-benzyl-9deazaguanine, were three to six times more potent than the most potent known inhibitor, 8-amino-9-(2thienylmethyl)guanine. The optimum spacer between the purine base and the aromatic substituent proved to be a single methylene group. Crystallographic data showed that generally the planes of the aromatic rings tend to orient in a reproducible conformation. The aromatic groups optimize their interaction with Phe 159 and Phe 200, which results in the classic “herringbone” arrangement reported in a variety of aromatic systems [26]. Inhibitors with cycloaliphatic substituents at N9 of deazaguanine were also as potent as the aromatic analogs. The cycloaliphatic substituents occupied the same general volume as the aromatic groups. As with the aromatic series, the optimum spacer between the 9-deazaguanine and the hydrophobic substituent is one carbon atom. X-ray analysis of the PNP complexes of 9-cyclohexyl-9-deazaguanine, a relatively poor inhibitor, and the complex of 9-cyclohexylmethyl-9-deazaguanine, a potent inhibitor, showed the two cyclohexyl groups occupy approximately the same space in the active site with the purine base pulled out of its optimal position in the former. The chemistry is more straightforward with the aromatic series. From modeling studies, we saw that the sugar binding pocket could be filled more

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completely by adding any of several chemical groupings to the benzene ring. The best fit came from adding a chlorine atom to position 3 of the benzene ring. C. Phosphate Site The final step added a group that would interact with the phosphate binding site either directly or via electrostatic interactions. We could not use phosphate itself, because phosphate-containing compounds are not metabolically stable and have difficulty passing through cell membranes intact. Acyclovir diphosphate, which is not membrane permeable and is subject to extracellular metabolism, is a good example. Our results suggested that an ideal PNP inhibitor in the 9-deazapurine series would contain an aromatic group and a substituent with affinity for the phosphate site interlinked by spacers with optimum lengths. Crystallographic and modeling studies suggested a two-to-four-atom spacer. Initial modeling studies encouraged us to prepare several structures, but they failed to improve the binding affinity of our twopart structure. Crystallographic analysis of a number of PNP inhibitor complexes revealed significant displacement of the inhibitors. These displacements appear to be the result of close contacts between the inhibitor and the ion in the phosphate binding site. Sulfate ions occupy the phosphate site in PNP crystals as they are grown from ammonium sulfate solution. These inhibitors were more potent when the binding was measured in 1 mM phosphate solution rather than in 50 mM phosphate. Kinetic studies showed that these inhibitors were competitive not only with inosine but also with phosphate, in keeping with the above observation. These results, summarized in Table 1, show that the IC50 (50 mM) is equal to or larger than the IC50 (1 mM), in some cases by as much as 100-fold. The ratio and the dimension of the 9-substituent show some correlation. Compounds such as 8-aminoguanosine and 8-amino-9-(2-thienylmethyl)guanine show no difference. Since the concentration of phosphate in intact cells is 1 mM, we routinely used this assay condition for all PNP inhibitors. Starting with a model of the 9-benzyl-9-deazaguanine/PNP complex, we concluded that two of the positions on the 9-benzyl group, namely the 2-position of the phenyl ring and one of the benzylic sites, appeared to be oriented so that a group attached to either one could interact favorably with the phosphate binding site. The first compound made in this series, 9-[2-(3-phosphonopropoxy)benzyl]guanine, turned out to be a poor PNP inhibitor. Subsequent crystallographic analysis revealed that the plane of the aromatic ring had rotated approximately 90° from its optimum position in the hydrophobic pocket. This reorientation of the ring was presumably necessary to accommodate the four atom spacer between the phenyl ring and the phosphonate group. A compound

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IC50, µM R2a

R2

(S)-3-Chlorophenyl 3-Chlorophenyl

CH2CO2H CH2CN

50 mM phosphateb

Ratiod 1 mM phosphatec

0.031

0.0059

5.3

1.8

0.010

180

2-Tetrehydrothienyl

H

0.22

0.011

20

3,4-Dichlorophenyl

H

0.25

0.012

21

3-Thienyl

H

0.08

0.020

4

3-Trifluoromethylcyclohexyl

H

0.74

0.020

37

Cyclopentyl

H

1.8

0.029

62

Cycloheptyl

H

0.86

0.030

29

Pyridin-3-yl

H

0.20

0.030

2-(Phosphonoethyl)phenyle

H

0.45

0.035

13

Cyclohexyl

H

2.0

0.043

47

2-Furanyl

H

0.31

0.085

3.6

CH2CO2H

0.90

0.16

5.6

H

42

1.0

(R)-3-Chlorophenyl 2-Phosphonopropoxyphenyle aCompounds

with R2 not equal to H are racemic mixtures unless the R or S isomer is designated.

bCalf

spleen PNP assayed in 50 mM phosphate buffer.

cCalf

spleen PNP assayed in 1 mM phosphate buffer.

dIC

at 50 mM phosphate divided by IC50 at 1 mM phosphate.

50

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42

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eGuanine

base.

Source: Ref. 27.

with a two-carbon spacer was a much better PNP inhibitor; however, it was clear from x-ray analysis that the aromatic ring was unable to form the ideal “herringbone” packing interaction. Alternatively, compounds were modeled in which the spacer to the phosphate binding site branched from the benzylic carbon, thus placing no restrictions on the tilt of the aromatic ring. Examination of the 9-benzyl-9-deazaguanine/PNP complex indicated that of the two benzylic positions, one (pro-R) pointed into a sterically crowded area within the active site, whereas the other (pro-S) pointed into a relatively empty space adjacent to the phosphate binding site. This analysis led to the synthesis of racemic 9-[1-(3chlorophenyl)-2-carboxyethyl]-9-deazaguanine. This compound adds an acetate group (CH2COO-) to the methylene carbon atom that joined 9-deazaguanine to the chlorinated benzene ring. This compound was resolved into its (S) and (R) enantiomers.

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As predicated, the (S) acid was a 30-fold more potent inhibitor of PNP than the (R) form. X-ray crystallographic analysis of the complexes revealed that the (S) acid was oriented properly for optimal interactions with all three subsites (Figure 4F), whereas the (R) acid was not. This series of compounds contains the most potent membrane-permeable inhibitors of PNP yet reported [27]. V. Summary Recently, scientists at BioCryst have successfully completed a project to design and synthesize potent inhibitors of the enzyme Purine Nucleoside Phosphorylase (PNP) using the three-dimensional structure of the active site. Crystallographic and modeling methods have been combined with organic synthesis to produce inhibitors. Our experience in creating a set of potential drugs—one of which (BCX-34) is now in human trials for treating psoriasis and a form of T-cell lymphoma—illustrates the process and the power of structure-based design. This structure-based inhibitor design approach led to a number of inhibitors more than 100 times more potent than any membrane-permeable inhibitor available at the beginning of this project. During the two and half years of this project, about 60 active compounds were synthesized. This is a remarkably small number compared with the extensive synthesis programs generally involved in drug discovery by trial and error techniques. The large number of active compounds and the enhancement of inhibitor potency stand as proof that crystallographic and modeling techniques are now capable of playing a critical role in the rapid discovery of novel therapeutic agents. The entire protocol, from choosing the target to creating a drug suitable for clinical trials, can probably be accomplished today in two or three years. A. Obstacles Encountered and Lessons Learned Crystallographic analysis was based primarily on the results of difference Fourier maps in which the interactions between residues in the active site and the inhibitor could be characterized. During these studies, about 35 inhibitor complexes were evaluated by x-ray crystallographic techniques. It is noteworthy that the resolution of the PNP model extends to only 2.8 Å and that all of the difference Fourier maps were calculated at 3.2 Å resolution, much lower than often considered essential for drug design. Crystallographic analysis was facilitated by the large solvent content that allowed for free diffusion of inhibitors into enzymatically active crystals. Initial inhibitor modeling attempts using the native PNP structure were far less successful than subsequent analyses in which coordinates for the guanine-

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PNP complex were used, mainly because of the magnitude of the changes that occur during substrate binding. We found that computer modeling required significant tuning in order to provide useful results. Crystallographic results were useful in testing and modifying modeling parameters. The most useful modeling results were achieved after incorporation of the conformational searching techniques described earlier and when the coordinates for the PNP-guanine complex model were used. Visual inspection and chemical intuition were very important. B. Perspectives of Treating Targeted Disease One of the inhibitors designed during the drug discovery process, 9-(3-pyridylmethy)-9-deazaguanine (BCX-34), was selected for initial clinical development. Current clinical trials utilize both topical and oral formulations of the drug. Researchers at the University of Alabama at Birmingham and Washington University School of Medicine have recently completed small Phase II clinical trials of two indicated applications, cutaneous T-cell lymphoma (CTCL) and psoriasis, using a topical formulation of BCX-34. Although patients showed improvement in both trials, the duration of each was too short (six weeks) to adequately assess the efficacy of the drug. Subsequently 80% of the patients from the CTCL trial (24 patients) entered an open label trial for treatment of their disease for up to twelve months. At the end of the first six months of treatment, seven of the patients were in complete remission (verified by biopsy), two patients showed a clinical complete response, and nine patients had shown definite improvement. The other six patients had shown no change or progression of disease. No serious, drug-related adverse events were reported during the study. The process of structure-based drug design helped to ensure that the inhibitor would be highly selective for the PNP enzyme, and thus far no other targets for the drug have been identified. The mechanism of action of BCX-34 appears to be entirely related to its effect on the proliferation of human T-cells. This high degree of specificity probably also contributes to the high safety profile of the drug. Although longterm studies in more patients will be necessary to substantiate these results, it appears likely that BCX34 will have a significant clinical effect on at least some T-cell mediated diseases. Based on the results from these three trials, BioCryst has initiated a multicenter Phase III trial for the treatment of CTCL, as well as a large, multicenter Phase II trial for psoriasis. In addition to the two clinical trials using the topical formulation, a Phase I clinical trial in CTCL and T-cell lymphoma/leukemia has begun using an oral formulation of BCX-34. In the future, a number of other Tcell mediated diseases or processes are possible targets for BCX-34, including rheumatoid arthritis, multiple sclerosis, inflammatory bowel disease, and organ transplant rejection.

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References 1. Parks RE Jr., Agarwal RP. In: Boyer PD, ed. The Enzymes. 3rd Ed., New York: Academic, 1972; 7:483–514. 2. Stoeckler JD. In: Glazer, RE, ed. Developments in Cancer Chemotherapy. Florida: CRC, Baco Raton, 1984; 35–60. 3. Friedkin M, Kalckar H. In: Boyer PD, Lardy H, Myrback K, eds. The Enzymes. 2nd Ed. New York: Academic, 1961; 5:237–55. 4. Agarwal KC, Agarwal RP, Stoeckler JD, Parks RE Jr. Purine nucleoside phosphorylase. Microheterogeneity and comparison of kinetic behavior of the enzyme from several tissues and species. Biochemistry 1975; 14:79–84. 5. Bzowska A, Kulikowska E, Shugar D. Properties of purine nucleoside phosphorylase (PNP) of mammalian and bacterial origin. Z Naturforschung C Biosci 1990; 45:59–70. 6. Stoeckler JD, Agarwal RP, Agarwal KC, Schmid K, Parks RE Jr. Purine nucleoside phosphorylase from human erythrocytes: physiocochemical properties of the crystalline enzyme. Biochemistry 1978; 17:278–83. 7. Williams SR, Goddard JM, Martin DW Jr. Human purine nucleoside phosphorylase cDNA sequence and genomic clone characterization. Nucleic Acids Res 1984; 12:5779–87. 8. LePage GA, Junga IG, Bowman B. Biochemical and carcinostatic effects of α'-deoxythiguanosine Cancer Res 1964; 24:835–40. 9. Giblett ER, Ammann AJ, Wara DW, Sandman R, Diamond LK. Nucleoside-phosphorylase deficiency in a child with severely defective T-cell immunity and normal B-cell immunity. Lancet 1975; 1:1010–3. 10. Otterness I, Bilven M. In: Rainsford K, Velo G, eds. New Developments in Antirheumatic Therapy. Inflammation and Drug Therapy Series. Norwell, MA: Kluwer Academic, 1989; 2:277–304. 11. Stoeckler JD, Cambor C, Kuhns V, Chu SH, Parks RE Jr. Inhibitors of purine nucleoside phosphorylase, C(8) and C(5') substitutions. Biochemical Pharmacology 1982; 31:163–71. 12. Shewach DS, Chern JW, Pillote KE, Townsend LB, Daddona PE. Potentiation of 2'-deoxyguanosine cytotoxicity by a novel inhibitor of purine nucleoside phosphorylase, 8-amino-9-benzylguanine. Cancer Res 1986; 46:519–23. 13. Stoeckler JD, Ryden JB, Parks RE Jr, Chu MY, Lim MI, Ren WY, Klein RS. Inhibitors of purine nucleoside phosphorylase: effects of 9-deazapurine ribonucleosides and synthesis of 5'-deoxy-5'-iodo-9deazainosine. Cancer Res 1986; 46:1774–8. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_168.html (1 of 2) [4/5/2004 5:02:33 PM]

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14. Tuttle JV, Krenitsky TA. Effects of acyclovir and its metabolites on purine nucleoside phosphorylase. J Biol Chem 1984; 259:4065–9. 15. Gilbertsen RB, Scott ME, Dong MK, Kossarek LM, Bennett MK, Schrier DJ, Sircar JC. Preliminary report on 8-amino-9-(2-thienylmethyl) guanine (PD 119,229), a novel and potent purine nucleoside phosphorylase inhibitor. Agents and Actions 1987; 21:272–4. 16. Cook WJ, Ealick SE, Bugg CE, Stoeckler JD, Parks RE Jr. Crystallization and preliminary X-ray investigation of human erythrocytic purine nucleoside phosphorylase. J Biol Chem 1981; 256:4079–80.

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17. Ealick SE, Rule SA, Carter DC, Greenhough TJ, Babu YS, Cook WJ, Habash J, Helliwell JR, Stoeckler JD, Parks RE Jr, Chen SF, Bugg CE. Three-dimensional structure of human erythrocytic purine nucleoside phosphorylase at 3.2 Å resolution. J Biol Chem 1990; 265:1812–20. 18. Narayana SVL, Bugg CE, Ealick SE. Refined structure of purine nucleoside phosphorylase at 2.75 Å resolution. Acta Cryst D 1996; accepted. 19. Babu YS, Refined structure of guanine: purine nucleoside phosphorylase at 2.8 Å resolution. In preparation. 20. Koshland DE Jr. The key-lock theory and the induced fit theory. Angew Chem Int Ed Engl 1994; 33:2375–8. 21. Mohamadi F, Richards NGJ, Guida WC, Liskamp R, Lipton M, Caufield C, Change G, Hendrickson T, Still WC. MacroModel—An integrated software system for modeling organic and bioorganic molecules using molecular mechanics. J Comput Chem 1990; 11:440–67. 22. Weiner SJ, Kollman PA, Case DA, Singh UC, Ghio C, Alagona S, Profeta S, Weiner P. New force field for molecular mechanical calculations simulations of proteins and nucleic acids. J Am Chem Soc 1984; 106:765–84. 23. Chang G, Guida WC, Still WC. An internal coordinate monte carlo method for searching conformational space. J Am Chem Soc 1989; 111:4379–86. 24. Goodford P. A computational procedure for determining energetically favorable binding sites on biologically important macromolecules. J Med Chem 1985; 28:849–57. 25. Montgomery JA, Niwas S, Rose JD, Secrist JA 3d., Babu YS, Bugg CE, Erion MD, Guida WC, Ealick SE. Structure-based design of inhibitors of purine nucleoside phosphorylase. 1. 9-(arylmethyl) derivatives of 9-deazaguinine. J Med Chem 1993; 36:55–69. 26. Burley SK, Petsko GA. Aromatic-aromatic interaction: a mechanism of protein structure stabilization. Science 1985; 229:23–8. 27. Ealick SE, Babu YS, Bugg CE, Erion MD, Guida WC, Montgomery JA, Secrist JA 3d. Application of crystallographic and modeling methods in the design of purine nucleoside phosphorylase inhibitors. PNAS USA 1991; 88:11540–4.

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6 Structural Implications in the Design of Matrix-Metalloproteinase Inhibitors John C. Spurlino 3-Dimensional Pharmaceuticals, Inc., Exton, Pennsylvania I. Matrix-Metalloproteinases The matrix metalloproteinases (MMPs) are a family of ubiquitous enzymes that are involved in extracellular matrix degradation and remodeling. They are critical for the processes of morphogenesis and wound healing, but are also implicated in many human diseases including arthritis, metastasis, and cancer tumor growth [1, 2]. This family includes matrilysin, fibroblast collagenase (HFC), neutrophil collagenase (HNC), stromelysin 1 (HFS), stromelysin-2, stromelysin-3, gelatinases A and B, collagenase3, and the membrane type MMP. In addition to the destructive involvement in diseases, MMPs play a critical role in the remodeling of the extracellular matrix [3]. The MMP enzyme family is part of the superfamily of metzincins. The metzincin superfamily is distinguished by a conserved zinc binding motif for the catalytic zinc and a Met-turn region [4]. The MMPs are unique in that they also contain a second structural zinc, however this zinc may be absent in the intact full-length enzyme [5]. The presence of one to four structural calcium ions has been detected in the MMPs that have been characterized to date. The importance of the zinc ions and at least one of the structural calcium ions to enzymatic activity has been proven [6]. The MMPs are secreted as inactive proenzymes, which are activated by proteolytic cleavage. Once activated they are subject to control by tissue inhibitors of metalloproteinases (TIMPs). It is the imbalance between the active enzymes and the TIMPs that leads to destructive tissue degradation that potent directed pharmaceuticals can overcome. These enzymes have been the target of

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drug design since the late 1970s [7]. Batimastat, [{4-N-hydroxyamino}-2R-isobutyl-3S-{thienylthiomethyl} succinyl]-L-phenyl-alanine-N-methylamide, a potent nonspecific MMP inhibitor from British Bio-Tech, is now in Phase III trials. The information gained from current studies indicate that there is some efficacy in the treatment of disease states by MMP inhibitors [8–10]. There is still much debate about whether a broad spectrum or directed MMP inhibitor is the best course of treatment for a variety of diseases, partly because the exact role of individual MMPs is still unclear. Both collagenase-3 and HFC are suspected to cause osteoarthritis [11]. It is currently believed that gelatinase and collagenase-3 have a role in breast cancer [12]. Gelatinase A and B have been implicated in hemorrhagic brain injury [13]. Gelatinase A and B, HFC, and stromelysin may all be involved in gastric cancer [14]. Matrilysin may be implicated in colon cancer [15]. Increased gelatinase A and B activity has also been seen in response to beta-amyloid production [16]. The role that individual MMPs play in causing these diseases, however, remains unclear. This uncertainty underscores the need to develop selective inhibitors of individual MMPs to ferret out the roles each play in the development of specific disease states. The MMPs consist of one or more structural domains (Figure 1). The first domain, the propeptide domain, confers a self-inhibitory action on the full-length MMP. The second domain contains the active site residues and is referred to as the catalytic domain. The catalytic domain is characterized by the conserved zinc-binding sequence (HEXGHXXGXXHS), which also contains the glutamate residue that is essential for activity [17]. The MMPs are activated by cleavage of the prodomain. All MMPs contain these first two domains. Matrilysin, the simplest of the MMPs, is an example of a two-domain enzyme, where the active enzyme consists solely of the catalytic domain. The remainder of the MMPs also contain a hemopexin-like domain connected to the catalytic domain by a proline-rich linker. This domain is involved in the interaction of the collagenases and stromelysins with collagen and, in the case of the collagenases, is essential for activity against collagen [18]. The cleavage of the proline-rich linker region in HFC and HNC is another route to control collagenase activity. The hemopexin domain of the gelatinases is not necessary for collagen binding, but may be involved in receptor recognition [19]. The gelatinases also contain a fibronectin-like insert in the catalytic domain that is involved in binding collagen [20]. The fibronectin domain has also been shown to be essential for elastolytic activity [21]. A structural picture of these additional domains is essential for an understanding of the mode of action for these larger MMPs, but is not necessary for a structure-based drug design strategy. Differences in the catalytic domains of the MMPs can be used to drive a targeted drug discovery effort.

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Figure 1 A ribbon model of the full-length collagenase structure (1fbl.pdb). The prodomain would precede and include the portion labeled in the figure. The catalytic domain is shown, with a highlighted region where the fibronectin-like domain of the gelatinases is inserted.

II. 3-Dimensional Structure of MMPs Catalytic domain structures for fibroblast collagenase [22–25], neutrophil collagenase [26,27], matrilysin [28], and stomelysin [29, 30] have all been determined and deposited in the Protein Data Bank [31]. The catalytic domains of MMPs, as seen in the archetypal collagenase structure (shown as a ribbon drawing [32] in Figure 1), consist of an upper 5-stranded β sheet flanked by two α helices on one side of the active site cleft and a long loop that contains the Met-turn flanked by a single α helix on the other side of the cleft. The active-site groove as seen in the solvent-accessible surface [33] is an obvious structural feature (Figure 2). The top wall of the cleft (as seen in Figure 2) is formed by the top strand of the β sheet and the loop that contains the calcium binding site. The lower wall of the cleft is formed from the residues on either side of the Met-turn. These residues can be considered as an interrupted strand which, together with the substrate, complete the twisted β sheet of the catalytic domain. The bottom of the cleft is formed by the second helix, which

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Figure 2 The accessible surface of HFC with a modeled substrate from human collagen showing the binding sites.

contains the HExxH motif, the catalytic zinc, and the S1' pocket. Substrates bind in an extended conformation that approximates an antiparallel strand. The cleft, however, is not large enough to accommodate a triple helix collagen molecule. A structure for the full-length active porcine synovial collagenase [34] has been determined. The structure of the catalytic domain of this full-length enzyme is equivalent to the structures of the isolated catalytic domains of HFC, HNC, and matrilysin. The flexible linker domain between the catalytic and hemopexin domains is disordered and the orientation of the hemopexin domain in the structure offers no real clue as to the mode of action for the full-length collagenases. Furthermore, the matrilysin structure of the full-length active enzyme has almost identical secondary structural features (a Ca overlap of

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Figure 3 The sequence alignment of MMPs with the catalytic domain region highlighted. The residues that line the subtrate pockets are marked: S3 (3), S2 (2), S1 (1), S1 (*), S2' (@), and S3' (#). The highlighted catalytic domain alignment was dominated by the structural alignment of the determined structures of HFC, HNC, and matrilysin.

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0.43 Å) as the catalytic-domain structure of HFC. This demonstrates that the absence or presence of the hemopexin domain does not affect the overall structure of the catalytic domain. The sequence homology of the catalytic domains of the collagenases is 62%. This can be extended to the other members of the MMP family as seen in Figure 3. An understanding of the structural features of the target enzyme is essential for structure-based drug design. In this example we will be looking at inhibiting MMPs by binding to the active site. The numbering system used throughout this chapter will be in regard to the HFC sequence used in 1hfc.pdb. III. Surface Features The surface features of matrilysin, fibroblast collagenase, and neutrophil collagenase are all similar (Figure 4). The active-site groove can be plainly seen on the surface: two main pockets punctuated by the active-site zinc. Modeling

Figure 4 The accessible surfaces of HFC (a), HNC (b), and matrilysin (c) are shown with a bound P'-side hydroxamate inhibitor.

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studies show that there is not sufficient room in the S1'-S3' cleft to accommodate the native coiled triple collagen bundle. The active site consists of a series of subsites on either side of the catalytic zinc. These subsites are numbered starting at the catalytic zinc and preceding from N to C as S1', S2', etc., corresponding to the residues (P1', P2', etc.) of the substrate that is bound (Figure 2). Likewise the subsites are numbered S1, S2, etc. outward from the other side of the catalytic zinc. While the interaction of the substrate (and the inhibitor) with the catalytic zinc is the most important interaction, the remainder of the substrate (inhibitor) also forms hydrogen bonds with residues from the top strand of the β sheet and the loop region posterior to the Met-turn. These interactions with the substrate in the binding pockets of the MMPs are the prime targets for engineering specific MMP inhibitors. An in-depth understanding of the differences of the properties of these pockets in the different MMPs and the interactions of specific residues within these pockets is essential for structure-based design of inhibitors. IV. Main-Chain Substrate Interactions Most of the hydrogen bonds between the substrate and the MMP occur with the top strand of the β sheet. The P3 residue does not make any direct hydrogen bonds with the MMP. The P2 residue makes two hydrogen bonds with residue 184, which is a conserved alanine residue in all the aligned MMP sequences. Residue 183 is a conserved histidine, which is bound to the structural zinc, further stabilizing the conformation of the top strand. The carbonyl oxygen of P1 is liganded to the catalytic zinc. The lefthand side of the substrate is thus held in place by only two hydrogen bonds with the enzyme and one interaction with the catalytic zinc. Although the P3 residue does not make any hydrogenbond contributions to substrate binding, it is essential for catalytic activity [43]. The right-hand side of the substrate is held much tighter. The P1' residue's carbonyl oxygen makes a hydrogen bond with the amide nitrogen of residue 181, which is a conserved leucine residue. The amide nitrogen of the P1' residue is hydrogen bonded to the conserved alanine-182 carbonyl oxygen. The P2' substrate residue is held in place by hydrogen bonds to proline 238 and tyrosine 240, two more conserved residues. The amide nitrogen of P3' makes a hydrogen bond with the carbonyl oxygen of residue 179, the only nonconserved residue, making a main-chain interaction with the substrate. The use of conserved residues to maintain the main-chain interactions along the substrate backbone makes differentiation of the MMPs through these interactions difficult. Instead, differences in the regions where the side chains of the substrate interact can be used to drive the discovery of specific MMP inhibitors. The hydrogen-bonding pattern also indicates that right-hand side (P'-

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side) inhibitors will bind with greater affinity. Indeed, most of the structures of MMPs were determined with right-hand side inhibitors, and most of the pharmaceuticals currently in development are also righthand side binders. A closer look at the binding pockets themselves also demonstrates the reasons for the preference of researchers for the right-hand side. V. Substrate-Binding Pockets The nonprimed or left-hand side of the cleft consists of a large shallow depression. The S1 pocket consists of a shallow ridge that complements the glycine residue of the collagen strands. Most of the interactions with the glycine residue are brought about due to its interaction with the catalytic zinc. Asparagine 180 approaches the P1 residue in HFC. Crystallographic evidence indicates an interaction of the thiophene ring of batimastat via electrostatic interactions of the p orbitals and the catalytic zinc and the possibility of a water-mediated hydrogen bond to the carbonyl O of residue 184 [35]. Larger substituents can be accommodated in regions adjacent to the P1 pocket, possibly in the large pocket above the S1 site (see Figure 2). Increased potency was noted for several compounds with cyclic imido P1 substituents that could bind here [2]. The S2 pocket is a large shallow depression offering no real binding cavities. One side of the pocket is made up from the conserved histidine at position 228 and the main chain from residue 227. The bottom of the pocket is formed by histidine 222. Both of these histidines are liganded to the catalytic zinc. The other side consists mostly of the residue 186 side chain with some hydrogen-bonding contacts possible from the tip of the glutamine side chain in the case of HFC and HNC. The S3 pocket offers a shallow cavity to bind the conserved proline. The proline residue of the substrate would lie between the side chains of His 183, Phe 185, and Ser172 [36]. Residues 183 and 185 are conserved among the MMPs with the minor exception of a tyrosine replacing phenylalanine 164 in stromelysin. Residue 172 shows some variability among the MMPs existing as a serine in HNC and HFC and a tyrosine in the remainder of the aligned MMPs. The primed or right-hand side of the active site exists as a narrow canyon with a large well at the beginning. The S1' pocket is a narrow, deep cavity providing an ideal binding site for inhibitor design. The S1' pocket is the most significant feature of the surface, extending as a tunnel completely through the enzyme in the case of neutrophil collagenase and stromelysin. This feature makes the S1' pocket an ideal candidate for use in designing an inhibitor with specificity for HNC, HFC, or HFS. The volumes of the S1' pockets vary greatly. Matrilysin has the smallest S1' pocket at 111 Å3. The fibroblast collagenase pocket is not much larger at

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Figure 5 (a) A cut away view of the S1' pocket of HFC showing the termination of the pocket by Arg214. Matrilysin also has a truncated pocket. (b) A cut away view of the S1' pocket of HNC showing the clear path through to the other side of the enzyme. The gelatinases are also likely to have large extended S1' pockets.

123 Å3. The pocket of the neutrophil collagenase that travels through the enzyme has a volume of 305 Å3. Figure 5 demonstrates the differences in the relative sizes of the S1' pockets of HFC and HNC. Stromelysin has a pocket that should be of similar size as that of neutrophil collagenase [21]. The gelatinases and collagenase-3, based on sequence alignment, should also posses long tunnel-like S1' pockets. The residues that line the S1' pocket are mostly hydrophobic residues (see Figure 3) and show a remarkable overall similarity. The specificity of a number of inhibitors of MMPs can be linked to differences in the S1' pockets. The S1' pocket of HFC is terminated by arginine 214, while matrilysin has a tyrosine residue that accomplishes the same thing. The remainder of the aligned MMPs have leucine residues at that position. In addition the conformation of the leucine residue is swung back, forming the tunnel. There are three additional residues that are significant in their differences within the S1' pocket: residues 239 and a two-residue insert, relative to HFC, after residue 242. These residues

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form the lower end of the tunnel. The major effect of the different residues found here is on the diameter of the exit hole in the S1' tunnel. However, there are some residues that could be targeted for hydrogenbond formation. The S2' binding cavity is a narrow cleft that can easily accommodate a peptide backbone, but with no room for a side chain. The interaction of the P2' side chain is made with the exterior surface of the enzyme. The S2' site is exposed to solvent and presents two possible interaction sites for bound inhibitors that are related by a rotation about χ1. These sites consist of residues 179–180 on one side and residues 238–240 on the other. The S3' binding cavity opens up out of the canyon and consists almost entirely of surface interactions. The side chain of P3' interacts with residues 210 and 240, which are mostly conserved tyrosine residues. Additional interactions could be formed with some of the additional residues found in the insertions after residue 242. VI. Structure-Based Design Structure-based drug design is an iterative process that starts with a lead compound, a structural model of the target, and a structure-activity relationship

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(SAR) model. The lead compound can come from compound screening, previously discovered inhibitors, or it can be based on a known substrate. The model can be obtained from x-ray crystallography, high-field nuclear magnetic resonance spectroscopy, or from homology-based model building. Inhibitor structures are developed and docked into the model of the binding site of interest, typically the active site of the enzyme. The interactions of the inhibitor-enzyme complex are evaluated and ranked. The most promising compounds are then synthesized and tested. Based on the results of the testing, additional enzyme-inhibitor structures are determined, the SAR model is updated, and the process beings again. As a model case of structure-based drug design for MMPs we will look at the design of a right-handed inhibitor based on the x-ray structures of HFC and HNC. VII. Zinc-Binding Group The design of active-site inhibitors based on the natural substrate of the collagenases has produced a variety of zinc-binding groups to anchor the inhibitor to the catalytic zinc. These group include hydroxamates, thiols, phosphorous acid derivatives (phosphinate, phosphonate, phosphoramidate), and carboxylates. The selection of a suitable zinc-binding group has been studied in depth [37–40]. The most potent zinc-binding group found for the collagenases to date is the hydroxamate. The structural comparison of hydroxamate, carboxylate, and sulfodiimine in matrilysin provided information on the contribution of the zinc ligand to the overall potency of the inhibitor [19]. The potency of the zinc-bind group can be directly related to the number of bonds in which it is involved for this instance. The hydroxamate is the perfect bidentate ligand to the zinc with both oxygens being within 2.2 Å of the zinc. The hydroxamate group also is involved in hydrogen bonds with Glu219 and the carbonyl oxygen of Ala182. The carboxylate group is also a bidentate ligand to zinc, however the oxygens are not equidistant from the zinc. The carboxylate forms only one additional hydrogen bond with Glu219 of the enzyme. The sulfodiimine bound to matrilysin is a monodentate zinc ligand and the weakest of the zinc-binding groups. The comparison of several inhibitors with both carboxylate and hydroxamate zinc-binding groups demonstrates this property in fibroblast and neutrophil collagenases as well. While the potency of inhibitors with different zinc-binding groups maps directly to the number of bonds formed by the zinc-binding group, some of the increase in potency of the hydroxamate group over charged groups most likely is a result of the decreased energetic cost of the desolvation of the neutral hydroxamate.

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VIII. S1' Interactions Matrix metalloproteinase structural studies of the P'-side inhibitors to date show a common set of inhibitor-enzyme interactions. This can be attributed primarily to the strong directional zinc-binding forces. Further stabilizing forces from the backbone hydrogen-bonding patterns common to a β sheet allow for minor adjustments due to the zinc interactions to be made while maintaining a common pharmacophore. The fairly rigid constraints of binding based on the known hydroxamate inhibitors allows the use of computer-aided modeling to play a useful role in the design of specific MMP inhibitors. Exploration of the S1' pocket was carried out by docking the P1' group within the cavity followed by rounds of energy minimization. In order to maintain the integrity of the MMP structure several limitations were used. All MMP atoms that are greater than 8 Å from the docked inhibitor were frozen. The Cα atoms of all residues within 8 Å of the inhibitor were initially constrained to their original position by a 20 kcal/mol Å2 force constant that was gradually relaxed to 1 kcal/mol Å2. Strong initial constraints were also placed on the conserved hydrogen bonds and zinc-ligand interactions. This method has several advantages and disadvantages over the common static treatment of target structures. The advantages are that it more closely approximates the actual dynamic state of protein structure and is not computationally prohibitive. The disadvantages include the increased computational cost over a static enzyme target and the fact that gross structural rearrangements can still not be accounted for. The structural similarity of the active site of the MMP family allows structure-based drug design to effectively be used for those enyzmes whose structures have not been determined yet. Examination of the S1' cavities of HFC and HNC clearly indicates a path for designing inhibitors that bind preferentially (Figure 5). The cavity of HFC is mostly filled by the leucine side chain of the preferred substrate, while the S1' pocket of HNC [26] and HFS [30] remains unfilled. The sequence similarities of the gelatinases with HNC indicate that they too can accommodate a much larger P1' group. A series of compounds was designed to explore filling the long S1' tunnel of gelatinase B [41, 42]. The optimum length for binding to gelatinase B was found. There was an increase in affinity for the phenolic ethers versus the benzylic ethers of the same overall length for binding to gelatinase B, but there was no preference seen in binding to HFS. Surprisingly the phenolic ethers also showed potent binding to HFC. The size of the bottom of the S1' pocket and the differences in the preferred torsion angle for the bond to the aromatic ring both play a role in this differential binding.

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IX. S2' Interactions The interactions at the S2' site display an interesting structure-activity relationship. While the interactions do not include any hydrogen bonds, favorable stacking interactions do play a significant role in binding. A glycine residue at P2' results in a loss of three orders of magnitude in potency for otherwise identical inhibitors [41]. There is a preference for an aromatic ring at this position in natural peptide substrates [43]. Structural considerations also allow the placement of a t-butyl group here. The potency of a t-butyl glycine P2' is less than that of a phenylalanine (Table 1), but the expected gain in bioavailability brought about by shielding the amide bond from solvation effects should compensate for the loss in potency. X. Conclusions High resolution x-ray crystallographic structure determination is an essential step in structure-based drug design. The need for high resolution structural data

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Figure 6 (a) The pocket of HFC with a leucine residue in the S1' pocket. (b) The volume of the S1' pocket of HFC can change when there are favorable interactions. The binding of the (CH2)4OPh can cause Arg214 to move, thereby making room for the extended side chain.

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to develop an appropriate SAR is demonstrated in the case of the inhibitors shown in Table 1. The unusual potency of a benzylic ether for HFC was unexpected and would not have been predicted with standard docking and minimization studies (Figure 6). The differences in the potency of the various 4-substituted analogs of inhibitor 10 against HFC suggest ππ interactions are the driving force for the displacement of arginine 214. The electron-withdrawing Cl substitution decreases the affinity for HFC while increasing the affinity for HFS. The leading 4-pentyl group can not effectively interact with arginine 214 in HFC; therefore, the rearrangement does not occur. The open channel that is present in HFS and gelatinase B presents no such impediment to binding and the affinity is essentially equal to the unsubstituted form. Not all structure-based design experiments are successful. Attempts to displace the arginine residue that caps the S1' pocket of HFC by forming a salt link with a P1' carboxylate or hydroxyl moiety were unsuccessful [42]. However, these failed attempts offer some redeeming features in the refinement of parameters that can be used to evaluate the energetic potentials for displacing buried water molecules as well as the inherent desolvation energies for polar compounds. The outlook for structure-based drug design is good. The advancement in both x-ray area detectors and computer hardware will make the determination of a series of compounds bound to a target enzyme for use in SAR development commonplace in drug-discovery efforts. The continued explosion of structural studies will lead to an increased understanding of the dynamics of protein interactions, which will, in turn, lead to better docking algorithms. The combination of structural information and greater computational power will also make more accurate predictions of protein—ligand interactions possible. References 1. Birkedal-Hansen H, Moore WGI, Bodden MK, Windsor LJ, Birkedal-Hansen B, DeCarlo A, Engler JA. Matrix metalloproteinases: a review. Crit. Rev. Oral. Biol. Med. 1993; 4:197–250. 2. Beckett RP, Davidson AH, Drummond AH, Huxley P, Whittaker M. Recent advances in matrix metalloproteinase inhibitor research. DDT 1996; 1:16–26. 3. Birkdell-Hansen H. Proteolytic remodeling of extracellular matrix. Cur. Op. Cell Biology 1995; 7:728–735. 4. Stocker W, Grams F, Baumann U, Gomis-Ruth F-X, McKay DB, Bode W. The metzincins topological and sequential relations between the astacins, adamalysins, serralysins, and matrixins (collagenases) define a superfamily of zinc peptidases. Protein Sci. 1995; 4:823–840.

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5. Willenbrock F, Murphy G, Phillips IR, Brocklehurst K. The second zinc atom in the matrix metalloproteinase catalytic domain is absent in the full-length enzymes: a possible for the C-terminal domain. FEBS Lett. 1995; 358:189–192. 6. Lowry CL, McGeehan G, Levine H. Metal ion stabilization of the conformation of a recombinant 19kDa catalytic fragment of human fibroblast collagenase. Proteins 1992; 12:42–48. 7. Hodgson J. Remodeling MMPIs. Bio Technology 1995; 13:554–557. 8. Brown PD. Matrix metalloproteinase inhibitors: a novel class of anticancer agents. Adv. Enzyme Regul. 1995; 35:293–301. 9. Watson SA, Morris TM, Robinson G, Crimmin MJ, Brown PD, Hardcastle JD. Inhibition of organ invasion by matrix metalloproteinase inhibitor batimastat (BB-94) in two human colon carcinoma metastasis models. Cancer Res. 1995; 16:3629–3633. 10. Sledge GW Jr, Qulali M, Goulet R. Bone EA, Fife R. Effect of matrix metalloproteinase inhibitor batimastat on breast cancer regrowth and metastasis in athymic mice. J. Natl. Cancer Inst. 1995; 87:1546–1150. 11. Mitchel PG, Magna HA, Reeves LM, Lopresti-Morrow LL, Yocum SA, Rosner PJ, Geoghegam KF, Hambor JE. Cloning, expression, and type II collagenolytic activity of matrix metalloproteinase-13 from human osteoarthritic cartilage. J. Clin. Invest. 1996; 3:761–768. 12. Freije JMP, Diez-Ita I, Balbin M, Sanchez LM, Blaco R, Toliva J, Lopez-Otin C. Molecular cloning and expression of collagenase-3, a novel human matrix metalloproteinase produced by breast carcinomas. J. Biol. Chem. 1994; 269:16766–16773. 13. Rosenberg GA. Matrix metalloproteinases in brain injury. J. Neurotrauma 1995; 12:833–842. 14. Nomura H, Fujimoto N, Seiki M, Mai M, Okada Y. Enhanced production of matrix metalloporteinase and activation of matrix metalloproteinase 2 (gelatinase A) in human gastric carcinomas. Int. J. Cancer 1996; 69:9–16. 15. Itoh F, Yamamoto H, Hinoda Y, Imai K. Enhanced secretion and activation of matrilysin during malignant conversion of human colorectal epithelium and its relationship with invasive potential of colon cancer cells. Cancer 1996; 77:1717–1721. 16. Deb S, Gottschall PE. Increased production of matrix metalloproteinases in enriched astrocyte and mixed hippocampal cultures treated with beta-amyloid peptides. J. Neurochem. 1996; 66:1641–1647. 17. Crabbe T, Zucker S, Cockett MI, Willenbrock F, Tickle S, O'Connell JP, Scothern JM, Murphy G, Docherty AJP. Mutation of the active site glutamic acid of human gelatinase A: effects on latency, catalysis and the binding of tissue inhibitor of metalloproteinase-1. Biochemistry 1994; 33:6684–6690. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_187.html (1 of 2) [4/5/2004 5:04:47 PM]

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18. Murphy G, Allan JA, Willenbrock, F, Cockett MI, Docherty AJP. The role of the C-terminal domain in collagenase and stromelysin specificity. J. Biol. Chem. 1992; 267:9612–9618. 19. Murphy G, Willenbrock F, Ward RV, Cockett MI, Eaton D, Docherty AJP. The C-terminal domain of 72 kDa gelatinase A is not required for catalysis, but is essential for membrane activation and modulates interactions with tissue inhibitors of metalloproteinase. J. Biochem. 1992; 328:637–641.

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20. Murphy G, Docherty AJP. Assessment of the role of fibronectin-like domain of gelatinase A by analysis of a deletion mutant. J. Biol. Chem. 1994; 269:6632–6636. 21. Shipley JM, Doyle GA, Fliszar CJ, Ye QZ, Johnson LL, Shapiro SD, Welgus HG, Senior RM. The structural basis for the elastolytic activity of the 92-kDa and 72-kDa gelatinases. Role of the fibronectin type II-like repeats. J. Biol. Chem. 1996; 271:4335–4341. 22. Spurlino, J, Smallwood AM, Carlton DC, Banks TM, Vavra KJ, Johnson JS, Cook EW, Falvo J, Wahl RC, Pulvino TA, Wendoloski JJ, Smith, DL. 1.56 Å structure of mature truncated human fibroblast collagenase. Proteins 1994; 19:98–109. 23. Lovejoy B, Cleasby A, Hassell AM, Longley K, Luther MA, Weigl D, McGeehan G, McElroy AB, Drewry D, Lambert MH, Jordan SR. Structure of the catalytic domain of fibroblast collagenase complexed with an inhibitor. Science 1994; 263:375–377. 24. Lovejoy B, Hassell AM, Luther MA, Weigl D, Jordan SR. Crystal structures of recombinant 19-kDa human fibroblast collagenase complexed to itself. Biochemistry 1994; 33:8207–8217. 25. Borkakoti N, Winkler FK, Williams DH, D'Arcy A, Broadhurst MJ, Brown PA, Johnson WH, Murray EJ. Structure of the catalytic domain of human fibroblast collagenase complexed with an inhibitor. Struct. Biol. 1994; 1:106–110. 26. Stams T, Spurlino JC, Smith DL, Wahl RC, Ho TF, Qoronfleh MW, Banks TM, Rubin B. Structure of human neutrophil collagenase reveals large S1' specificity pocket. Struct. Biol. 1994; 1:119–123. 27. Bode W, Reinemer P, Huber R, Kleine T, Schnierer S, Tschesche H. The X-ray crystal structure of the catalytic domain of human neutrophil collagenase inhibited by a substrate analogue reveals the essentials for catalysis and specificity. EMBO J. 1994; 13:1263–1269. 28. Browner MF, Smith WW, Castelhano AL. Matrilysin-inhibitor complexes: common themes among metalloproteinases. Biochemistry 1995; 34:6602–6610. 29. Wetmore DR, Hardman KD. Roles of the propeptide and metal ions in the folding and stability of the catalytic domain of stromelysin (matrix metalloproteinase 3). Biochemistry 1996; 35:6549–6558. 30. Dhanaraj V, Ye Q–Z, Johnson LL, Hupe DJ, Ortwine DF, Dunbar JB, Rubin JR, Pavvlovsky A, Humblet C and Blundell TL. X-ray structure of a hydroxamate inhibitor complex of stromelysin catalytic domain and its comparison with members of the zinc metalloproteinase superfamily. Structure 1996; 4:375–386. 31. Bernstein FC, Koetzle TF, Williams GJB, Meyer EF, Brice MD, Rodgers JR, Kennard O, Shimanouchi T, Tasumi M. The Protein Data Bank: a computer-based archival file for macromolecular structures. J. Mol. Biol. 1977; 112:535–542.

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32. Carson M. Ribbon models of macromolecules. J. Mol. Graphics 1987; 5:103–106. 33. Connolly ML. The molecular surface package J. Mol. Graphics 1993; 11:139–141. 34. Li J, O'Hare MC, Skarzynski T, Lloyd LF, Curry VA, Clark IM, Bigg HF, Hazleman BL, Cawston TE, Blow DM. X-ray structure of a hydroxamate inhibitor complex of stromelysin catalytic domain and its comparison with members of the zinc metalloproteinase superfamily. Structure 1996; 4:375–386. 35. Grams F, Crimmin M, Hinnes L, Huxley P, Pieper M, Tschesche H, Bode W. Structure determination and analysis of human neutrophil collagenase complexed with a hydroxamate inhibitor. Biochemistry 1995; 34:14012–14020.

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36. Bode W, Reinemer P, Huber R, Kleine T, Schnierer S, Tschesche H. The X-ray crystal structure of the catalytic domain of human neutrophil collagenase inhibited by a substrate analogue reveals the essentials for catalysis and specificity. EMBO J. 1994; 13:1263–1269. 37. Schwartz MA, Van Wart HE. In: Ellis GP, Luscombe DK, eds. Progress in Medicinal Chemistry, Vol. 29. London: Elsevier Publishers, 1992: Chapter 8. 38. Johnson WH, Roberts NA, Borkakoti N. J. Enzyme Inhibition 1987; 2:1–22. 39. Wahl RC, Dunlop RP, Morgan BA. In: Bristol JA, ed. Annual Reports in Medicinal Chemistry. New York: Academic Press, 1990: Chapter 19. 40. Henderson B, Docherty AJP, Beeley NRA. Drugs of the Future 1990; 15:495–408. 41. Wahl RC, Pulvino TA, Mathiowetz AM, Ghose AK, Johnson JS, Delecki D, Cook ER, Gainer JA, Gowravaram MR, Tomczuk BE. Hydroxamate inhibitors of human gelatinase B (92kDa). Biorg. and Med. Chem. Lett. 1995; 5:349–352. 42. Gowravaram MR, Tomzcuk BE, Johnson JS, Delecki D, Cook ER, Ghose AK, Mathiowetz AM, Spurlino JC, Rubin B, Smith DL, Pulvino T, Wahl RC. Inhibition of matrix metalloproteinases by hydroxamates containing heteroatom-based modifications of the P1' group. J. Med. Chem. 1995; 38:2570–2581. 43. Netzel-Arnett S, Fields G, Birkdal-Hansen H, Avan Wart HE. Sequence specificities of human fibroblast and neutrophil collagenases. J. Biol. Chem. 1991; 206:6747–7855.

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7 Structure—Function Relationships in Hydroxysteroid Dehydrogenases Igor Tsigelny and Michael E. Baker University of California, San Diego, La Jolla, California I. Introduction Steroid hormones regulate a multitude of physiological processes in humans. Androgens and estrogens regulate sexual development and reproduction; glucocorticoids are important in the response to stress; vitamin D is important in bone growth; progestins are important for a viable fetus during pregnancy; mineralocorticoids regulate sodium and potassium balance to maintain normal blood pressure. Moreover, the growth of some breast and prostate tumors depends on steroids. With this multitude of medically important steroid-dependent actions, much research has gone into understanding their mode of action, with most of the effort concerned with the receptors that mediate the actions of steriods. A. High Blood Pressure and 11β-Hydroxysteroid Dehydrogenase It is only in the last decade that the role of hydroxysteroid dehydrogenases (Figure 1) in regulating the actions of steroids has been appreciated [1–6]. This mechanism for regulating steroid hormone action was uncovered in several laboratories studying various aspects of high blood pressure. One source was the study in the 1970s that identified the syndrome, Apparent Mineralocorticoid Excess (AME), a genetic disease that results in high blood pressure in children [7–10]. Also important is the work from laboratories investigating paradoxes in the mechanism of action of aldosterone in the kidney [1–4,9,10]. These studies identified 11β-hydroxysteroid dehydrogenase (11β-HSD) as a key enzyme.

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Figure 1 Reactions catalyzed by 11β-hydroxysteroid and 17β-hydroxysteroid dehydrogenases. (a) 11 β-hydroxysteroid dehydrogenase type 1, an NADPH-dependent enzyme, catalyzes the conversion of the inactive steroid, cortisone, to cortisol, which is the biologically active glucocorticoid. 11β-hydroxysteroid dehydrogenase type 2, an NAD+-dependent enzyme, catalyzes the reverse direction. (b) 17β-hydroxysteroid dehy-drogenase type 1, an NADPH-dependent enzyme, catalyzes the reduction of estrone to estradiol. Type 2, an NAD+-dependent enzyme, catalyzes the oxidation of estradiol to estrone. Type 3, an NADPH-dependent enzyme, catalyzes the reduction of androstene dione to testosterone. Type 4, an NAD+-dependent enzyme, catalyzes the oxidation of estradiol to estrone, and androstenediol to dehydroepiandrosterone.

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The enzyme 11β-HSD interconverts the active glucocorticoid cortisol and cortisone, an inactive metabolite (Figure 1a). By the oxidation of cortisol to cortisone, 11β-HSD prevents glucocorticoids from deleterious actions in certain cell types. For example, excess glucocorticoids in Leydig cells inhibit testosterone synthesis [3,11]. Expression of 11β-HSD in Leydig cells prevents this effect of glucocorticoids. In this way, 11β-HSD is important in androgen action. The enzyme 11β-HSD is also important in aldosterone action in the distal tubule of the kidney. Glucocorticoids have high affinity for the mineralocorticoid receptor [12] and can stimulate the mineralocorticoid response—uptake of sodium from urine—one effect of which is to increase blood pressure. Local expression of 11β-HSD in the distal tubule prevents this effect of glucocorticoids. The steroid aldosterone, which is not metabolized by 11β-HSD, can bind to the mineralocorticoid receptor and regulate sodium and potassium balance. Thus, 11β-HSD has an important role in regulating the biological actions of both glucocorticoids and mineralocorticoids. As would be expected, interference with 11β-HSD activity due to a mutation [13,14] or by an inhibitor such as licorice (Figure 2) [1–3,15] has a variety of physiological effects including high blood pressure due to mineralocorticoid actions of glucocorticoids in the kidney's distal tubule. Thus, studies to unravel genetic hypertension in children and the actions of aldosterone in the kidney yielded the general insight that, at specific times, altered expression of 11β-HSD in specific tissues is an important mechanism for regulating glucocorticoid, mineralocorticoid, and androgen action. A similar mechanism has been found for 17β-hydroxysteroid dehydrogenase (17β-HSD), the enzyme that regulates the concentrations of estradiol and testosterone in human [5,16,17] (Figure 1b). Genetics diseases associated with mutations in this enzyme lead to developmental abnormalities [18]. Enzymes that regulate the concentrations of retinoids [19] and prostaglandins [20] may also have a similar role [6]. B. Multiple Divergent 11β-Hydroxysteroid and 17β-Hydroxysteroid Dehydrogenases The cloning and sequencing of 11β-HSD [21–25] and 17β-HSD [16–18,26] revealed two 11β-HSDs and four 17β-HSDs with very different sequences

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Figure 2 Structure of licorice and carbenoxolone. Glycyrrhizic acid, a constituent of licorice extract, is found in the root of Glycyrrhiza glabra. The glycosidic group at C3 is cleaved by bacteria in the small intestine to form glycyrrhetinic acid, the compound that inhibits 11β-hydroxysteroid dehydrogenase. Carbenoxolone, a water soluble synthetic analog of glycyrrhetinic acid, is widely used to regulate 11β-HSD in vitro and in vivo.

(Figure 3, Table 1). This was surprising, as one would expect the two 11β-HSDs to be similar because they recognize the same substrates. Instead, the two 11β-HSDs have less than 20% sequence identity, after including gaps in the alignment (Table 1). The same degree of sequence divergence is found in the four 17β-HSDs [6]. This sequence divergence is reflected in differences in their catalytic properties. For example 11β-hydroxysteroid dehydrogenase-type 1 (11β-HSD-1) is an NADPH-dependent enzyme that converts cortisone to cortisol, and 11β-hydroxysteroid dehydrogenase-type 2 is an NAD+-dependent enzyme that oxidizes cortisol to cortisone. The enzyme 17βHSD-1 is an NADPH-dependent enzyme that converts estrone to estradiol, and 17βHSD-2 is an NAD+-dependent enzyme that oxidizes estradiol to estrone and testosterone to androstenedione.

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Figure 3 Alignment of 11β-and 17β-hydroxysteroid dehydrogenases. As seen in this Figure and Table 1, the sequences of the two 11β-HSDs and four 17β-HSDs are very divergent. Boxes denote sites where either 5 or 6 residues are conserved, which are likely to be functionally important.

There is considerable interest in understanding the structural bases for these differences because this information would be very useful in designing steroids and other compounds to selectively regulate the activity of one or more steroid dehydrogenases as a means of treating hormone-responsive diseases. There is precedent for this kind of treatment in the use of licorice extract from the root of the plant Glycyrrhiza glabra [15,27] to treat Addison's disease, http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_195.html (1 of 2) [4/5/2004 5:05:15 PM]

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Figure 3 (Continued)

which is characterized by insufficient levels of cortisol. Licorice inhibits 11β-HSD-2, which raises the circulating levels of cortisol and provides some relief from the symptoms of Addison's disease. This use of licorice is an example of a plant-derived compound having important uses in mammalian steroid hormone physiology and indicates another reason why elucidation of the structure of steroid dehydrogenases is of medical interest. Plants contain a wide variety of compounds, many of which have been purified and had their structures determined; however, we don't know which of these compounds inhibit steroid

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Page 197 Table 1 Percent Identity Between Hydroxysteroid Dehydrogenase Sequences Shown in Figure 3

11β-HSD-2

11β-HSD-2

17β-HSD-2

17β-HSD-3

17β-HSD-1

11β-HSD-1

17β-HSD-4

0.00

46.1

20.3

28.6

20.6

18.1

0.0

21.1

21.2

17.7

18.9

0.0

19.1

19.6

17.5

21.1

20.4

0.0

17.1

17β-HSD-2 17β-HSD-3 17β-HSD-1 11β-HSD-1

0.0

17β-HSD-4

0.0

dehydrogenases. Knowledge of structure-activity relationships for the binding site on steroid dehydrogenases will be helpful in identifying novel compounds from plants and other sources that could be useful in regulating steroid dehydrogenases. At this time, we are just beginning to work on this ambitious goal. Structural information is limited. The 3-D structure of 17β-HSD type 1 has been determined [28], but without the steroid or cofactor in the binding site. Fortunately, 11β-HSD and 17β-HSD belong to a large family of enzymes that are called short-chain alcohol dehydrogenases [29–31] or sec-alcohol dehydrogenases [32]. The structures of dehydrogenase homologs in bacteria, plants, and animals have been determined [33–37] and we used them as templates for modeling 11β-HSD and 17β-HSD [38,39]. There also is information about the effects of mutations on catalytic activity in 11β-HSD-1 [40] and 17β-HSD-1 [41] and in homologs, especially for Drosophila alcohol dehydrogenase (ADH) [42–48]. Together they enable us to begin to understand the relationship between structure and function in hydroxysteroid dehydrogenases, as we discuss in this chapter. II. Methods A. Molecular Modeling

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Important for the validity of the models that we constructed is the evidence from models of other proteins indicating that two proteins can have as little as 20 to 25% sequence identity and still have very similar 3D structures, especially in α helices and β stands [49–52]. Variation is found in the loops and coiled structures. A relevant example for this chapter is the comparison of the tertiary structure of rat dihydropteridine reductase [33] and Streptomyces hydrogenans 20β-hydroxysteroid dehydrogenase [34]. As noted by Varughese et al. [33], despite less than 20% sequence identity between dihydropteridine reductase and

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Figure 4 Amino acids important in cofactor and catalysis in human 11b-hydroxysteroid dehydrogenase types 1 and 2. (a) 11b-HSD type 1. Preference of 11b-HSD type 1 for NADPH resides in lysine-44 and arginine-66, which have positively charged side chains that stabilize the binding of the 2'-phosphate on NADPH. These residues also counteract the repulsive interaction between glutamic acid 69 and the phosphate group. (b) 11b-HSD type 2. Preference of 11b-HSD type 2 for NAD+ is due to favorable bonds with aspartic acid-91, serine-92, and threonine-112. Moreover there is a coulombic repulsion between aspartic acid-91 and NADP+, which destabilizes binding of NADP+ 11b-HSD type 2 lacks a nearby amino acid with a positively charged side chain that could diminish the repulsive interaction between NADP+ and aspartic acid-91. Also shown are threonine residues that could hydrogen bond to nicotinamide's carboxamidemoiety.

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20β-hydroxysteroid dehydrogenase, the root mean square deviation for the two tertiary structures is 2 Å over 160 Cα carbon atoms. We aligned human 11β-HSD-1 [21] and 11β-HSD-2 [22] with S. hydrogenans 20β-hydroxysteroid dehydrogenase and Escherichia coli 7α-hydroxysteroid dehydrogenase [37] for 3D modeling. Human 11β-HSD has extra segments at the amino terminus and carboxyl terminus. Previously reported alignments [30,31,53] were used to find the core structure consisting of about 225 residues that are structurally similar to the template. The first 190 residues of the 255 residues are reasonably well conserved among the hydroxysteroid dehydrogenases. Alignment of the C-terminal 65 residues is less certain as this part contains gaps and insertions. Fortunately, the core 190 residues contains the catalytic site and the cofactor binding site. We also superimposed the two 11β-HSD structures on mouse carbonyl reductase [37]. The 11β-HSD 3D structures superimpose nicely on α helices E and F and other helices and strands that are important in binding of cofactor and substrate. Then, we extracted NADPH from carbonyl reductase and NAD+ from 7α-hydroxysteroid dehydrogenase and inserted the cofactors into the structures of the two 11β-HSDs. The α helix F in 17β-HSD-1 [16], 17β-HSD-2 [17], 17β-HSD-3 [18], and porcine 17β-HSD-4 [26] was constructed by modeling on α helix F in 20β-hydroxysteroid dehydrogenase. Comparisons with other 3D structures [33–37] have demonstrated that this α helix is highly conserved. The modeled dimers were not minimized as a dimer complex to avoid the artifactual adjustment of α helix F side chains. The Homology program (Biosym Technologies, Inc., 1995) was used to model a 255-residue segment of 11β-HSD and α helix F of 17β-HSD on the S. hydrogenans 20β-hydroxysteroid dehydrogenase template. To produce the final model (Figures 4 and 5) this program finds an optimal configuration of the residues when arranged in the template structure by minimizing unfavorable interactions between amino acid side chains. The side chains of each monomer were then minimized (1,000 iterations of the conjugate gradient) using the Discover program (Biosym Technologies Inc., 1995).

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Figure 5 Structure of α helix F interface of human 11β-hydroxysteroid dehydrogenase types 1 and 2. The α helix F part of the dimer interface on 11β-HSD-1 and -2 is shown along with side chains of the highly conserved tyrosine and lysine residues and other residues that are oriented into the cavity that binds substrate and nucleotide cofactor.

III. Results and Discussion A. NADPH Binding Site on 11β-Hydroxysteroid Dehydrogenase Types 1 and 2 Several lines of evidence—sequence analysis, mutagenesis studies, and the solved 3D structure of homologs of 11β-HSD—indicate that the nucleotide binding site in these enzymes has many similarities to that in other classes of dehydrogenases. For many dehydrogenases, the nucleotide binding domain

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consists of a β strand α helix, β strand in a fold that provides a hydrophobic pocket for the adenosine monophosphate (AMP) part of the nucleotide cofactor [51,54,55]. In short-chain alcohol dehydrogenases, this βαβ fold is at the amino terminus. The turn between the first β strand and the α helix is a glycine-rich segment of the form Gly-X-X-X-Gly-X-Gly. This glycine-rich segment forms a hydrophobic pocket that allows close association of the AMP part of the cofactor. However, this glycine-rich segment has other functions in short-chain alcohol dehydrogenases. Tanaka et al. [37] and our 3D modeling [60] indicate that this glycine rich segment has an important role in cofactor specificity and binding of the nicotinamide moiety to 11β-HSD. B. 11β-HSD-1 Preference for NADPH Figure 4 shows our 3D model of human 11β-HSD types 1 and 2. These models identify residues important in preference of 11β-HSD-1 for NADPH and 11β-HSD-2 for NADH. In 11β-HSD type 1, lysine-44 and arginine-66 have favorable coulombic interactions with the 2'-phosphate on NADP+ that stabilize binding (Figure 4a). Moreover, their positively charged side chains compensate for the negative interaction between glutamic acid-69 and the 2'-phosphate group. Tanaka et al. [37] found a similar function for lysine-14 and arginine-39 in the preference of mouse carbonyl reductase for NADPH. C. 11β-HSD-2 Preference for NAD+ The 3D structure of 11β-HSD-2 shows that NAD+ has stabilizing interactions between the ribose hydroxyl and aspartic acid-91, serine-92, and threonine-112. Replacement of NAD+ with NADP+ reveals a coulombic repulsion between the 2'-phosphate group and aspartic acid-91. However, 11β-HSD type 2 lacks a nearby amino acid with a positively charged side chain that could compensate for the negative charge on aspartic acid-91. This explains the preference of 11β-HSD-2 for NAD+. D. Amino Acids Important in Binding the Nicotinamide Ring and Carboxamide Moiety Both 11β-HSD types 1 and 2 contain residues in the C-terminal half that interact with the nicotinamide ring and carboxamide moiety to limit rotations about the N-glycosidic bond. These intersections are important in positioning the cofactor for proS hydride transfer at C4. In 11β-HSD-1, cysteine-213 stabilizes the nicotinamide ring; threonine-220 and threonine-222 stabilize the carboxamide moiety. In 11β-HSD-2, there are more interactions: proline-262, phenylalanine-265, threonine-267, serine-

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269, and valine-270 are close to either the nicotinamide ring or the carboxamide moiety. In addition, the face of the side chain of phenylalanine-94 is below the nicotinamide ring and its carboxamide group. This latter interaction is unusual because phenylalanine-94 is between the two canonical glycine residues in the βαβ fold, which is usually thought of as interacting mainly with the AMP part of the cofactor. In 11β-HSD-2, there is an interesting configuration of amino acids with aromatic side chains that are below the nicotinamide ring and which provide a hydrophobic cushion for NAD+. E. Catalytic Site Comparison of 11β-HSD-1 with homologs identifies tyrosine-183 and lysine-187 as being highly conserved residues. Mutagenesis of these residues [40] and the homologous tyrosine and lysine in 17βHSD-1 and Drosophila alcohol dehydrogenase [44,45] shows that these residues are important for catalytic function. The 3D model of 11β-HSD-1 presented in Figure 4 shows that tyrosine-183 is 3.6 Å from the nicotinamide C4, where hydride transfer occurs. Similarly, in 11β-HSD-2, tyrosine-232 is 4 Å from C4 on NAD+. Their positions support the notion that tyrosine is the catalytically active residue. However, a problem with this model is that the pKa of tyrosine is about 10, which would make this residue a poor nucleophile at neutral pH. To resolve this problem for the homologous tyrosine in Drosophila alcohol dehydrogenase, Chen et al. [44] proposed that the pKa of tyrosine is lowered by a nearby positively charged lysine. The 3D structure of the two 11β-HSDs shows that lysine-187 and lysine-236 are close to the proposed catalytically active tyrosine residues, which supports the hypothesis of Chen et al. [44]. F. Dimer Interface and the Catalytic Site Most short-chain alcohol dehydrogenases are active as either dimers or tetramers. Analysis of rat dihydropteridine reductase by Varughese et al. [33] indicates that the dimer interface consists of α helix E and α helix F from each monomer arranged in a four α helix bundle, a structure in which the hydrophobic surfaces on the helices form a core that yields very stable structure in a wide variety of proteins [56–59]. A four-helix bundle also appears to stabilize S. hydrogenans 20β-hydroxysteroid dehydrogenase, a tetrameric enzyme [34]. The α helix F contains the conserved tyrosine and lysine residue, which adds a constraint to changes in the sequence of this helix. It has at least two functions: stabilizing the dimer and orienting tyrosine and lysine and other residues for optimal interaction with substrate and nucleotide cofactor. The role of a specific site on the outer hydrophobic surface of α helix F in dimerization was suggested recently when a Drosophila ADH mutant that does

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not form stable dimers was sequenced [48]. This ADH mutant has alanine-159 replaced with threonine. A 3D model of Drosophila ADH shows alanine-159 on the opposite surface of α helix F from tyrosine153 and lysine-157 [48]. Alanine-159 along with alanine-158 form a hydrophobic anchor that stabilizes the dimer interface. These two residues of ADH and the homologous residues in other short-chain alcohol dehydrogenases have been overlooked in sequence analyses because they are not absolutely conserved. In fact, at least five amino acids are found in these positions among the different sec-alcohol dehydrogenases. G. Dimer Interface in 11β-and 17β-Hydroxysteroid Dehydrogenases Because α helix F at the dimer interface also contains the catalytic tyrosine and the nearby lysine residue, any structural analysis of the catalytic site must also consider the structure of this part of the dimer interface. For this reason, we modeled α helix F on 11β-HSD-1 and -2 and 17β-HSD-1, -2, -3, and -4 to gain an insight into stabilizing interactions and how they may affect the catalytic site. H. Human 11β-HSD-1 Figure 5 shows the modeled structure for the α helix F interface in human 11β-HSD-1, in which phenylalanine-188 and alanine-189 form an anchor. Alanine-189 is 3.5 Å and 4.7 Å from alanine-189 and alanine-185, respectively, on the other subunit. The phenylalanine-188 side chain is 3.2 Å from glycine-192. There is a hydrogen bond between serine-185 and serine-196, which are 3.2 Å apart. Alanine-185 is 4.7 Å from phenylalanine-193. There also is a hydrophobic interaction between phenylalanine-193 and alanine-181, which are 3.9 Å apart. This web of interaction between side chains on the outer surface of α helix F on each subunit influences residues that have side chains oriented to the interior where catalysis occurs. Alanine-185, which is stabilized by interactions with phenylalanine-193, and serine-184, which interacts with serine-196, are between the conserved tyrosine-183 and lysine-187. Phenylalanine-193 is adjacent to phenylalanine194, which is positioned into the catalytic site. I. Human 11β-HSD-2 Figure 5 shows the α helix F interface of human 11β-HSD-2. Alanine-237 is about 3 Å from leucine241; alanine-238 is about 3.7 Å from both alanine-238 and threonine-234. Threonine-234 has a stabilizing hydrophobic interaction

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Figure 6 Structure of α helix F interface of mammalian 17β-hydroxysteroid dehydrogenases. The α helix F part of the dimer interface on 17β-hydroxysteroid dehydrogenases is shown along with side chains of the highly conserved tyrosine and lysine residues and three other residues that are oriented into the cavity that binds substrate and nucleotide cofactor. (a) Modeled structure of human 17β-hydroxysteroid dehydrogenase type 1. (b) Modeled structure of human 17β-hydroxysteroid dehydrogenase type 2. (c) Modeled structure of human 17β-hydroxysteroid dehydrogenase type 3. (d) Modeled structure of porcine 17β-hydroxysteroid dehydrogenase type 4.

with leucine-242, which is 4.2 Å distant. Threonine-245 is 4.2 Å from the Cα carbon of glycine-230.

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The type 1 and type 2 enzymes preferentially catalyze the reduction of the 11-keto group and the oxidation of the 11-hydroxyl group, respectively, on glucocorticoids. The chemistry of the side chains on methionine-243 in the type-2

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enzyme and of phenylalanine-194 on 11β-hydroxysteriod dehydrogenase type 1 is quite different; it may be important in the different catalytic properties of these two enzymes. J. Human 17β-HSD-1 Figure 6a shows the modeled α helix F interface in human 17β-hydroxysteroid dehydrogenase type 1 in which phenylalanine-160 and alanine-161 form an anchor. Both residues have important stabilizing interactions across the dimer interface. Alanine-161 is 4.1 Å from alanine-161 on the other subunit. Alanine-161 has a hydrophobic interaction with alanine-157, which is in the segment between the conserved tyrosine-155 and lysine-159. There is a hydrophobic

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interaction between alanine-157 and leucine-165, which are about 3.8 Å apart. Phenylalanine-160 is 4 Å from glycine-164. There also is a hydrogen bond between cysteine-156 and serine-168, which are 3.2 Å apart. This is an interesting structural property of residues in the segment between the conserved tyrosine and lysine residues: this segment is important in stabilizing dimers. This pattern is repeated in the other 11β- and 17β-hydroxysteroid dehydrogenases suggesting conservation of this stabilizing structure, although the residues are not as well conserved as the tyrosine and lysine. K. Human 17β-HSD-2 Figure 6b shows the modeled α helix F interface in human 17β-hydroxysteroid dehydrogenase type 2. Alanine-237 is 3 Å from the hydrophobic part of the side chain of methionine-241 on the other subunit. Methionine-241 is 3.2 Å from serine-234. Alanine-230 is 3.7 Å from phenylalanine-242 and 4.5 Å from valine-245. Alanine-238, the other anchoring residue, is 4.1 Å from alanine-238 on the other subunit. L. Human 17β-HSD-3 Figure 6c shows the modeled α helix F interface in human 17β-hydroxysteroid dehydrogenase-type 3. Alanine-203 is 3.1 Å from alanine-207. Phenylalanine-204 is 3.2 Å from the other phenylalanine-204 and alanine-200. These are the only stabilizing interactions that we find in our analysis. Human 17βhydroxys-teroid dehydrogenase type 3 has the weakest interactions across the α helix F interface among the four types of 17β-hydroxysteroid dehydrogenases. The conformation of this part of 17βhydroxysteroid dehydrogenase type 3 could change upon binding of substrate, leading to other stabilizing interactions. And, of course other parts of the protein may have intersubunit interactions that stabilize the dimer. Alternatively, the hydrophobic surface of α helix F may interact with another protein or a membrane surface, a potentially important regulatory mechanism that we discuss later in this paper. M. Pig 17β-HSD-4 Figure 6d shows the modeled α helix F interface in pig 17β-hydroxysteroid dehydrogenase type 4. Leucine-169 is 2.9 Å from glycine-173. Leucine-174 is 4.3 Å from alanine-162 and 3.3 Å from alanine166. There also is a hydrogen bond between serine-165 and serine-175, which are 3 Å apart. N. Prospects for the Application of Structure-Function Analysis of Steroid Dehydrogenases in Hormone Therapy In the last two years there has been impressive progress in understanding the structure of steroid dehydrogenases that are important in regulating blood pres

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sure and the actions of reproductive hormones. This progress has come from several directions. First, the cloning and sequencing the dehydrogenases that regulate the actions of aldosterone, cortisol, estradiol, and testosterone. Second, determination of the 3D structure of 17β-HSD-1 and several homologs. Analyses of their 3D structures confirm a general principle that structural similarity is much higher than sequence similarity. This supports proposed molecular models of medically important steroid dehydrogenases using the alignment of their sequences onto the templates of 3D structural homologs. Models of 11β-HSD-1 and -2 are beginning to reveal important properties about these enzymes. We now have a good picture of the structural basis for specificity for NADPH and NADH in 11β-HSD-1 and -2. With this information, we can now turn our attention to modeling cortisol in these two enzymes. This information will open up the possibility for developing analogs to regulate the actions of these two enzymes for use in regulating blood pressure and other physiological processes. The development of carbenoxolone, a water-soluble synthetic analog of glycyrrhetinic acid, shows that chemists can create compounds that have high affinity for 11β-HSD. The next task is to synthesize compounds that are specific for 11β-HSD-1 or 11β-HSD-2. Molecular modeling can contribute important information to solving this kind of problem. Similarly important information will come from the 3D models of 17β-HSD-1, -2, -3 and -4. These models will be useful in developing compounds to regulate estrogen and androgen action, which have important application in reproductive medicine and in treating estrogen-dependent breast tumors and androgen-dependent prostatic tumors. Many compounds in plants have estrogenic and androgenic activity; some of these compounds are likely to work via inhibition of one of the 17β-HSD enzymes [27]. Analogous to the development of carbenoxolone to regulate 11β-HSD, we can seek synthetic compounds that regulate specific types of 17β-HSD, which may be useful in reproductive medicine and in treating cancers. Considering the explosive pace of biomedical research and the new developments in computers for sophisticated structural analyses, the next few years promise to yield important advances in design of new hormone therapies based on the knowledge of the structure of steroid dehydrogenases. Acknowledgments We thank Drs. Tanaka, Nonaka, Nakanishi, Deyashiki, Hara, and Mitsui for providing us with the x-ray crystallographic coordinates of carbonyl reductase and 7α-hydroxysteroid dehydrogenase. The support of the Supercomputer Center of the University of California, San Diego is gratefully acknowledged.

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References 1. Funder JW, Pearce PT, Smith R, Smith AI. Mineralocorticoid action: target tissue specificity is enzyme, not receptor, mediated. Science 242; 1988:583–586. 2. Edwards CRW, Stewart PM, Burt D, Brett L, McIntyre MA, Sutanto WS, De Kloet ER, Monder C. Localization of 11β-hydroxysteroid dehydrogenase-tissue specific protector for the mineralocorticoid receptor. Lancet 2; 1988:986–989. 3. Monder C. Corticosteroids, receptors, and the organ-specific functions of 11β-hydroxysteroid dehydrogenase. FASEB J5; 1991:3047–3054. 4. White PC. Disorders of aldosterone biosynthesis and action. New Eng J Med 331; 1994:250–258. 5. Andersson S. 17β-hydroxysteroid dehydrogenase: isozymes and mutations. J Endocrinol 146; 1995:197–200. 6. Baker ME. Unusual evolution of 11β- and 17β-hydroxysteroid and retinol dehydrogenases. Bioessays 18; 1996:63–70. 7. New MI, Levine LS, Biglieri EG, Pareira J, Ulick S. Evidence for an unidentified steroid in a child with apparent mineralocorticoid hypertension. J Clin Endocrinol Metab 44; 1977:924–933. 8. Ulick S, Levine LS, Gunczler P, Zanconato G, Ramirez LC, Rauh W, Rosler A, Bradlow HL, New MI. A syndrome of apparent mineralocorticoid excess associated with defects in the peripheral metabolism of cortisol. J Clin Endocrinol Metab 49; 1979:757–764. 9. Funder JW, Pearce PT, Myles K, Roy LP. Apparent mineralocorticoid excess, pseudohypoaldosteronism, and urinary electrolyte excretion: toward a redefinition of mineralocorticoid action. FASEB J 4; 1990:3234–3238. 10. Stewart PM, Edwards CRW. The cortisol-cortisone shuttle and hypertension. J Steroid Biochem Molec Biol 40; 1991:501–509. 11. Monder C. Comparative aspects of 11β-hydroxysteroid dehydrogenase. Testicular 11βhydroxysteroid dehydrogenase: development of a model for the mediation of Leydig cell function by corticosteroids. Steroids 59; 1994:69–73. 12. Arriza JL, Weinberger C, Cerelli G, Glaser TM, Handelin BL, Housman DE, Evans RM. Cloning of the human mineralocorticoid receptor complementary DNA: structural and functional kinship with the glucocorticoid receptor. Science 237; 1987:268–275.

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13. Wilson RC, Krozowski ZS, Li K, Obeyesekere VR, Razzaghy-Azar M, Harbison MD, Wei JQ, Shackleton CHL, Funder JW, New MI. A mutation in the HSD11B2 gene in a family with apparent mineralocorticoid excess. J Clin Endocrinology Metab 80; 1995:2263–2266. 14. Mune T, Rogerson FM, Nikkila H, Agarwal AK, White PC. Human hypertension caused by mutations in the kidney isozyme of 11β-hydroxysteroid dehydrogenase. Nature Genetics 10; 1995:394–399. 15. Baker ME. Licorice and enzymes other than 11β-hydroxysteroid dehydrogenase. Steroids 59; 1994:136–141. 16. Peltoketo H, Isomaa V, Vihko R. Genomic organization and DNA sequences of human 17βhydroxysteroid dehydrogenase genes and flanking regions. Eur J Biochem 209; 1992:459–466. 17. Wu L, Einstein M, Geissler WM, Chan HK, Elliston KO, Andersson S. Expression cloning and characterization of human 17β-hydroxysteroid dehydrogenase type 2,

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a microsomal enzyme possessing 20α-hydroxysteroid dehydrogenase activity. J Biol Chem 268; 1993:12964–12969. 18. Geissler WM, Davis DL, Wu L, Bradshaw KD, Patel S, Mendonca BB, Elliston KO, Wilson JD, Russell DW, Andersson S. Male pseudohermaphrodites, caused by mutations to testicular 17βhydroxysteroid dehydrogenase-3. Nature Genetics 7; 1994:34–39. 19. Napoli JL, Boerman MHEM, Chai X, Zhai Y, Fiorella PD. Enzymes and binding proteins affecting retinoic acid concentrations. J Ster Biochem Molec Biol 55; 1995:589–600. 20. Baker ME. Evolution of enzymatic regulation of prostaglandin action: novel connections to regulation of human sex and adrenal function, antibiotic synthesis and nitrogen fixation. Prostaglandins 42; 1991:391–407. 21. Tannin GM, Agarwal AK, Monder C, New MI, White PC. The human gene for 11β-hydroxysteroid dehydrogenase. J Biol Chem 266; 1991:16653–16658. 22. Albiston AL, Obeyesekere VR, Smith RE, Krozowski ZS. Cloning and tissue distribution of the human 11β-hydroxysteroid dehydrogenase type 2 enzyme. Mol Cell Endocrinol 105; 1994:R11–R17. 23. Agarwal AK, Mune T, Monder C, White PC. NAD+-dependent isoform of 11β-hydroxysteroid dehydrogenase. Cloning and characterization of cDNA from sheep kidney. J Biol Chem 269; 1994:25959–25962. 24. Naray-Fejes-Toth A, Fejes-Toth G. Expression cloning of the aldosterone target cell-specific 11βhydroxysteroid dehydrogenase from rabbit collecting duct cells. Endocrinology 136; 1995:2579–2586. 25. Cole TJ. Cloning of the mouse 11β-hydroxysteroid dehydrogenase type 2 gene: tissue specific expression and localization in distal convoluted tubules and collecting ducts of the kidney. Endocrinology 136; 1995:4693–4696. 26. Leenders F, Adamski J, Husen B, Thole, HH, Jungblut PW. Molecular cloning and amino acid sequence of the porcine 17β-estradiol dehydrogenase. Eur J Biochem 222; 1994:221–227. 27. Baker ME. Endocrine activity of plant-derived compounds: an evolutionary perspective. Proc Soc Exper Biol Med 208; 1995: 131–138. 28. Ghosh D, Pletnev VZ, Zhu D-W, Wawrkak Z, Duax WL, Pangborn W, Labrie F, Lin S–X. Structure of human estrogenic 17β-hydroxysteroid dehydrogenase at 2.20 Aº resolution. Structure 3; 1995:503–513.

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29. Baker ME. Genealogy of regulation of human sex and adrenal function, prostaglandin action, snapdragon and petunia flower colors, antibiotics, and nitrogen fixation: functional diversity from two ancestral dehydrogenases. Steroids 56; 1991:354–360. 30. Persson B, Krook M, Jornvall H. Characteristics of short-chain alcohol dehydrogenases and related enzymes. Eur J Biochem 200; 1991:537–543. 31. Krozowski Z. 11β-hydroxysteroid dehydrogenase and the short chain alcohol dehydrogenase (SCAD) superfamily. Mol Cell Endocrinol 84; 1992:C25–C31. 32. Baker ME. Protochlorophyllide reductase is homologous to human carbonyl reductase and pig 20βhydroxysteroid dehydrogenase. Biochem J 300; 1994:605–607. 33. Varughese KI, Xuong NH, Kiefer PM, Matthews DA, Whiteley JM. Structural and mechanistic characteristics of dihydropteridine reductase: a member of the Tyr- (Xaa)3-Lys-containing family of reductases and dehydrogenases. proc Natl Acad Sci USA 91; 1994:5582–5586.

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34. Ghosh D, Wawrzak Z, Weeks CM, Duax WL, Erman M. The refined three-dimensional structure of 3α,20β-hydroxysteroid dehydrogenase and possible roles of the residues conserved in chart-chain dehydrogenases. Structure 2; 1994:629–640. 35. Dessen A, Quemard A, Blanchard JS, Jacobs Jr, WR, Sacchettini JC. Crystal structure and function of the isoniazid target for Mycobacterium tuberculosis. Science 267; 1995:1638–1641. 36. Rafferty JB, Simon JW, Baldock C, Artymiuk PJ, Stuitje AR, Slabas AR, Rice DW. Common themes in redox chemistry emerge from the X-ray structure of oilseed rape, Brassica napus, enoyl acyl carrier protein reductase. Structure 3; 1995:927–938. 37. Tanaka N, Nonaka T, Nakanishi M, Deyashiki Y, Hara A, Mitsui Y. Crystal structure of the ternary complex of mouse lung carbonyl reductase at 1.8 Å resolution: the structural origin of coenzyme specificity in the short-chain dehydrogenase/reductase family. Structure 4; 1996:33–45. 38. Tsigelny I, Baker ME. Structures stabilizing the dimer interface on human 11β-hydroxysteroid dehydrogenase-types 1 and 2 and human 15-hydroxyprostaglandin dehydrogenase and their homologs. Biochem Biophys Res Commun 217; 1995:859–868. 39. Tsigelny I, Baker ME. Structures important in mammalian 11β-and 17β-hydroxysteroid dehydrogenases. J Ster Biochem Molec Biol 55; 1995:589–600. 40. Obeid J, White PC. Tyr-179 and lys-183 are essential for enzymatic activity of 11β-hydroxysteroid dehydrogenase. Biochem Biophys Res Comm 188; 1992:222–227. 41. Puranen TJ, Poutanen MH, Peltoketo HE, Vihko PT, Vihko RK. Site-directed mutagenesis of the putative active site of human 17β-hydroxysteroid dehydrogenase type 1. Biochem J 304; 1994:289–293. 42. Chen Z, Lu L, Shirley M, Lee WR, Chang SH. Site-directed mutagenesis of glycine-14 and two “critical” cysteinyl residues in Drosophila alcohol dehydrogenase. Biochemistry 29; 1990:1112–1118. 43. Chen Z, Lin Z-G, Lee WR, Chang SH. Role of aspartic acid-38 in the cofactor specificity of Drosophila alcohol dehydrogenase. Eur J Biochem 202; 1991:263–267. 44. Chen Z, Jiang JC, Lin Z-G, Lee WR, Baker ME, Chang SH. Site-specific mutagenesis of Drosophila alcohol dehydrogenase: evidence for involvement of tyrosine-152 and lysine-156 in catalysis. Biochemistry 32; 1993:3342–3346. 45. Cols N, Marfany G, Atrian S, Gonzalez-Duarte R. Effect of site-directed mutagenesis on conserved positions of Drosophila alcohol dehydrogenase. FEBS Lett 319; 1993:90–94.

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46. Ribas dePoplana L, Fothergill-Gilmore LA. The active site architecture of a short chain dehydrogenase defined by site-directed mutagenesis and structure modeling. Biochemistry 33; 1994:7047–7055. 47. Chen Z, Tsigelny I, Lee WR, Baker ME, Chang SH. Adding a positive charge at residue 46 of Drosophila alcohol dehydrogenase increases cofactor specificity for NADP+. FEBS Lett 356; 1994:81–85. 48. Chenevert S, Fossett N, Lee WR, Tsigelny I, Baker ME, Chang SH. Amino acids important in enzyme activity and dimer stability for Drosophila alcohol dehydrogenase. Biochem J 308; 1995:419–423. 49. Chothia C, Lesk AM. The relation between the divergence of sequence and structure in proteins. EMBO J 5; 1986:823–826.

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50. Greer J. Comparative modeling of homologous proteins. Methods Enzymol 202; 1991:239–252. 51. Branden C, Tooze J. Introduction to protein structure. New York: Garland Publishing, 1991. 52. Ring CS, Cohen FE. Modeling protein structures: construction and their applications. FASEB J 7; 1993:783–790. 53. Baker ME. Sequence analysis of steroid-and prostaglandin- metabolizing enzymes: application to understanding catalysis. Steroids 59; 1994:248–258. 54. Wierenga RK, De Maeyer MC, Hol WGJ. Interaction of pyrophosphate moieties with α-helixes in dinucleotide binding proteins. Biochemistry 24; 1985:1346–1357. 55. Wierenga RK, Terpstra PP, Hol WGJ. Prediction of the occurrence of the ADP-binding βαβ-fold in proteins, using an amino acid sequence fingerprint. J Mol Biol 187; 1986:101–107. 56. Weber P, Salemme FR. Structural and functional diversity in four-α-helical proteins. Nature 287; 1980:82–84. 57. Chou K-C, Maggiora GM, Nemethy G, Scheraga HA. Energetics of the structure of the four-α-helix bundle in proteins. Proc Natl Acad Sci USA 85; 1988:4295–4299. 58. Presnell SR, Cohen FE. The topological distribution of four-α-helical proteins. Proc Natl Acad Sci USA. 86; 1989:6592–6596. 59. Harris NL, Presnell SR, Cohen FE. Four helix bundle diversity in globular proteins. J Mol Biol 236; 1994:1356–1368.

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8 Design of ATP Competitive Specific Inhibitors of Protein Kinases Using Template Modeling Janusz M. Sowadski,* Charles A. Ellis,* Rolf Karlsson* University of California, San Diego, La Jolla, California Madhusudan Scripps Research Institute, La Jolla, California I. Protein Kinases and Diseases The protein kinase family encompasses more than three hundred members of critically important enzymes, each one with a specific role or function within the cell. These enzymes, ATPphosphotransferases, recognize target proteins and through the phosphorylation of specific sites either activate or deactivate a particular pathway of signal transduction. Many of these signaling pathways are associated with cell surface receptors, which are located in the membranes that surround cells. The difference between the families of protein kinases is that they have different targets and generally fall into two major classes: The serine/threonine protein kinases transfer a phosphate from ATP to a serine or threonine residue in the target protein. This class of enzymes are generally associated with cytoplasmic signaling events. The tyrosine protein kinases transfer phosphate from ATP to tyrosine residues in the target protein and are generally associated with receptors that become activated after binding a growth factor or other ligand. Protein kinases are significant targets for therapeutic drug development and have been implicated as the disease causing components of numerous tumor *Current

affiliation: Tufts University, Boston, Massachusetts.

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viruses. Specifically, it is the deregulation of the activity of protein kinases that leads to disease by tumor viruses. The importance of this deregulation can be dramatically illustrated by the large number of viral oncogenes (or cancer causing genes) that encode structurally modified protein kinases. These deregulated enzymes are able to bypass the normal tightly regulated processes of growth control, leading to acute malignant transformation. These oncogenes are one of the first examples of the identification of disease-causing genes. Many of these viral genes have subsequently been implicated in human diseases. Malignant tissues also share the common characteristic of an acquired independence from controls. The receptor—for example, PDGF and EGFR—can be stimulated by a ligand coming either from the cell itself (autocrine) or from nearby tissues (paracrine). Regardless of the mechanism leading to receptor activity, the resulting kinase activity results in a cascade of signals that turn on cellular proliferation programs. Therefore, selective inhibition of receptor tyrosine kinase will block tyrosine kinase driven cell proliferation resulting in antitumor activity. In addition to cancer, a growing number of nonmalignant proliferative diseases, (e.g., psoriasis, atherosclerosis, restenosis, fibrosis, etc.) or inflammatory responses (e.g., septic shock, asthma, osteo and rheumatoid arthritis, etc.) involve dysfunctional signaling pathways. Successful development of drugs that target this class of enzymes will depend on the discovery of selective inhibitors designed for the appropriate protein kinase within the family. In the past several years there has been an explosion of structural studies within the protein kinase family [1–8]. These studies, initiated by the crystal structure of Protein Kinase A [9–12] (CAPK) have shown that all members of the protein kinase family fold into a uniform three-dimensional catalytic core. Yet this uniform three-dimensional fold exhibits both different surface charges and at least two major conformations. II. Protein Kinase Template The stereo view of the ribbon diagram of cAPK is presented in Figure 1a. The overall topology of the core extending from strand 1 through helix h, Figure 1b, is identical (except helix B) with the eight other structures of protein kinases determined to this point. Furthermore, Figure 2 presents an overall structural comparison of the catalytic cores of the five kinases, cAPK, CDK2, CDK2-CYCLIN, IR, and MAP. The N-terminal helix A, which is present only in the cAPK crystal structure, is anchored by myristic acid in the mammalian bovine heart of cAPK. Myristic acid inserts itself into the hydrophobic pocket of the lower lobe of the enzyme, which results in the structural ordering of helixA and Ser10[13], one of the four autophosphorylation sites.

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Figure 1 Diagram of cAPK fold. (a) Stereo MOLSCRIPT diagram. (b) The key loops are as follows: phosphate anchor located between strand 1 and strand 2, catalytic loop located between strands 6 and 7, DFG motif located between strands 8 and 9, activation loop including P+1 site between strand 9 and helix F. Phosphorylation site Thr197 is indicated by a large circle, inhibitor PKI (5–24) is colored in dark, and the P site in the peptide is shown in the dark circle. This figure has been generated using MOLSCRIPT [27].

Following the connectivity diagram, helix A is connected to β-strand 1, then to the phosphate anchor encompassing signature motif Gly50XGly 52XXGly55. The β-strand 2 is followed by β-strand 3 carrying invariant Lys72. Three antiparallel beta strands create the unique fold of the nucleotide binding site of the protein kinase. The β-strands 3 is followed by helix B, which is present only in cAPK, helix C, and β-strands 4 and 5. Helix C shows the largest displacements among many different protein kinase structures and consists of invariant Glu91, which forms a salt bridge with Lys72. This salt bridge is absent in the inactive CDK-2 structure [1] but present in the crystal structure of the complex of CDK-2 and its activator-cyclin [7]. Displacement of helix C is perhaps most pronounced in the case of the insulin receptor tyrosine kinase structure (IRK) [3]. In PKA, Phe185 resides in the hydrophobic pocket formed by the hydrophobic residues of helix C (upper lobe) and Tyr164 (lower lobe). In the IRK crystal structure, the hydrophobic residues of helix C, which provide the pocket for invariant Phe185 (of DFG motif), no longer interact with

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Figure 1 (Continued)

this residue. The DFG motif in IRK occupies the ATP site, which blocks the access of ATP. The division between the upper and lower lobes of the enzyme is well defined by the two major conformations of the upper lobe observed in the crystal structures of cAPK. One conformation has been observed in the orthorhombic crystals of recombinant cAPK [9,10,14] and another in the cubic

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Figure 2 The diagrams of the Cα trace of the five kinases, cAPK, CDK2, CDK2-CYCLIN, IR, MAP. The thin line represents crystallographically determined homologous regions of the five kinases (R. Karlsson and J.M. Sowadski, personal communication).

crystals of bovine heart mammalian cAPK [15,16]. A comparison between the two structures has shown that there is rotation of the upper lobe by 15 degrees and a translation of 1.9 Å in the mammalian structure, which results in the opening of the nucleotide binding cleft. The motion of this lobe, which includes His87, one of the ligands to Thr197 [15], indicates that the phosphorylation of this site will be important for conformational diversity of the upper lobe. This is confirmed by the varying degrees of displacement of the upper lobe of all structures of the inactive unphosphorylated protein kinases (see review [17]). The lower lobe of the enzyme starts with helix D, which is followed by helix E and β-strands 6 and 7. The catalytic loop connecting both strands consists of a critical set of residues with Tyr164 and Arg165 at the beginning of the loop. The Tyr164 residue forms a hydrogen bond with invariant Asp220. The Arg165 residue, which is present in a great majority of protein kinases, provides two hydrogen bonds to the oxygens of the phosphate of Thr197. Invariant Asp166 (the catalytic base) and Asn171 (the ligand to one of the metal sites) are also located within this loop.

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The catalytic loop is the region of divergence between Ser/Thr and Tyr kinases. In cAPK and all Ser/Thr Kinases, Lys168 interacts with the γ phosphate of ATP during catalysis [12]. The role of Lys is replaced by Arg [9] and the insulin receptor tyrosine kinase structure [3] shows Arg1136 in a similar position as Lys168 in the active site of cAPK. The catalytic loop and β-strand 7 are followed by β-strand 8 and a short DFG conserved motif. This conserved motif consists of invariant Asp184, a ligand to the metal site, and invariant Phe185. The DFG motif is followed by β-strand 9, an activation loop that includes Thr197. The activation loop differs considerably among unphosphorylated kinases, CDK-2, ERK-2, and insulin receptor kinase. Furthermore, two crystal structures, twitchin protein kinase [2] and phosphorylase kinase [5], both lack a phosphorylation site in this region. In the first structure, twitchin protein kinase, Thr197 and Arg165, are replaced by hydrophobic residues, Val6098 and Leu6062 respectively [2]. This was predicted using modeling [18] and subsequently confirmed by the three-dimensional structure. In the second structure, phosphorylase kinase, the Thr197 is replaced by Glu182, which interacts with Arg148 [5]. Hence, in both structures the regulatory function of the phosphorylation site is replaced by a stable scaffold secured by either hydrophobic or electrostatic interactions. Since it has been shown that this loop provides a stable template for PKI(5–24) binding in cAPK, the status of phosphorylation of the activation loop critically affects the substrate binding. This is demonstrated in c-Src, a homolog of the Rous Sarcoma virus oncogene by mutation of Arg385, which is predicted to interact with the phosphate of Tyr416, and results in loss of activity toward the exogenous substrate [19]. The activation loop is followed by a P+1 loop which accommodates the P+1 site of the substrate. The P+1 loop is followed by invariant Glu208, which forms a salt bridge with invariant Arg280. This conserved pair plays a structural role and as the structure of CK-1 [16] shows, it can be replaced by other charged residues that maintain the same fold of the lower lobe. The P+1 loop and Glu 208 are followed by helix F consisting of invariant Asp220, followed by helices G, H, and I. The helix J and the C-terminal tail of cAPK, which are absent in other protein kinases, undergo a large motion during the cleft opening. The opening of the cleft results in loss of hydrogen bonds provided by the γ phosphate of ATP and the peptide that would bridge the lower and upper lobe of the enzyme. The motion of the upper domain increases the accessibility of the ATP binding site and one can envision that in the “open” conformation ATP binds. Yet, in the “closed” conformation ATP and its γ phosphate are positioned for a nucleophilic attack on the substrate. The motion of this lobe—which includes His87, one of the ligands to Thr197 [15]—indicates that the phosphorylation of this site will be important for conformational diversity of the upper lobe. This is confirmed by the varying degrees of displacement of the upper lobe

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of all structures of the inactive unphosphorylated protein kinases (see review [17]). The various displacements of the conserved upper domain of the catalytic cores of various kinases documented by crystallographic work suggest that this is the important underlying mechanism of catalysis. Analysis of crystal contacts of various kinases is however required to define the extent of displacement due to the lattice forces. In the case of cAPK, the displacement as observed for mammalian cAPK in the cubic crystal form is due to the intermolecular interaction in the lattice [20]. Analysis of the two crystal structures of the cell cycle-controlling kinases clearly shows two binding modes of ATP. In the inactive state without cyclin, ATP binding of its triphosphate moieties is different from that in the active form with cyclin bound. The major difference is the re-arrangement upon cyclin binding of the conserved Lys33-Glu51 pair, which is responsible for the binding of the α and β phosphates of ATP. III. Crystallographic Analysis of Substrate Specificities of Individual Kinases The most important contribution of subsequent crystallographic studies has been the confirmation of the structural homology extending through the members of this family of enzymes. The crystal structures of CDK-2 [1], ERK-2 [5], twitchin [2], insulin receptor kinase [3], phosphorylase kinase, CK-1 [6], along with structure of calcium/calmodulin-dependent protein kinase I [8] provide solid proof for the structural conservation of the catalytic core in the family. This is further confirmed by the recent structure of the active complex of CDK2/cyclin, which shows that Lys33-Glu51 pair is at a distance of 3.0 Å [7] as predicted in the model of CDK-2 based on the cAPK structure [21]. The structure of the complex has also confirmed the binding of cyclin to helix C and to the upper lobe, demonstrating the mechanism of activation by cyclin that results in bridging the invariant residues into the common network of distances required by structural homology of the protein kinase catalytic core. The crystallographic analysis of the structural homology of protein kinases can now be carried out using structures of various kinases to find a common search model to be used in molecular replacement methods (J.M. Sowadski and R. Karlsson private communication). The structures of various kinases have been used as search models to solve the structure of the cAPK using cAPK diffraction data. The best search model consists of fragments of the catalytic core excluding the activation loop, inserts, and upper lobe due to rotational motion observed in each structure. A structure solution has been found for several protein kinases using this selected model as shown in Figure 2. One of the most critical aspects of this analysis is the presence of the structurally

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conserved substrate-binding cleft as observed in the crystal structure of cAPK:PKI complex (Figure la,b). This finding allows the charges within this cleft to be predicted using the amino acid sequence of any given kinase. In the analysis of the structural data of other protein kinases, it is noted that only cAPK has been crystallized with its specific peptide inhibitor. Nevertheless, three other structures of protein kinases compared with the structure of the cAPK-PKI complex provide substantial evidence for the conservation of the substrate binding cleft. The substrate binding cleft of the phosphorylase kinase structure has been analyzed in detail and it is clear that all amino acids of the known specific substrate can be built into the PKI model and all required corresponding charges can be found in the cleft of the phosphorylase kinase structure. In the CK-1 structure determined without a peptide, the requirement of the peptide specificity resides on the P-3 site, which has to be phosphorylated. An analysis of the surface charges of the cleft of the CK-1 structure reveals the exact correspondence of the residues required to interact with a phosphorylated substrate at this site. Finally, the tripeptide of the pseudosubstrate site of IRK consisting of the Asp-Tyr-Tyr motif has corresponding charges in the structure of the enzyme's substrate cleft and confirms the data obtained from the degenerated peptide library for the unique sequence motifs of nine tyrosine kinases [22]. It is becoming increasingly clear that the wealth of structural data of protein kinases with cAPK as a prototype provides evidence for two important features concerning substrate binding. First, the substrate binding cleft is structurally conserved and second, the surface charges of this cleft and hydrophobic cavities on the surface are very diverse and correspond to the specificity requirement of the substrate for individual protein kinases (see Figure la). It is now possible to use the structural conservation of the substrate binding cleft to predict the charges and hydrophobic residues of the cleft to define substrate specificities for individual kinases. IV. Crystallographic Analysis of the ATP Binding Site Reveals Distinct Differences Utilized for the Further Design of Specific Inhibitors The diagram elucidating detailed interactions of ATP with the enzyme is presented in Figure 3a,b. Six out of nine invariant residues of the catalytic core of protein kinase are involved in ATP binding and catalysis. The key residues that hold the β and γ phosphates in position are the phosphate anchor, the metal sites, and Lys168. The amides of the residues—Phe54, Gly55, and Ser53—are essential for the position of the γ and β phosphates. The metal site coordinated by invariant Asp184 is also sequestered by the β and γ phosphates and the metal

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Figure 3 (a) Ternary complex of MnATP with the inhibitor peptide PKI (5–24). Glu121 and Val123 of the conserved linker region of the protein kinase catalytic core form the bidentate hydrogen bond with the 6-amino group and N1 nitrogen of the purine base. Thr183, non-conserved, forms a hydrogen bond with the N7 position of purine. 2'-OH of ribose interacts with the side chain of Glu127 of helix D and P-3 Arg of the specific inhibitor while the 3'-OH interacts with Glu170 of the catalytic loop. (b) The local environment of serine nucleophile at P site (left site) and local environment of phosphorylated P site serine (right side). The side chain of the catalytic base is at hydrogen bond distance from-OH of the Ser nucleophile which defines the conserved substrate binding P site.

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site coordinated by invariant Asn171 is also sequestered by the α and γ phosphates. The residue Lys168 is at hydrogen-bond distance from the γ phosphate. The postulated in-line mechanism of phosphotransfer in cAPK [23] can be examined through an analysis of the MnATP and PKI (5–24), Ser substrate peptide and ADP complexes [24]. A comparison between cAPK and IRK structures indicates that Ser versus Tyr specificity is obtained by displacement of the substrate binding site in such a way that the hydroxyl of nucleophiles of both Ser and Tyr fall in the same point in the active site facing corresponding catalytic bases Asp1132 and Asp166 for IRK and cAPK respectively. The purine base of ATP is anchored to the enzyme by three hydrogen bonds, two of them involve the 6amino group and N1 nitrogen, which interact with the backbone atoms of Glu121 and Val123. the 6amino group and N1 nitrogen of the purine base form the hydrogen bonds also in the structure of CK-1 and in the structure of phosphorylase kinase. Yet, in the structure of inactive CDK-2, the purine base forms only one hydrogen bond via its N6 position. While N7 nitrogen interacts directly in cAPK with the side chain of Thr183 in CK-1, this nitrogen interacts directly with Glu55 and Tyr59 via two hydrogen-bonded water molecules and in phosposhorylase kinase via one water molecule. Hence, the N7 nitrogen and its interaction with the enzyme is a region for potential modification of ATP competitive inhibitors. Ribose is held by both enzyme (Glu127 and Glu170) and inhibitor (P-3 Arg). While the side chain of Glu127 interacts with 2'-OH of ribose in cAPK, in CK-1 the 2'-OH interacts via two water molecules with Ser91 and Asp94. In ERK-2, 2'-OH interacts with Asp109. This region has been utilized to design ATP-based specific inhibitors by modifications of the ATP-competitive nonspecific inhibitor staurosporine [25], see Figure 4. In the model cAPK with bound staurosporine inhibitor, the lactam amide group of the inhibitor functions as a bidentate hydrogen bond donor-acceptor

Figure 4 Chemical structures of staurosporine inhibitor, left, and CGP52411.

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Figure 5 (a) Staurosporine molecule docked in the ATP binding site of PKA. hydrogen bonds anchoring staurosporine molecule in the active site of PKA consists of the 6-amino group and N1 nitrogen and carbonyl of Glu121 and the amide hydrogen atom of Val123. This bidentate hydrogen bond formation has been observed in all complexes of protein kinases and ATP solved so far. (b) Inhibitor of CGP 52411 and ATP docked on the active site of PKA. Residues of Glu127 and Glu170 are also shown and these are not conserved in the EGFR kinase.

(Figure 5a). This key observation is supported by chemical data of lactam amide derivatives, which provide a plausible model of staurosporine inhibition. This is in the protonated boat-type conformation found to fit in the ATP binding cleft with minimal steric hindrance. In this model, the 4-amino group forms hydrogen bonds with the backbone carbonyl of Glu170 and the carboxylate group of Glu127 of cAPK. A model of the EGFR kinase shows that Glu127 and 170 are replaced by Cys and Arg, respectively (Figure 5b). Replacement of Glu127 by Cys is critical according to the model and explains the several-fold decrease of potency of staurosporine inhibitor toward the EGFR kinase (IC50=630 nM

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Figure 5 (Continued)

versus IC50=15 nM for cAPK). Yet enhancement of specificity utilizing the inhibitor CGP52411 (Figure 4), whose selectivity originates in the occupancy by one of the anilino moieties of the inhibitor in the region of the enzyme cleft that normally binds the ribose ring of ATP, is considerable. The inhibitor CGP52411 inhibits the EGFR tyrosine kinase with an IC50 value of 300 nM while it is less active by at least two orders of magnitude on a panel of protein kinases including cAPK, phosphorylase kinase, casein kinase, protein kinase C (most isoforms), and v-abl, c-lyn, c-fgr tyrosine kinases. Hence, the three-dimensional model of EGFR tyrosine kinase rationalizes the specificity of the CGP52411 inhibitor. This model suggests that analysis of the putative regions of the ribose binding of ATP in other kinases through template modeling would provide the required chemical modification of pharmacophores to enhance their selectivity. In modeling, the use of the bidendate hydrogen bonding provided by the linker region (Figure 3a) of the conserved

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protein kinase core is essential. Yet to predict the region of the structure of the inhibitor that will form these strong Watson-Crick hydrogen bonds remains difficult. The use of the 6-amino group and N1 nitrogen of the purine base to model the pharmacophore, as seen with staurosporine, is not possible in two other adenine-type inhibitors. Both isopentenyl adenine, a nonspecific inhibitor of protein kinases, and olomoucin, a more specific inhibitor of Ser/Thr protein kinases, are modified only at the 6-amino group position. Thus, bidentate hydrogen-bond formation as seen in the ATP purine base is not possible. Furthermore, there are inhibitors that do not contain the chemical structure of adenine, for example des-chloro-flavopyridol, a potent inhibitor of cdc-2 cell cycle kinase. In crystallographic analysis of the binding of three inhibitors, olomoucin (OLO), isopentenyl adenine (ISO) and des-chloro-flavopyridol (DFP) to inactive CDK-2 cell cycle protein kinase, Kim and coworkers [26] have provided additional insight into the binding of adenine-and nonadenine-based inhibitors. Inhibitors with purine rings (OLO and ISO) bind in relatively the same area of the binding cleft as the adenine ring of ATP. Relative orientation of each purine ring with respect to the protein is different for all three ligands. This is most likely due to the fact that the 6-amino group of adenine in ATP is replaced by an isopentenylamino group in ISO and by the bulky benzylamino group in OLO. In the case of the third inhibitor, which is not an adenine derivative, the benzopyran ring occupies approximately the same region as the purine ring of ATP. The two ring systems overlap in the same plane but benzopyran is rotated about 60 degrees relative to the adenine of ATP. In this orientation, two strong bidendate hydrogen bond are formed with the oxygens in the 4th and 5th positions of the inhibitor. Furthermore, these bonds are the same ones formed by the 6-amino group and N1 nitrogen of the adenine ring. Crystallographic analysis has shown that both the substrate and ATP-binding clefts are structurally conserved yet differ in the surface charges between individual protein kinases. The structural template of the protein kinase family as discovered in the structure solution of cAPK predicts these differences. Template modeling provides a rational basis for the design of specific inhibitors for protein kinases based on the ATP binding site [25]. This is the first significant step in the design of specific inhibitors targeted at the ATP site. Recent work has now shown the conformational diversity of inhibitors binding in the interdomain ATPbinding cleft [26]. Although the residues of the protein kinase catalytic core that form the bidentate donor-acceptor bond with inhibitors are identical throughout different structures, the residues of the inhibitors vary greatly. All inhibitors use this common bidentate bond yet the specificity lies in several other bonds formed between the inhibitor and specific regions of the individual protein kinases. Furthermore, it is difficult to model

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the bidentate bond-forming residues of the inhibitor and it is through protein crystallography that these are determined. Template modeling can then be used to determine the specific surface charges of the modeled kinase that can be exploited by the inhibitor for its specificity. Modeling and focused combinatorial chemistry are the tools to achieve the goal of inhibitor specificity. Acknowledgments This work was supported by CTR grant 4237. We thank Drs. Furet, P. Traxler, and N. Lydon of Ciba for their drawings presented in Figure 5. We also thank Dr. Nikola P. Pavletich for the coordinates of the CDK2-CYCLIN complex used in Figure 2. References 1. De Bondt HL, Rosenblatt J, Jancarik J, Jones HD, Morgan DO, Kim SH. Crystal structure of cyclindependent kinase2. Nature 1993; 363:595–602. 2. Hu S-H, Parker MW, Lei JY, Wilce MCJ, Benian GM, Kemp BE. Insights into autoregulation from the crystal structure of twitchin kinase. Nature 1994; 369:581–584. 3. Hubbard SR, Wei L, Ellis L, Hendrickson WA. Crystal structure of the tyrosine kinase domain of the human insulin receptor. Nature 1994; 372:746–754. 4. Zhang F, Strand A, Robbins D, Cobb MH, Goldsmith EJ. Atomic structure of the MAP kinase ERK2 at 2.3 Å resolution. Nature 1994; 367:704–710. 5. Owen DJ, Noble MEM, Garman EF, Papageorgiou AC, Johnson LN. Two structures of the catalytic domain of phosphorylase kinase; an active protein kinase complexed with substrate analogue and product. Structure 1995; 3:467–482. 6. Xu R-M, Carmel G, Sweet RM, Kuret J, Cheng X. Crystal structure of casein kinase-1, a phosphatedirected protein kinase. The EMBO Journal 1995; 14:1015–1023. 7. Jeffrey PD, Russo AA, Polyak K, Gibbs E, Hurwitz J, Massague J, Pavletich NP. Mechanism of CDK activation revealed by the structure of a cyclinA-CDK2 complex. Nature 1995; 376:313–320. 8. Goldberg J, Nairn AC, Kuryian J. Structural basis for the autoinhibition of calcium/calmodulindependent protein kinase I. Cell 1996; 84:875–887. 9. Knighton DR, Zheng J-H, Ten Eyck LF, Xuong N-H, Taylor SS, Sowadski JM. Structure of a peptide inhibitor bound to the catalytic subunit of cyclic adenosine monophosphate-dependent protein kinase. Science 1991; 253:414–420.

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10. Knighton DR, Zheng J-H, Ten Eyck LF, Ashford VA, Xuong N-H, Taylor SS, Sowadski JM. Crystal structure of the catalytic subunit of cyclic adenosine monophosphate dependent protein kinase. Science 1991; 253:407–414. 11. Zheng J-H, Trafny EA, Kninghton DR, Xuong N-H, Taylor SS, Ten Eyck LF, Sowadski JM. 2.2Å refined crystal structure of the catalytic subunit of cAMP-dependent protein kinase complexed with MnATP and a peptide inhibitor. Acta Cryst 1993; D49:362–365.

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12. Zheng J-H, Knighton DR, Ten Eyck LF, Karlsson R, Xuong N-H, Taylor SS, Sowadski JM. Crystal structure of the catalytic subunit of cAMP-dependent protein kinase complexed with MgATP and peptide inhibitor. Biochemistry 1993; 32:2154–2161. 13. Sowadski JM, Ellis C, Madhusudan. Detergent binding to unmyristylated protein kinase A—structural implications for the role of myristate. Journal of Bioenergetics and Biomembranes 1996; 28:7–12. 14. Knighton DR, Bell S, Xuong N-H, Ten Eyck LF, Taylor SS, Sowadski JM. 2.0Å refined crystal structure of catalytic subunit of cAMP-dependent protein kinase complexed with a peptide inhibitor and detergent. Acta Cryst 1993; D49:357–361. 15. Karlsson RF, Zheng J-H, Xuong N-H, Taylor SS, Sowadski JM. Crystal structure of the mammalian catalytic subunit of cAMP-dependent protein kinase and an inhibitor peptide displays an open conformation. Acta Cryst 1993; D49:381–388. 16. Zheng J-H, Knighton DR, Xuong N-H, Taylor SS, Sowadski JM, Ten Eyck LF. Crystal structures of the myristylated catalytic subunit of cAMP-dependent protein kinase reveal open and closed conformations. Proteins Science 1993; 2:1559–1573. 17. Taylor SS, Radzio-Andzelm E. Three protein kinase structures define a common motif. Structure 1994; 2:345–355. 18. Knighton DR, Pearson RB, Sowadski JM, Means AR, Ten Eyck LF, Taylor SS, Kemp BE. Structural basis of the intrasteric regulation of myosin light chain kinase. Science 1992; 258:130–135. 19. Senften M, Schenker G, Sowadski JM, Ballmer-Hofer K. Catalytic activity and transformation potential of v-Src require arginine 385 in the substrate binding pocket. Oncogene 1995; 10:199–203. 20. Karlsson RF, Madhusudan Taylor SS, Sowadski JM. Intermolecular contacts in various crystal forms related to the open and closed conformational states of the catalytic subunit of cAMP-dependent protein kinase. Acta Cryst 1994; D50:657–662. 21. Marcote MJ, Knighton DR, Basi G, Sowadski JM, Brambilla P, Draetta G, Taylor SS. A threedimensional model of the cdc2 protein kinase: identification of cyclin and suc1 binding regions. Molecular and Cellular Biology 1993; 13:5122–5133. 22. Songyang, Z et al., Catalytic specificity of protein tyrosine kinase is critical for selective signalling. Nature 1991; 373:536–539. 23. Ho M-F, Bramson HN, Hansen DE, Knowles JR, Kaiser ET. Stereochemical course of the phospho group transfer catalyzed by cAMP-dependent protein kinase. J Am Chem Soc 1988; 110:2680–2681.

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24. Madhusudan Xuong N-H, Ten Eyck LF, Taylor SS, Sowadski JM. cAMP-dependent protein kinase: Crystallographic insights into substrate recognition and phosphotransfer. Protein Science 1994; 3:176–187. 25. Furet P, Caravattti G, Priestle J, Sowadski J, Trinks U, Traxler P. Modeling study of protein kinase inhibitors: Binding mode of staurosporine-origin of the selectivity of CGP 52 411. J Comp Aid Mol Design 1995; 9:465–472. 26. Azevedo WF Jr, Mueller-Diechmann H-J, Schulze-Gahmen U, Worland PJ, Sausville E, Kim S-H. Proc Natl Acad Sci 1996; In press. 27. Kraulis PJ, MOLSCRIPT—A program to produce both detailed and schematic plots of protein structures. Journal of Applied Crystallography 1991; 24:946–950.

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9 Structural Studies of Aldose Reductase Inhibition David K. Wilson and Florante A. Quiocho Baylor College of Medicine, Houston, Texas J. Mark Petrash Washington University School of Medicine, St. Louis, Missouri I. Introduction Aldose reductase (ALR2; EC 1.1.1.21) is an ~36 kDa enzyme that catalyzes the reduction of a wide range of carbonyl-containing compounds to their corresponding alcohols. It is a member of an extensive aldo-keto oxidoreductase enzyme family, a collection of structurally similar proteins expressed in both animals and plants. Most members of the enzyme family possess similarities in molecular mass, pH optimum, coenzyme dependence, and demonstrate overlapping specificity for many substrates and inhibitors. While no essential physiological function has been established for ALR2, extensive experimental evidence suggests that it plays an important role in the development of diabetic complications affecting the visual, nervous, and renal systems [1]. The linkage between ALR2 and pathogenesis of diabetic complications lies in the polyol pathway of glucose metabolism (Figure 1). In hyperglycemic tissues such as in diabetes mellitus, the capacity of hexokinase to shunt glucose to glycolysis and other major pathways of glucose metabolism is exceeded. Consequently, enhanced flux of glucose through the polyol pathway occurs. The enzyme ALR2 catalyzes the first step in this pathway, producing sorbitol, an active osmolyte. The polyol pathway is completed by the NAD+-dependent oxidation of sorbitol to fructose, mediated by sorbitol dehydrogenase. Extensive evidence exists to suggest a linkage between the pathogenesis of diabetic complications and enhanced glucose metabolism via the polyol pathway. The polyol pathway functions in all tissues susceptible to clinically

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Figure 1 Schematic of the polyol pathway showing the NADPH-dependent reduction of open chain D-glucose to sorbitol, which is catalyzed by ALR2. This step is followed by the NAD+-dependent oxidation of sorbitol by sorbitol dehydrogenase to yield D-fructose.

significant diabetic complications. Transgenic animals overexpressing ALR2 in target tissues of diabetic complications are more prone to development of experimentally induced diabetic complications [2,3]. The most extensive body of evidence linking ALR2 to the pathogenesis of diabetic complications comes from numerous successes in the treatment of experimental animals with a variety of ALR2 inhibitors (ARI) [4]. Many of these studies demonstrated that ARIs substantially delay or in some cases prevent the onset of complications. Clinical trials of ARIs have yielded encouraging results in alleviating painful symptoms of diabetic complications. However, unacceptable side effects related to toxicity or inadequate pharmocokinetic profiles have rendered most of the drug candidates undesirable. Nevertheless, several ARIs are commercially available in some countries and more appear to be in the pipeline. The therapeutic rationale for treatment of human diabetics with ARIs to delay or prevent onset of diabetic complications is compelling. Animal models with experimentally induced hyperglycemia develop complications that are morphologically and functionally similar to that seen in the human diabetic patient. Many structurally

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diverse ARIs have been shown to substantially delay or completely prevent the onset of such complications in experimental animal models. While some studies indicate that ALR2 may play a functional role in osmotic homeostasis in the kidney, evidence from animal studies suggests that it is metabolically dispensible. Long-term complications exact a terrible toll of morbidity and mortality on patients with diabetes mellitus. For example, patients with diabetes have about a 25-fold increased risk for becoming blind over that of the general population. Diabetic retinopathy is one of the most common causes of visual loss and accounts for about 12% of new cases of blindness each year in the United States alone [5]. II. Drug Design Prior to Structural Data Many inhibitors have been developed over the past two decades without the advantage of a structural understanding of the enzyme [4,6,7]. Significant improvement has been made since the discovery of the first such orally active compound to show in vivo activity, alrestatin (Figure 2), which had an IC50 in the low micromolar range [8]. Many high-affinity inhibitors with IC50s in the low nanomolar range are now under study. Recent drug-design efforts have yielded compounds usually with one of two chemical motifs: carboxylates or spirohydantoins. A number of these compounds such as tolrestat [9], ponalrestat [10], epalrestat [11], sorbinil [12], and zopolrestat [13] have progressed to the point of clinical trials. Unfortunately, clinical ineffectiveness and/or unacceptable side effects have limited the usefulness of most of those that had been shown to be effective in vitro. The latter problems may be associated with a lack of specificity since many aldose reductase inhibitors inhibit both ALR2 and aldehyde reductase [14]. For this reason, ALR2 as well as the other members of the aldo-keto reductase family have been the subject of crystallographic studies with the hope of determining a structural basis for inhibitor specificity and ultimately to provide a basis for enhancing binding affinity. III. Structural Studies of Aldose Reductase The first crystal structures available for ALR2 were those of the porcine form complexed with the NADPH analog 2'-monophosphoadenosine-5'-diphosphoribose [15] and the human enzyme complexed with the NADPH cofactor [16]. Further studies have been conducted on mutants of the human enzyme [17] and ternary complexes of the human enzyme with an inhibitor [18]. All of these

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Figure 2 A number of ALR2 inhibitors that have entered clinical trails.

structures show the protein to fold into a (β/α)8 barrel (Figure 3). This fold has emerged as the most common enzyme motif [19] although most of the proteins adopting this structure share no sequence homology. The ALR2 enzyme is, however, the first NAD(P)H binding protein to adopt this fold. It contains an extra β hairpin preceding the first β strand, which caps the N-terminal end of the barrel. It also has two helices that are not part of the regular barrel. One precedes α7 and the other follows α8. A. Cofactor Binding The NADPH cofactor is bound in an extended conformation across the C-terminal end of the β barrel. The catalytically active nicotinamide moiety is located

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Figure 3 Cα trace of the ALR2 holoenzyme looking down the (β/α)8 barrel. The NADPH cofactor is seen bound across the carboxy-terminal end of the β-barrel with the active nicotinamide moiety in the center. Figure produced using the MOLSCRIPT program [48].

at the center of the barrel while the adenosine extends away to bind between α7 and α8. A belt composed of residues 213 to 227 folds over the pyrophosphate of the NADPH to sequester a large part of the cofactor from the solvent. It is fastened to the other side of the NADPH binding site via Asp216 on the loop that forms bifurcated salt links with Lys21 and Lys262. The dominant interactions holding the coenzyme in place are directional hydrogen bonds and salt links from positively charged side chains to the phosphates. The interaction between the 2' phosphate on the NADPH and the side chains from Lys262 and Arg268 account for the enzyme's preference for NADPH over NADH. Earlier biochemical studies had shown that it is the 4-pro-R hydride that is transferred from the nicotinamide to the substrate [20]. This is ensured by a hydrogen-bonding network using side chains from Ser159 and Asn160 and the main chain of Gln183 to orient the amide. It is also determined by the stacking interactions with Tyr209, which is adjacent to the 4-pro-S side of the nicotinamide. B. Mechanism The catalytic site was unambiguously identified using the location of the nicotinamide moiety of the NADPH cofactor in the holoenzyme structure. The region surrounding the catalytic site is a 12-Å deep groove that measures

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approximately 7 by 13 Å and is lined primarily with hydrophobic side chains. This is entirely consistent with earlier experiments showing that the enzyme has a marked preference for lipophilic substrates versus polar substrates such as sugars [21]. The catalytic site also suggested a model for the enzyme's chemical mechanism, which is substantially similar to most other NAD(P)H-dependent oxido reductases. Upon binding of the reduced cofactor, the enzyme is able to form a ternary complex with the substrate. The pro-R hydride from the C-4 of the nicotinamide is transferred to the carbonyl carbon of the substrate, which in turn causes the carbonyl oxygen to abstract a proton from a general acid, which is presumably located on the protein, to form the alcoholic product. Three proton-donating side chains are located within 6 Å of the C-4 atom in the NADPH cofactor that could potentially fulfill this role: Tyr48, His110, and Cys298. Since it is not conserved in other members of the aldo-keto reductase family that exhibit enzymatic activity (Figure 8) Cys298 was unlikely as a candidate proton donor. The histidine is surrounded by several hydrophobic residues including Val47, Trp79, and Trp111, which would serve to lower the pKa of the side chain, making it less effective as a proton donor at physiological pHs. The tryrosine, which ordinarily has a pKa of approximately 11 engages in an interaction with the charged ammonium group of Lys77, which in turn charge-pairs with Asp43. This network serves to depress the pKa of the phenolic oxygen, increasing the exchangability of the proton. Subsequent activity studies involving site-directed mutants support this model [17,22]. The Tyr48 rarrow.gif Phe mutation shows a complete lack of activity while the Asp43 rarrow.gif Asn, Lys77 rarrow.gif Met, His110 rarrow.gif Asn, and Cys298 rarrow.gif Ser showed losses in catalytic efficiency of approximately 100-, 1000-, 106-, and 10-fold respectively when compared with the wildtype enzyme. These results correlate well with the functions predicted for each residue with the exception of the histidine. The structure of the ALR2 holoenzyme showed that the catalytic site was situated atop the nicotinamide moiety of the NADPH cofactor. The substrate binding site, which would determine the enzyme's specificity and also presumably bind inhibitors, appeared to be composed of a deep cleft (Figures 3 and 4). It extended away from the catalytic site towards the loop composed of residues between β4 and α4 and the last 20 residues of the carboxy-terminal meander. This hypothesis was supported by the appearance of poorly resolved density that occupied this region, which suggested the presence of an endogenously bound substrate or inhibitor in the structure of the holoenzyme [16]. Subsequent studies indicate that this electron density may be a citrate molecule, one of the components included in the crystallization mixture. Activity studies indicate that citrate is indeed one of the many inhibitors of the enzyme with a Ki in the millimolar range [23].

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Figure 4 Surface representation of the ALR2 holoenzyme in an orientation similar to Figure 3. The nicotinamide moiety that defines the active site of the enzyme is seen in the center. The groove that extends down from it is highly hydrophobic and was initially assumed to be the inhibitor binding site. Figure prepared using the GRASP program [49].

IV. Aldose Reductase Complexed with Inhibitor While a large number of high-potency inhibitors for ALR2 have been developed [4], a structural understanding of the exact molecular features that foster this affinity have been only vaguely understood. Several general chemical motifs such as hydrophobic ring systems, a spirohydrantoin group or carboxylate group are seen repeatedly when examining a list of known inhibitors (Figure 2) but little was known about the specific role for each in inhibitor binding. The structure of the ALR2/NADPH/zopolrestat ternary complex [18] has provided some answers about the mode of binding of zopolrestat (Figure 2), a high-affinity, carboxylate-containing compound developed by Pfizer, Inc. [13]. While the overall structure was preserved, the inhibitor binding induced a conformational change of the enzyme. This change, which involved the movement of several loops in the active site of the molecule, was large enough to cause a change in crystal packing relative to the holoenzyme. As a consequence of the shifting of the active-site loops, a cavity is created inside the protein in which the benzothiazole ring is seated and the groove that was implicated in substrate and inhibitor binding by the holoenzyme structure vanishes. This illustrates the unpredictability of conformational changes within a protein in response to substrate or inhibitor binding. It also implies that modeling compounds

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in the active site of an enzyme or drug-design algorithms such as inhibitor docking may be inappropriate in some cases if they do not adequately model considerable plasticity in the binding site. A closer look at the binding of zopolrestat shows that it is dominated by extensive hydrophobic contacts between the protein and the inhibitor. These include side chains from Trp20, Tyr48, Trp79, Trp111, Phe115, Phe122, Trp219, Ala299, Leu300, Tyr309, and Pro310 (Figure 7). This is not surprising given the apolar nature of the enzyme's active site as determined by the holoenzyme structure. What is surprising is that the inhibitor created the part of its own binding site that the benzothiazole rings occupy by “burrowing” into the hydrophobic core of the protein to carve out a region with very good steric complementarity to this moiety. It does this rather than binding in the solvent-exposed hydrophobic binding groove that is seen in the holoenzyme structure. The remaining interactions involving hydrogen bonds and salt links also appear to be very important in inhibitor binding. With the exception of one of the fluorine atoms, all atoms that are able to engage in hydrogen bonding do so. The carboxylate, which is seen in so many aldose reductase inhibitors, is saltlinked to His110, which is located very near the catalytic site (Figure 7). Presumably, the carboxylate in the other inhibitors plays the same role and could be used as an anchor when modeling these into the active site. Inhibition studies involving ALR2 have indicated noncompetitive inhibition for virtually all compounds examined to date when the forward (reduction) reaction is monitored. This mode of inhibition is often interpreted as meaning that the inhibitor binds to a site on the enzyme that is independent of the catalytic site. Kinetic and competition studies have both led to this conclusion in the case of ALR2 [24,25]. The crystal structure of the enzyme complexed with both the NADPH cofactor and zopolrestat, however, clearly shows the inhibitor occupying the region directly above the nicotinamide of the NADPH and, therefore, the active site (Figures 5, 6, and 7). Most previous inhibition studies reported noncompetitive and/or uncompetitive inhibition patterns when aldose reductase inhibitors were examined in the forward direction, i.e. inhibition of NADPH-dependent aldehyde reduction. With the finding that the overall rate-limiting step in the direction of aldehyde reduction is at the level of structural isomerization following alcohol product release [26,27], it is not surprising that lack of competitive inhibition would be observed in such standard double reciprocal plots. To further complicate matters, many aldose reductase inhibitors were not recognized in previous studies as tight-binding inhibitors and were inappropriately evaluated using Michaelis-Menten kinetics. Thus, noncompetitive or uncompetitive inhibition patterns were previously reported for inhibitors that were subsequently shown to bind directly at the active site. Recent structure-function and kinetic studies have revealed important details concerning the structural basis for the catalytic

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Figure 5 Schematic representation of the ALR2/NADPH/zopolrestat ternary complex. The NADPH is bound across the enzyme from the center to the right while the zopolrestat binds atop the nicotinamide and extends to the lower left. Conformational changes are seen in the C-terminal loop below the zopolrestat in the picture and the loop to the right of the inhibitor between β4 and α4.

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Figure 6 A surface representation of the ternary complex as seen in Figure 5. Note that the inhbitor creates part of its binding site by “burrowing” into the protein rather than binding entirely in the groove seen Figure 4.

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Figure 7 Stereo of zopolrestat binding to the active site of ALR2. The salt link made by the carboxylate of the inhibitor and hydrogen bonds are depicted with dashed lines. The remainder of the interactions are apolar with the residues shown.

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and inhibition mechanisms and provided a clarification for the mechanism by which inhibitors of both the carboxylate [28] and spirohydantoin [29] classes bind at the active site of ALR2. V. Aldo-Keto Reductase Family A. Effects of Sequence on Drug Design The problem of structure-based drug design for ALR2 and the drug-design effort in general is compounded by the fact that this enzymes is a member of a large family of aldo-keto reductases with overlapping substrate specificity. In humans at least three such enzymes have been found: ALR2 [30], aldehyde reductase (ALR1) [30], and chlordecone reductase [31]. Other members of the family that have been isolated in other species include rat 3α-hydroxysteroid dehydrogenase [32], murine fibroblast growth factor induced protein [33], bovine prostagladin F synthase [34], murine vas deferens protein [35], frog ρ-crystallin [36], the P100/11E gene product in Leishmania major [37], and Corynebactium diketogluconate reductase [38]. This large number suggests that there may be more such enzymes to be found in humans. The similarities between the proteins with respect to both the sequence and substrate specificity implies that the nature of the substrate binding sites are similar across the family. This has indeed been the case in all the structures determined from this family to date (see below). While detailed binding studies of various inhibitors with all the different enzymes have not been conducted, it is likely that drugs intended for ALR2 are likely to “cross react” with many of the other enzymes within the family. One such case that has recently been studied both crystallographically and biochemically is the murine FR-1 protein described below. The binding sites of all of these enzymes are characterized by their large size and hydrophobicity suggesting that ideal substrates may be steroids or molecules of a similar size and nature. Sequence comparisons of all the proteins, including those whose structure has not yet been determined, show that there is a large amount of similarity involving residues implicated in substrate binding (see Figure 8). One region that diverges somewhat is the 15-amino acid segment at the carboxy terminus of the protein. This segment is likely to be responsible for what little differences in substrate specificity exhibited by the enzymes. It is the same segment that is seen adopting a different conformation upon zopolrestat binding to ALR2. It may then be possible that it is not only the chemical nature of this loop that—in making positive and negative interactions with the substrate/inhibitor—modulates specificity, but also the flexibility conferred by the amino acid sequence. Such a difference is seen when contrasting the structures of ALR2 and FR-1 bound to zopolrestat.

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Figure 8 Sequence alignment of several members of the aldo-keto reductase family. Abbreviations used are HALR2, human aldose reductase (30); HALR1, human aldehyde reductase [30]; 3α-HSD, 3α-hydroxysteroid dehydrogenase [32]; FR-1, murine FR-1 [33]; BPGFS, bovine prostaglandin F synthase [34]; CCDR12, human chlordecone reductase [31]; CDGR, Corynebacterium diketogluconate reductase [38]; MVDP, murine vas deferens protein [35], JFRC, Japanese frog ρ crystallin [36].

B. Structures Structures of several other members of the aldo-keto reductase family have also been determined. These include aldehyde reductase [39,40], FR-1 [41] and 3α-hydroxysteroid dehydrogenase [42]. Since each of these proteins retain a large amount of sequence identity and homology with human ALR2, it is not surprising to note that the overall tertiary structures are very similar. Root-meansquare Cα deviations between the human ALR2 holoenzyme structure and the rest of the family are in the range of 1–2Å

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Closer examination of the active site shows that the residues involved in catalysis (most notably the residues analogous to Asp43, and Tyr48, and Lys77 in ALR2) are structurally conserved among each of these proteins (Figure 8), suggesting that the mechanism is also conserved throughout the family. Many of the residues found in the binding site (defined as those making contact with the zopolrestat in the ternary complex) are also largely conserved with the exception of a number of residues in the carboxy terminus of the protein. This is indeed where the most structure variation appears to be concentrated among the proteins. These residues compose a loop that is the same loop that shifts upon binding of zopolrestat in ALR2. Inhibitors with improved specificity will very likely take advantage of the subtle structural differences that are introduced by the variation in sequences in this area. VI. Future Design of Aldose Reductase Inhibitors The availability of structural data for ALR2 in its holoenzyme and different ternary forms is likely to lead to improvements in the affinity of future generations of inhibitors. As the architecture and plasticity of the binding site are better understood, increasingly potent inhibitors may be designed to occupy it. Although these structures provide a positive target for drug design, there are a number of negative targets. Increased in vivo potency is likely to be derived from the specific inhibition of ALR2 that would entail the avoidance of other members of the aldo-keto reductase family. Determination of the structures of other members of the family may increase the specificity of compounds by providing structures of targets to avoid. While the incorporation of the negative targets in the drug-design process relies on the determination of other structures and is likely to be complicated, conventional computational techniques may be applied to the problem of the positive target. Two such methods that may hold promise are docking [43] and computational thermodynamic perturbation [44]. A. Inhibitor Docking to the Enzyme Our initial efforts to exploit the ALR2 holoenzyme structure for drug design utilized the program DOCK [43]. This program is capable of finding depressions on the surface of the enzyme that could serve as binding sites for substrates or inhibitors. Once the correct area is defined, the program rotates structures of candidate compounds within this space and scores each compound based upon its steric complementarity with the binding site. However, the program does not include the potential polar interactions between the inhibitor and protein when scoring. The search was further constrained by the inability to include conformational variations both in the test compounds as well as the protein, due to computational limitations.

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This method was used to screen a significant portion of the contents of the Cambridge Structural Database [45] against the ALR2 holoenzyme binding site (D.K. Wilson, J. M. Petrash, and F. A. Quiocho, unpublished data). Among the approximately 30 highest scoring compounds were several aromatic aldoximes that had inhibition constants in the micromolar range. These were similar to aldoximes such as benzaldoxime, which has been previously observed to have similar inhibition constants [46]. A disappointing result was that this search did not “rediscover” any of the known high-affinity ALR2 inhibitors that were contained in the search library. Before the determination of the ternary complex of the enzyme with zopolrestat, this was interpreted as meaning that these compounds bound to the enzyme in a conformation somewhat different than the one adopted in the crystal structure used for the search. The structure of the ternary complex showed this to be a wrong assumption; it was the protein that changed conformation upon inhibitor binding, creating a pocket that did not exist in the holoenzyme structure. When bound to the protein, zopolrestat is actually quite similar in conformation to its small molecule x-ray structure. It is therefore very possible that different ALR2 inhibitors and substrates may cause the enzyme to flex in different ways, creating binding sites that may be different in size and chemical nature. B. Computational Thermodynamic Perturbation Computational thermodynamic perturbation is a powerful, albeit computationally expensive, group of techniques that are designed to estimate relative binding affinities of two closely related drugs, given the structure of at least one of them complexed with the target protein [44]. This approach has the potential to assay candidate compounds in the computer for improvements in inhibitor binding, thereby removing the necessity to sythesize and assay these compounds in the lab. For a number of reasons, ALR2 promises to be a good system for the application of this technique and the experimental verification of the results. The structure is very well determined in complex with zopolrestat, a high-affinity inhibitor. A number of zopolrestat derivatives with various functional groups decorating the compound have been sythesized and partially characterized with respect to ALR2 inhibition [13,47]. These compounds could serve as a sort of basis set of controls for the theoretical calculations. If parameters used in these calculations can be selected such that the computationally derived binding energies agree even qualitatively with the experimentally determined binding energies, serious consideration should be given to new compounds that are predicted to bind with enhanced affinity. Since ALR2 is crystallizable with zopolrestat bound, there is every reason to believe that crystals of the enzyme complexed with similar compounds will be obtainable. Such structures could provide the basis for further rounds of drug improvement.

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VII. Conclusions The observation of a protein undergoing a conformational change when binding to an inhibitor, as seen with ALR2 and zopolrestat, illustrates a common problem associated with structure-based drug design. It is tempting to view proteins as static structures since their crystal structures are static. Attempts to design drugs to fit the apparent active site of an enzyme may fail when the plasticity of the protein is not taken into account. While the disorder associated with amino acid side chains can be modeled with a moderate computational effort, larger conformational changes—such as the loop movement seen in ALR2—are virtually impossible to predict. Until this becomes possible, x-ray crystal structures of complexes will continue to be indispensible. Finally, it can be easy to forget that a compound's affinity for the protein is not the only consideration when designing inhibitors of enzymes from a structural point of view. The structures of aldose reductase and the FR-1 protein complexed with the drug zopolrestat, a compound with a very high affinity for ALR2, can serve as a reminder of how specificity can also be a very important factor. This is particularly true when a protein is a member of a family of proteins that share sequence homology and are apt to have overlapping specificities. Structure may then play a key role in the determination of features that are unique to the target protein and therefore prime considerations when designing inhibitors. Acknowledgments We thank T. Reynolds who assisted with the production of the figures. This work was supported by a grant from Research to Prevent Blindness, Inc. and grants EY05856, EY02687, and DK20579 to J. Mark Petrash. Florante A. Quiocho is an investigator of the Howard Hughes Medical Institute. References 1. Kinoshita JH, Nishimura C. The involvement of aldose reductase in diabetic complications. DiabetesMetebolism Rev 1988; 4:323–337.

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2. Lee AYW, Chung SK, Chung SSM. Demonstration that polyol accumulation is responsible for diabetic cataract by the use of transgenic mice expressing the aldose reductase gene in the lens. Proc Natl Acad Sci USA 1995;92:2780–2784. 3. Yamaoka T, Nishimura C, Yamashita K, Itakura M, Yamada T, Fujimoto J, Kokai Y. Acute onset of diabetic pathological changes in transgenic mice with human aldose reductase cDNA. Diabetologia 1995;38:255–261. 4. Sarges R, Oates P. Aldose reductase inhibitors: Recent developments. Prog Drug Res 1993; 40:99–161.

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5. National Advisory Eye Council (1994). In: Vision Research: A National Plan 1994–1998. National Institutes of Health Publication No. 93–3186. 6. Kador PF. The role of aldose reductase in the development of diabetic complications. Med Res Rev 1988; 8:325–352. 7. Dvornik D. Aldose Reductase Inhibition: An Approach to the Prevention of Diabetic Complications. New York: Biomedical Information Corporation 1987. 8. Dvornik D, Simard-Duquesne N, Krami M, Sestanj K, Gabbay KH, Kinoshita JN, Varma SD, Merola LO. Polyol accumulation in galatosemic and diabetic rats: control by an aldose reductase inhibitor. Science 1973; 182:1146–1148. 9. Sestanj K, Bellini F, Fung S, Abraham N, Treasurywala A, Humber L, Simard-Duquesne N, Dvornik D. N-[[5-(trifluoromethyl)-6-methoxy-1-naphthalenyl]thioxomethyl]-N-methylglycine (Tolrestat), a potent, orally active aldose reductase inhibitor. J Med Chem 1984;27:255–256. 10. Ward WHJ, Sennitt CM, Ross H, Dingle A, Timms D, Mirrlees DJ, Tuffin DP. Ponalrestat: a potent and specific inhibitor of aldose reductase. Biochem Pharmacol 1990; 39:337–346. 11. Terashima H, Hama K, Yamamoto R, Tsuboshima M, Kikkawa R, Hatanaka 1, Shigeta Y. Effects of a new aldose reductase inhibitor on various tissues in vitro. J Pharmacol Exp. Ther 1984;229:226–230. 12. Peterson MJ, Sarges R, Aldinger CD. CP-45634: a novel aldose reductase inhibitor that inhibits polyol pathway activity in diabetic and galactosemic rats. Metabolism 1979; 28(supp 1):456–461. 13. Mylari BL, Larson ER, Beyer TA, Zembrowski WJ, Aldinger CE, Dee MF, Siegel TW, Singleton DH. Novel, potent aldose reductase inhibitors: 3,4-dihydro-4-oxo-3-[[5-(trifluoromethyl)-2benzothiazolyl]methyl]-1-phthalazine-acetic acid (zopolrestat) and congeners. J Med Chem 1991; 34:108–122. 14. Srivastava SK, Petrash JM, Sadana AJ, Partridge CA. Susceptibility of aldose and aldehyde reductases to aldose reductase inhibitors. Curr Eye Res 1982; 2:407–410. 15. Rondeau JM, Tete-Favier F, Podjarny A, Reymann JM, Barth P, Biellmann JF, Moras D. Novel NADPH-binding domain revealed by the crystal structure of aldose reductase. Nature 1992; 355:469–472. 16. Wilson DK, Bohren KM, Gabbay KH, Quiocho FA. An unlikely sugar substrate site in the 1.65 Å structure of the human aldose reductase holoenzyme implicated in diabetic complications. Science 1992; 257:81–84. 17. Borhani DW, Harter TM, Petrash JM. The crystal structure of the aldose reductase-NADPH binary complex. J Biol Chem 1992; 267:24841–24847. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_244.html (1 of 2) [4/5/2004 5:09:29 PM]

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18. Wilson DK, Tarle I, Petrash JM, Quiocho FA. Refined 1.8 Å structure of human aldose reductase complexed with the potent inhibitor zopolrestat. Proc Natl Acad Sci USA 1993; 90:9847–9851. 19. Branden CI. The TIM barrel—the most frequently occurring folding motif in proteins. Curr Opin Struct Biol 1991; 1:978–983. 20. Feldman HB, Szczepanik PA, Havre P, Corrall RJM, Yu LC, Rodman HM, Rosner BA, Klein PD, Landau, BR. Stereospecificity of the hydrogen transfer catalyzd by human placental aldose reductase. Biochim Biophys Acta 1997; 480:14–20. 21. Wermuth B, Buergisser HB, Bohren KM, von Wartburg JP. Purification and characterization of human brain aldose reductase. Eur J Biochem 1982; 127:279–284.

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22. Tarle I, Borhani DW, Wilson DK, Quiocho FA, Petrash JM. Probing the active site of human aldose reductase: site directed mutagenesis of Asp-43, Lys-77 and His-110. J Biol Chem 1993; 268:25687–25693. 23. Harrison DH, Bohren KM, Ringe D, Petsko GA, Gabbay KH. An anion binding site in human aldose reductase: mechanistic implications for the binding of citrate, cacodylate, and glucose 6-phosphate. Biochemistry 1994; 33:2011–2020. 24. Kador PF, Sharpless NE. Pharmacophor requirements of the aldose reductase inhibitor site. Mol Pharmacol 1983; 24:521–531. 25. Kador PF, Goosey JD, Sharpless NE, Kolish J, Miller DD. Stereospecific inhibition of aldose reductase. Eur J Med Chem 1981; 16:293–298. 26. Kubiseski TJ, Hyndman DJ, Morjana NA, Flynn TG. Studies on pig muscle aldose reductase. Kinetic mechanism and evidence for a slow conformational change upon coenzyme binding. J Biol Chem 1992; 267:6510–6517. 27. Grimshaw CE, Shahbaz M, Putney CG. Mechanistic basis for nonlinear kinetics of aldehyde reduction catalyzed by aldose reductase. Biochemistry 1990; 29:9947–9955. 28. Grimshaw CE, Bohren KM, Lai CJ, Gabbay KH. Human aldose reductase: pK of tyrosine 48 reveals the preferred ionization state for catalysis and inhibition. Biochemistry 1995; 34:14374–14384. 29. Liu SQ, Bhatnagar A, Srivastava SK. Does sorbinil bind to the substrate binding site of aldose reductase? Biochem Pharmacol 1992; 44:2427–2429. 30. Bohren KM, Bullock B, Wermuth B, Gabbay KH. The aldo-keto reductase superfamily: cDNAs and deduced amino acid sequences of human aldehyde and aldose reductases. J Biol Chem 1989; 264:9547–9551. 31. Winters CJ, Molowa DT, Guzelian PS. Isolation and characterization of cloned cDNAs encoding human liver chlordecone reductase. Biochemistry 1990; 29:1080–1087. 32. Pawlowski JE, Huizinga M, Penning TM. Cloning and sequencing of the cDNA for rat liver 3αhydroxysteroid/dihydrodiol dehydrogenase. J Biol Chem 1991; 266:8820–8825. 33. Donohue PJ, Alberts GF, Hampton BS, Winkles JA. A delayed-early gene activated by fibroblast growth factor-1 encodes a protein related to aldose reductase. J Biol Chem 1994; 269:8604–8609. 34. Watanabe K, Fujii Y, Nakayama K, Ohkubo H, Kuramitsu S, Kagamiyama H, Nakanishi S, Hayaishi O. Structural similarity of bovine lung prostaglandin F synthase to lens ε crystallin of the European common frog. Proc Natl Acad Sci U S A 1988; 85:11–15.

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35. Pailhoux EA, Martinez A, Veyssiere GM, Jean CG. Androgen-dependent protein from mouse vas deferens: cDNA cloning and protein homology with the aldo-keto reductase superfamily. J Biol Chem 1990; 265:19932–19936. 36. Fujii Y, Watanabe K, Hayashi H, Urade Y, Kuramitsu S, Kagamiyama H, Hayashi O. Purification and characterization of p-crystallin from Japanese common bullfrog lens. J Biol Chem 1990; 265:9914–9923. 37. Samaras N, Spithill TW. The developmentally regulated P100/11E gene of Leishmania major shows homology to a superfamily of reductase genes. J Biol Chem 1989; 264:4251–4254. 38. Anderson S, Marks CM, Lazarus R, Miller J, Stafford K, Seymour J, Light D, Rastetter W, Estell D. Production of 2-keto-L-gulonate, an intermediate in L-

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ascorbate synthesis, by a genetically modified Erwinia herbicola. Science 1985; 230:144–149. 39. El-Kabbani O, Green NC, Lin G, Carson M, Narayanam SVL, Moore K, Flynn TG, DeLucas LJ. Structures of human and porcine aldehyde reductase: an enzyme implicated in diabetic complications. Acta Crystallogr D 1994; 50:859–868. 40. El-Kabbani O, Judge K, Ginell SL, Myles Daa, DeLucas LJ, Flynn TG. Structure of porcine aldehyde reductase holoenzyme. Nat Struct Biol 1995; 2:687–692. 41. Wilson DK, Nakano T, Petrash JM, Quiocho FA. 1.7 Å structure of FR-1, a fibroblast growth factorinduced member of the aldo-keto reductase family complexed with coenzyme and inhibitor. Biochemistry 1995; 34:14323–14330. 42. Hoog SS, Pawlowski JE, Alzari PM, Penning TM, Lewis M. Three-dimensional structure of rat liver 3α-hydroxysteroid/dihydrodiol dehydrogenase: a member of the aldo-keto reductase superfamily. Proc Natl Acad Sci USA 1994; 91:2517–2521. 43 Schoichet B, Bodian D, Kuntz I. Molecular docking using shape descriptors. J Comp Chem 1992; 13:380–397. 44 Straatsma TP, McCammon JA. Computational alchemy. Annu Rev Phys Chem 1992 43:407–435. 45 Allen FG, Bellar SA, Brice MD, Cartwright BA, Doubleday A, Higgs H, Hummelink T, HummelinkPeters BG, Kennard O, Motherwell WDS, Rodgeres JR, Watson DG. The Cambridge Crystallographic Data Centre: Computer based search retrieval, analysis and display of information. Acta Crystallogr B35:2331–2339. 46 Shen C, Sigman DS. New inhibitors of aldose reductase: anti-oximes of aromatic aldehydes. Arch Biochem Biophys 1991; 286:596–603. 47 Mylari BL, Beyer TA, Scott PJ, Aldinger CE, Dee MF, Siegel TW, Zembrowski WJ. Potent, orally active aldose reductase inhbitors related to zopolrestat: surrogates for benzothiazole side chain. J Med Chem 1992; 35:457–465. 48 Kraulis PJ. MOLSCRIPT: a program to produce both detailed and schematic plots of protein structures. J Appl Crystallog 1991; 24:946–950. 49 Nicholls A, Sharp KA, Honig B. Protein folding and association: insights from the interfacial and thermodynamic properties of hydrocarbons. Proteins 1991; 11:281–296.

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10 Structure-Based Design of Thrombin Inhibitors Patricia C. Weber and Michael Czarniecki Schering-Plough Research Institute, Kenilworth, New Jersey I. Roles of Thrombin in Hemostasis and the Therapeutic Utility of Thrombin Inhibitors Thrombin is a serine protease that plays critical roles in both blood clot formation and anticoagulation. In the penultimate step of the coagulation cascade, thrombin cleaves soluble fibrinogen to form insoluble fibrin. Thrombin also activates other coagulation factors including Factor XIII, the enzyme responsible for crosslinking fibrin to further stabilize the thrombus. Additional clot-promoting functions include stimulation of platelet aggregation by cleavage of the thrombin receptor. In contrast to its roles in clot formation, thrombin participates in anticoagulant functions. For example, thrombin-mediated activation of protein C, a protease involved in anticoagulation, is enhanced when thrombin is complexed with thrombomodulin, and in this complex, thrombin can neither cleave fibrinogen nor activate platelets. The interrelationship among thrombin's many roles in hemostasis is complex and presents several mechanisms for inhibition of thrombus formation. For recent reviews see References 1 through 5. Most drug discovery efforts focus on thrombin inhibition as a means to prevent the serious consequences of thrombus formation in myocardial infarction and stroke. Thrombin inhibitors may also prevent clot formation in patients prone to deep vein thrombosis or repeat heart attack. In combination with thrombus dissolution therapies, thrombin inhibitors may decrease the incidence of reocclusion due, in part, to the release of active clot-bound thrombin. In this article, recent examples of small molecule inhibitors interacting at the fibrinogen primary specificity pocket and with residues of the catalytic triad

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are given. Inhibitors designed to make more extended interactions with thrombin are also presented. II. Structure of Thrombin Thrombin consists of two polypeptides, an A chain of 36 residues and a 259-residue B chain, linked by a disulfide bond. The crystallographic structure of thrombin reveals a globular protein organized about two β barrels with the overall folding pattern of the chymotrypsin serine protease family [6,7]. The catalytic triad and nearby oxyanion hole are located roughly between the β barrels and adopt the geometric arrangement required for serine-protease-assisted, peptide bond cleavage (Figure 1). Thrombin's multifunctionality and regulation of activity are achieved by specialized subsites on the enzyme's surface (Figure 2). Fibrinogen cleavage, for example, involves interactions at the primary specificity pocket, the extended fibrinogen recognition exosite, and an additional specificity pocket. Subsite interactions differ for cleavage of other thrombin substrates including the thrombin receptor and protein C. Additional and overlapping subsites exist for thrombin effector molecules including heparin, antithrombin III, and heparin cofactor II [8,9].

Figure 1 Stereoscopic view of the crystallographic structure of thrombin complexed with N-acetyl-(D-Phe)-Pro-boroArg-OH. Helical regions are represented in the standard way and arrows indicate regions of β sheet. Solid lines show the thrombin bound conformation of N-acetyl-(D-Phe)-Pro-boroArg-OH (taken from Reference 10). Active-site residues, His57 and Ser195, are shown with a ball-and-stick representation. The authors thank Dr. C. L. Strickland for the drawing.

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Figure 2 Schematic representation of Subsite Utilization in Thrombin Complexes (after Reference 8). Fibrinogen interacts with three thrombin subsites (here thrombin is represented by a large oval and the interconnected subsites by an irregular three-armed shape). Physiological effectors of thrombin and thrombin inhibitors form distinct interactions at these subsites. Additional subsites, such as the heparin-binding site, exist on the thrombin surface and are not indicated here. The catalytic triad is represented by three circles at the vertices of a triangle.

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III. Thrombin Inhibitors Directed at the Fibrinopeptide a Binding Pocket The majority of synthetic thrombin inhibitors interact at the fibrinopeptide A binding pocket, which includes the catalytic residues Ser195 and His57, hydrogen-bonding capabilities within the oxyanion hole, peptide backbone functional groups that hydrogen bond with the peptide backbone of the substrate, and residues involved in amino acid recognition (Figure 3). Many of these binding determinants are utilized by N-acetyl-(D-Phe)-Pro-Arg-chloromethylketone (PPACK [7]) and its boronic acid analog (DUP714 [10]). The crystallographic structures of these molecules complexed with thrombin have both served as starting points for structure-based drug design and as reference structures for comparison of binding modes of other inhibitors. The use of arginine boronate esters as transition-state mimetics results in potent peptidyl thrombin inhibitors. These inhibitors, however, exhibit significant affinity for other serine proteases that have in common a specificity for substrates with basic residues at P1 (e.g. trypsin, Factor Xa, and plasmin). Earlier work demonstrated that neutral side chains of P1 boronate esters impart greater selectivity for thrombin. The boropeptide shown in Figure 4 was investigated as the prototype of neutral side chain, tripeptide thrombin inhibitors [11]. It had a Ki against thrombin of 7 nM and shows selectivity relative to other trypsin-like plasma proteases. Since these inhibitors have a neutral residue at the P1 site, Deadman and coworkers [11] sought to demonstrate the mode of binding to thrombin in the absence of a salt bridge with Asp189.

Figure 3 Schematic diagram of binding determinants within the fibrinopeptide A binding pocket of thrombin and their utilization by N-acetyl-(D-Phe)-Pro-boroArg-OH.

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Figure 4 Schematic representation of the active-site orientation of a “neutral” P1 boronic acid thrombin inhibitor.

Boron-11-NMR, a sensitive probe of the chemical environment around boronate esters, can distinguish between trigonal and tetrahedral forms of boron. The 11B-NMR spectrum of this inhibitor complexed with thrombin showed a single peak at -17 ppm that remained constant for 7 hours. The chemical shift suggests boron adopts a tetrahedral geometry on binding to thrombin and is consistent with the orientation of the inhibitor in the active site shown in Figure 4. While the 11B-NMR revealed an interaction within the catalytic site, it could not distinguish between bonding with Ser195 or His57. Kahn and coworkers [12] recently investigated the application of synthesized peptidomimetics as novel inhibitors of thrombin. Fibrinogen peptide A mimetic (FPAM, Figure 5) incorporates a bicyclic peptidomimetic within the turn region of fibrinogen peptide A. The bicyclic peptidomimetic confers conformational stability to the turn region as suggested by x-ray crystal structures of fibrinogen peptide complexes as well as complexes of BPTI with thrombin. X-ray crystallographic studies of FPAM complexed with thrombin (Figure 5) showed that the S1 subsite is occupied by the arginine guanidinium [12]. The Val group of FPAM makes extensive hydrophobic contacts within the S2 apolar binding site. The Gly at P3 interacts with thrombin via a β-sheet-type hydrogen bond with the carbonyl group of Gly216 and appears to be important in the positioning of the bicyclic ring corresponding to the β bend. This bicyclic ring, although not aromatic, forms an edge-toface contact with Trp215. One of the phenyl rings shows hydrophobic contact with lle174, while the other shows no significant interactions with thrombin. By comparison to other inhibitors complexed to thrombin, FPAM appears to have a new binding mode that differs from that of substrate, or hirudin, or argatroban.

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Figure 5 Schematic representation of the principal intermolecular interactions of a fibrinogen peptide A mimetic within the active site of thrombin.

Obst et al. [13], departing from the peptide template, designed and synthesized novel nonpeptide inhibitors of thrombin. They began with a cyclic template having attachment sites for three side chains that would be complementary to the S1, S2, and S3 sites in thrombin. Important to the design was a rigid template that would avoid hydrophobic collapse of the side chains and loss of conformational degrees of freedom upon complex formation with thrombin. Using computational approaches (Insight II/Discover/CVFF force field), possible templates were modeled within the active site of thrombin. These studies resulted in the synthesis of thirteen analogs that shared a common template. The most active molecule (Ki = 90 nM, 8-fold selective versus trypsin) was studied further by x-ray crystallography (Figure 6). The positively charged benzamidine binds into the S1 pocket of thrombin forming a bidentate hydrogen bond with Asp189. The proximal carbonyl of the rigid template acts as a hydrogen-bond acceptor for the amide NH of Gly216. The methylene dioxybenzyl group at P3 interacts with thrombin in two ways. An edge-to-face interaction was observed with Trp215, and an oxygen of the methylenedioxy group acts as an acceptor for a hydrogen bond with the OH hydrogen of Tyr60A. Recent communications from Bristol-Myers Squibb [14,15] describe peptidomimetic inhibitors (Figure 7) that were designed to bind thrombin with an N- to C-polypeptide chain sense opposite that of the substrate and form interactions similar to those made by the first three residues of hirudin (lle1, Thr2, Tyr3). In the x-ray crystal structure of BMS-183507 (Ki = 17.2 nM) with thrombin [15], the N terminus is facing the catalytic site while the methyl ester is

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Figure 6 Schematic representation of the principal intermolecular interactions of a nonpeptide bicyclic inhibitor within the active site of thrombin.

exposed to solvent. No specific interactions were observed with the catalytic triad. A bound water molecule hydrogen bonded to the Ser195 hydroxyl. The complex is stabilized by a network of hydrogen bonds as well as hydrophobic interactions. The Phe1-O and the Phe3-NH form hydrogen bonds with Gly216, and the Phe1-NH hydrogen bonds to the backbone carbonyl of Ser214. The Phe1 phenyl group occupies the S2 site, while Phe3 interacts within the S3 site. The retro-inhibitors contain a 4-guanidinobutanoyl group that extends into the S1 specificity site. Rather than forming two hydrogen bonds between the guanidine and Asp189 in a manner similar to PPACK, BMS-183507 forms only one, with the second hydrogen bond being directed to the carbonyl oxygen of Gly219. Binding affinity, as evidenced by loss of more than two orders of magnitude in affinity on addition of one or two methylene groups, was sensitive to chain length at this position. The allo-Thr hydroxyl oxygen accepts a hydrogen bond from the backbone NH of Gly219. This additional interaction accounts, at least in part, for the increase in affinity when compared to the inhibitor with Leu in this position. Comparison of the crystal structures of thrombin complexed with BMS-183507 and with hirudin reveals that the hirudin residue, Thr2, and the allo-Thr of BMS-183507 interact differently with thrombin. The hirudin Thr2 binds at S2,

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Figure 7 Schematic representation comparing the principal inhibitor-to-thombin interactions of related inhibitors with either Leu (7a) or allo-Thr (7b) at P3.

whereas the allo-Thr sidechain is oriented toward the protein exterior and is partially exposed to solvent.

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Cyclotheonamide A (CtA), a macrocyclic marine natural product derived from the Japanese sponge, Theonella sp., inhibits thrombin with an IC50 value of

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Figure 8 Schematic representation of the principal intermolecular interactions of cyclotheonamide A within the active site of thrombin.

100 nM and represents a novel structural class of serine protease inhibitors. An x-ray crystal structure of CtA complexed with thrombin was used to determine the molecular basis for this inhibition (Figure 8 [16]). The Arg-Pro unit binds to the S1 and S2 sites in a manner similar to the Arg-Pro of PPACK. The Arg guanidinium group forms a bidentate hydrogen bond with Asp189 while the Pro establishes a βsheet interaction with the Ser214-Gly216 backbone. The α-ketoamide acts as a transition-state mimetic forming a tetrahedral hemiketal with the hydroxyl of Ser195. Within the complex, CtA adopts a relatively open conformation with the Pro orthogonal to the macrocycle and confined by a hydrophobic pocket defined by Tyr60A, Trp60D, and Leu99. Two aromatic residues are involved in stacking interactions with Tyr60A and Trp60D. Cyclotheonamide A, however, does not effectively match the S3 interactions provided by the D-Phe group found in PPACK. In CtA, the formamide group is too polar to effectively complement the S3 site adjacent to Trp215. The authors note that the complex of CtA with thrombin does not appear optimal and suggest that synthetic analogs could significantly improve both potency and selectivity. Starting with the known thrombin inhibitors Argatroban and Nα(2-naphthyl-sulfonyl-glycyl)-DL-pamidinophenylalanyl-piperidine (NAPAP), a group at Roche initiated a medicinal chemistry program to develop thrombin inhibitors with reduced toxicity and an improved hemodynamic profile [17].

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The discovery program proceeded in four iterative phases which are shown in Table 1. Initial screening of low molecular weight organic bases led to the discovery of 1-amidinopiperidine (1–1) as a new surrogate for the guanidine and amidine functionality in Argatroban and NAPAP, respectively. A distinct advantage of 1-amidinopiperidine is its intrinsic selectivity for thrombin over

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trypsin. Application of three-fold iterative strategy of design involving synthesis, x-ray crystallography, and molecular modeling, this group elaborated the 1-amidinopiperidine from structures that inhibited in the micromolar range to some inhibiting in the picomolar range. In doing so significant improvements in the selectivity of thrombin relative to trypsin were also achieved. In the case of the D-amino acid series (1–2), a “second inhibitor binding mode” that differed from that of Argatroban was identified. In this new and unexpected binding mode, the S2 pocket is unoccupied and the napthalenesulfonyl group fills the S3 site and overlaps the front of the S2 site. The benzyl group of the phenylalanine is oriented toward the protein surface and is partially exposed to solvent. The Argatroban or “inhibitor binding” mode was favored by the more potent L-amino acid series (1–3 and 1–4) where the piperidide (1–3) or Nbenzyl (1–4) binds to the S2 site and the aryl groups are found in the S3 site. IV. Bivalent Thrombin Inhibitors Directed at the Fibrinopeptide a Binding Pocket and the Fibrinogen Recognition Site A strategy to prepare highly selective thrombin inhibitors involves linkage of molecules capable of interacting at distinct subsites. This approach should result in inhibitors more specific for thrombin: while serine proteases possess common structural features related to catalysis and some serine proteases—including the coagulation enzyme Factor Xa—also exhibit primary substrate specificity for positively charged residues, only thrombin possesses recognition subsites for fibrinogen and effector molecules such as thrombomodulin. Nature has used this strategy in the evolution of hirudin, the anticoagulant protein produced by the medicinal leech. When this effective anticoagulant binds thrombin [18–20], the N-terminal domain blocks the primary specificity pocket while the C-terminal residues adopt an extended conformation and make multiple interactions within the fibrinogen recognition exosite. Guided by structural and biochemical information, small molecules capable of simultaneous interactions with both the primary specificity pocket and the fibrinogen recognition exosite were designed and synthesized. These bivalent inhibitors are composed of three regions: a group to block the primary specificity pocket, a sequence to bind the fibrinogen recognition site, and a chemical linker. The bivalent inhibitor approach was first executed with peptides [21–22]. In 1990, DiMaio et al. (3–3 [22]) used the peptide sequence from hirudin to link (d-Phe)-Pro-Arg-Pro, known to bind at the primary specificity pocket [23], with hirudin C-terminal residues, known to bind at the fibrinogen recognition site. Polyglycine linkers were also used to connect these sequences (Maraganore

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et al. [21]). Among these hirudin analogs, the tetraglycine linker appeared optimal (3–8, Ki = 2.3 nM, Table 2). Most of the peptide-based bivalent inhibitors were slowly cleaved by thrombin. Incorporation of a ketomethylene pseudo peptide bond (3–4) resulted in a noncleavable bivalent inhibitor that retained high thrombin affinity [24]. Decreased proteolysis in bivalent inhibitors increasingly nonpeptide in character continues to be observed. Chemically simpler linkers were made using multiple methylene-containing glycine variants [25]. The dependence of affinity on placement of amide linkage within linkers containing the same number of atoms indicated some specific thrombin-to-linker interactions (3–13,14,15,16). This was confirmed in the crystal structure of hirutonin-6:thrombin complex (3–26 [26]) where continuous electron density was observed for the entire bivalent inhibitor including the linker region. The extended nature of the fibrinogen recognition site complicates attempts to reduce inhibitor molecular weight while maintaining affinity. Although of similar molecular weight, substitution of the sequence -Asp-Tyr-Glu-Pro-lle-Pro-Glu-Glu-Ala-cyclohexylalanine-(D-Glu) for -Asp-Phe-Glu-Glu-llePro-Glu-Glu-Tyr-Leu-Gin increases affinity an order of magnitude (compare 3–17 and 3–18). Within a series of bivalent inhibitors, inclusion of sulfated tyrosine, the naturally occurring residue of hirudin, increases affinity 5 to 6 fold (3–8 compared to 3–11, and 3–1 to 3–2). Only seven residues are present in one of the smallest bivalent inhibitors (3–26). Increasingly nonpeptide substituents have been incorporated into the primary specificity pocket binding portion of the bivalent inhibitors. Higher affinity for thrombin was achieved by replacement of the (DPhe)-Pro-Arg with either dansyl-Arg-(D-pipecolic acid) (3–17, [27]) or 4-tert-butylbenzenesulfonyl-Arg(D-pipecolic acid) (3–18, [27]). While the arginine side chain of these and the (D-Phe)-Pro-Argcontaining inhibitors make similar interactions with the aspartic acid within the S1 specificity pocket, the dansyl-Arg-(D-pipecolic acid) inhibitors bind in a nonsubstrate mode [27]. This initial result suggests that other nonpeptide thrombin inhibitors may be successfully incorporated into bivalent inhibitors. Recently, a pyridinium methyl ketone bivalent inhibitor capable of forming a reversible covalent complex with thrombin was synthesized (3–26, [28]). Crystallographic analysis of its complex with thrombin showed the ketone carbonyl becomes tetrahedrally coordinate by bonding to the side chain of thrombin's active site residue, Ser195. Substitutions of cyclohexylalanine for phenylalanine (3–4 compared to 3–5) and the cyclohexylalanine-containing fibrinogen recognition peptide for the hirudin sequence (3–17 compared to 3–18) also contribute to the increased affinity of this bivalent inhibitor.

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V. Inactivated Thrombin as an Inhibitor of Clot Formation A means to selectively inhibit thrombin's role in coagulation while preserving its anticoagulant functions involves site-directed mutagenesis of thrombin itself. By introduction of a single mutation, Gibbs et al. [29] altered thrombin's relative specificity for fibrinogen and protein C. The engineered thrombin's increased activation of protein C over fibrinogen cleavage offers the possibility of inhibiting clot formation with a modified human protein, a molecule likely to exhibit few side effects. VI. The Role of Structural Information The discovery of thrombin inhibitors has benefited from available protein structural information. Models of the thrombin overall structure and its active site geometry, constructed from available structures of related serine proteases [30], aided in the design of the mechanism-based inhibitors such as PPACK [31] and its boroarginine analog [10]. The unexpected, nonsubstrate binding mode of early thrombin inhibitors such as NAPAP was revealed by x-ray crystallographic analyses [32]. Iterative structurebased design methods have been critical in the optimization of bivalent inhibitors and inhibitors directed at the primary specificity pocket. Structures of inhibitor:thrombin complexes are essential for the optimization of substitutents forming interactions within the aryl-binding site of the primary specificity pocket. In some cases (e.g. Table 1), seemingly minor alterations of the inhibitor can result in dramatic changes in the inhibitor's overall interactions with thrombin [17]. Drug discovery efforts have also been strongly influenced by results of structural studies of thrombin complexed with effectors and substrate peptides. For example, recently the structures of thrombin complexed with fibrinopeptide A [33] and human prothrombin fragment F1 [34] have been determined. In addition to their role in design of high-affinity inhibitors, these structures provide valuable insights for design of drugs specific for the various subsites and conformational states of thrombin. VII. Conclusion Discovery of therapeutically effective thrombin inhibitors involves issues such as affinity and selectivity, bioavailability, and formulation. In addition to these relatively common concerns, the complex in vivo mechanisms designed to

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balance its pro- and anticoagulant activities present additional challenges in the discovery of therapeutically effective thrombin inhibitors. References 1. Tapparelli C, Metternich R, Ehrhardt C, Cook NS. Synthetic low-molecular weight thrombin inhibitors: molecular design and pharmacological profile. TIPS 1993; 14:366–376. 2. Stubbs MT, Bode W. Structure and specificity in coagulation and its inhibition. Trends Cardiovasc Med 1995; 5:157–166. 3. Stone SR. Thrombin Inhibitors: A new generation of antithrombotics. Trends Cardiovasc. Med. 1995; 5:134–140. 4. Harker LA. Strategies for inhibiting the effects of thrombin. Blood Coagulation and Fibrinolysis 1994; 5:47–58. 5. Claeson G. Synthetic peptides and peptidomimetics as substrates and inhibitors of thrombin and other proteases in the blood coagulation system. Blood Coagulation and Fibrinolysis 1994; 5:411–436. 6. Bode W, Mayr I, Baumann U, Huber R, Stone SR, Hofsteenge J. The refined 1.9 Å crystal structure of human α-thrombin: interaction with D-Phe-Pro-Arg chloromethylketone and significance of the TyrPro-Pro-Trp insertion segment. EMBO J. 1989; 8:3467–3475. 7. Bode W, Turk D, Karshikov A. The refined 1.9-Å X-ray crystal structure of D-Phe-Pro-Arg chloromethylketone-inhibited human α-thrombin: structure analysis, overall structure, electrostatic properties, detailed active-site geometry, and structure-function relationships. Protein Science 1992; 1:426–471. 8. Stubbs MT, Bode W. A Player of many parts: the spotlight falls on thrombin's structure. Thrombosis Research. Vol. 69. Pergamon Press, 1993; 1–58. 9. Whinna HC, Church FC. Interaction of thrombin with antithrombin, heparin cofactor II and protein C inhibitor. Journal of Protein Chemistry 1993; 12:677–688. 10. Weber PC, Lee S-L, Lewandowski FA, Schadt MC, Chang C“ H, Kettner C. Kinetic and crystallographic studies of thrombin with Ac-(D)Phe-Pro-boroArg-OH and its lysine, amidine, homolysine and ornithine analogs. Biochemistry 1995; 34:3750–3757. 11. Deadman JJ, Elgendy S, Goodwin C, Green D, Baban J, Patel G, Skordalakes E, Chino N, Claeson G, Kakkar V, Scully M. Characterization of a class of peptide boronates with neutral P1 side chains as highly selective inhibitors of thrombin. J Med Chem 1995; 38:1511–1522.

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12. Wu T-P, Yee V, Tulinsky A, Chrusciel R, Nakanishi H, Shen R, Priebe C, Kahn M. The structure of a designed peptidomimetic inhibitor complex of α-thrombin. Protein Engineering 1993; 5:471–478. 13. Obst U, Gramlich V, Diederich F, Weber L, Banner DW. Design of novel, nonpeptidic thrombin inhibitors and structure of a thrombin-inhibitor complex. Angew Chem Int Ed Engl 1995; 34:1739. 14. Iwanowicz EJ, Lau WF, Lin J, Roberts DGM, Seiler SM. Retro-binding tripeptide thrombin active inhibitors: discovery, synthesis and molecular modeling. J Med Chem 1994; 37:2122–2124.

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15. Tabernero L, Chang CÁ, Ohringer SL, Lau WF, Iwanowicz EJ, Han W-C, Wang T, Seiler S, Roberts D, Sack JS. Structure of a retro-binding peptide inhibitor complexed with human α-thrombin. J Mol Biol 1995; 246:14–20. 16. Maryanoff BE, Qiu Z, Padmanabhan KP, Tulinsky A, Almond HR, Andrade-Gordon P, Greco M, Kauffman J, Nicolaou KC, Liu A, Brung P, Fusetani N. Molecular basis for the inhibition of human αthrombin by the macrocyclic peptide cyclotheonamide A. Natl Academy Sci USA 1993; 90:8048–8052. 17. Hilpert K, Ackermann J, Banner DW, Gast A, Gubernator K, Hadvary P, Labler L, Muller K, Schmid G, Tschopp T, Van de Waterbeemd H. Design and synthesis of potent and highly selective thrombin inhibitors. J Med Chem 1994; 37:3889–3901. 18. Rydel TJ, Tulinsky A, Bode W, Ravichandran KG, Huber R, Roitsch R, Fenton JW, II. The structure of a complex of recombinant hirudin and human α-thrombin. Science 1990; 249:277–280. 19. Grutter MG, Priestle JP, Rahuel J, Grossenbacher H, Bode W, Hofsteenge J, Stone SR. Crystal structure of the thrombin-hirudin complex: a novel mode of serine protease inhibitor. EMBO J 1990; 9:2361–2365. 20. Rydel TJ, Tulinsky A, Bode W, Huber R. Refined structure of the hirudin-thrombin complex. J Mol Biol 1991; 221:583–601. 21. Maraganore JM, Bourdon P, Jablonski J, Ramachandran KL, Fenton JW, II. Design and characterization of hirulogs: a novel class of bivalent peptide inhibitors of thrombin. Biochemistry 1990; 29:7095–7101. 22. DiMaio J, Gibbs B, Munn D, Lefebvre J, Ni F, Konishi Y. Bifunctional thrombin inhibitors based on the sequence of hirudin. J Biol Chem 1990; 265:21698–21703. 23. Kettner C, Shaw E. D-Phe-Pro-Arg CH2C1—A selective affinity label for thrombin. Thromb Res 1979; 14:969–973. 24. DiMaio J, Ni F, Gibbs B, Konishi Y. A new class of potent thrombin inhibitors that incorporates a scissile pseudopeptide bond. FEBS 1991; 282:47–52. 25. DiMaio J, Gibbs B, Lefebvre J, Konishi Y, Munn D, Yue SY. Synthesis of a homologous series of ketomethylene arginyl pseudodipeptides and application to low molecular weight hirudin-like thrombin inhibitors. J Med Chem 1992; 35:3331–3341. 26. Zdanov A, Wu S, DiMaio Y, Konishi Y, Li Y, Wu X, Edwards B, Martin P, Cygler M. Crystal structure of the complex of human α-thrombin and nonhydrolyzable bifunctional inhibitors, hirutonin-2 and hirutonin-6. PROTEINS: Structure, Function and Genetics 1993; 17:252–265.

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27. Tsuda Y, Cygler M, Gibbs BF, Pedyczak A, Fethiere J, Yue SY, Konishi Y. Design of potent bivalent thrombin inhibitors based on hirudin sequence: incorporation of nonsubstrate-type active site inhibitors. Biochemistry 1994; 33:14443–14451. 28. Rehse PH, Steinmetzer T, Li Y, Konishi Y, Cygler M. Crystal structure of a peptidyl pyridinium methyl ketone inhibitor with thrombin. Biochemistry 1995; 34:11537–11544. 29. Gibbs CS, Coutre SE, Tsiang M, Li WX, Jain AK, Dunn KE, Law VS, Tao CT, Matsumura SY, Mejza SJ, Paborsky LR, Leung LLK. Conversion of thrombin into an anticoagulant by protein engineering. Nature 1995; 378:413–416. 30. Greer J. Comparative model-building of the mammalian serine proteinases. J Mol Biol 1981; 153:1027–1042.

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31. Kettner C, Shaw E. D-PHE-PRO-ARGCH2Cl-1 selective affinity label for thrombin. Thrombosis Research 1979; 14:969–973. 32. Brandstetter H, Turk D, Hoeffken W, Grosse D, Sturzebecher J, Martin PD, Edwards BFP, Bode W. X-ray crystal structure of thrombin complexes with the benzamidine- and arginine-base inhibitors NAPAP, 4-TAPAP and MQPA: a starting point for elaborating improved antithrombotics. J Mol Biol 1992; 226:1085–1099. 33. Martin PD, Robertson W, Turke D, Bode W, Edwards BFP. The structure of residues 7–16 of the Aα-chain of human fibrinogen bound to bovine thrombin at 2.3 Å resolution. J Biol Chem 1992; 267:7911–7920. 34. Arni RK, Padmanabhan K, Padmanabhan KP, Wu TP, Tulinsky A. The structure of the non-covalent complex of prothrombin kringle 2 with PPACK-thrombin. Chem Phys Lipids; 1994; 67–68:59–66. 35. Stone SR, Hofsteenge J. Kinetics of the inhibition of thrombin by hirudin. Biochemistry 1986; 25:4622–4628. 36. Witting JI, Bourdon P, Maraganore JM, Fenton JW II. Hirulog-1 and -B2 thrombin specificity. Biochem J 1992; 287:663–664. 37. Bourdon P, Jablonski J, Chao BH, Maraganore JM. Structure-function relationships of hirulog peptide interactions with thrombin. FEBS 1991; 294:163–166. 38. Szewczuk Z, Gibbs BF, Yue SY, Purisima E, Zdanvo A, Cygler M, Konishi Y. Design of a linker for trivalent thrombin inhibitors: interaction of the main chain of the linker with thrombin. Biochemistry 1993; 32:3396–3404.

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11 Design of Antithrombotic Agents Directed at Factor Xa William C. Ripka Corvas International, Inc., San Diego, California I. Introduction Serine proteases have long been recognized as important players in a number of biochemical processes and their specific and selective inhibition provides multiple therapeutic opportunities [1]. In particular, the blood coagulation process is the result of an amplified cascade of proteolytic events in which several specific zymogens of serine proteases in blood are activated sequentially by selective cleavages to produce active enzymes [2,3]. This process, in pathological circumstances, may lead to the formation of a thrombus—an insoluble matrix of fibrin and platelets. Thrombosis is a serious medical problem in the United States and Europe as exemplified by the fact that half the people who die each year die of cardiovascular related problems. While much recent work in antithrombotic therapeutic approaches has focused on inhibition of thrombin, the central role that Factor Xa plays in the coagulation response to vascular injury also makes it an ideal pharmacological target for antithrombotic drug development. The recent report of the x-ray crystal structure of native Factor Xa [4] allows, for the first time, a wellfounded structure-based drug design approach for inhibitors. A number of reviews describing the biology [5–10] and chemistry [11,12] of Factor Xa inhibitors have appeared. II. Coagulation Cascade In the coagulation cascade (Figure 1), a highly amplified process leads to the formation of thrombin, which is the primary mediator for the conversion of fibrinogen to fibrin, as well as activation of platelets through the thrombin

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Figure 1 The coagulation cascade.

receptor. Thrombin generation is itself, however, the result of Factor Xa in complex with Factor Va on a phospholipid surface (the prothrombinase complex) acting on prothrombin. Vascular injury is the initiating event in the coagulation process, causing the activation of Factor Xa by the Factor VIIa/tissue factor complex. Factor Xa is, therefore, a central and crucial enzyme directly leading to the production of thrombin and its inhibition should be effective in blocking thrombogenesis. As a consequence of its key role early in the coagulation cascade process Factor Xa represents a potentially valuable therapeutic target for potent and specific inhibition. III. Proof of Principle for a Factor Xa Inhibitor In recent years the method by which certain hematophageous organisms maintain blood flow during feeding has been determined. Interestly, several of these organisms utilize Factor Xa inhibitors to prevent coagulation [13–15]; the tick anticoagulant peptide (TAP), a small protein isolated from the Ornithidoros moubata tick [13], and antistasin isolated from the Haementeria officinalis leech [14] are both potent and selective inhibitors of Factor Xa. As expected, these molecules are effective antithrombotics in several animal models of thrombosis (Table 1) and provide an important proof of principle with regard to the potential effectiveness of Factor Xa inhibitors as therapeutic anticoagulants.

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Page 267 Table 1 Factor Xa Inhibitors (TAP, Antistasin) in Experimental Models of Thrombosis Rat and rabbit models of venous thrombosis [25] Canine model of high shear, coronary arterial thrombosis [6,26] Canine model of femoral arterial thrombosis [27,28] Rhesus monkey model of acute disseminated intravascular coagulation [29,30] Baboon model of platelet dependent arterial thrombosis [9,31,32]

In an alternate approach, it has been shown that a covalently blocked, activesite-modified Factor Xa (DEGR-Xa) [16] as well as a catalytically impaired recombinant form [17] can be effective anticoagulants in models of deep-vein thrombosis [18] and in canine arterial thrombosis models [8]. In these examples, the active-site-inactivated Factor Xa competes with the active form for incorporation into the active prothrombinase complex since the binding of Factor Xa to Factor Va in this complex is independent of the active site [20]. These studies with Factor Xa inhibitors suggest that inhibiting earlier in the coagulation cascade, as well as inhibiting the production of thrombin by inactivating Factor Xa in the prothrombinase complex, may have certain therapeutic advantages. IV. Factor Xa—Structure and Function Factor Xa is a 59 kilodalton protein synthesized in the liver and secreted into the blood as an inactive zymogen (Figure 2) [21]. Prior to secretion the singlechain molecule undergoes co- and posttranslational modifications including removal of a signal sequence [22–24], gamma carboxylation of several glutamic acids (Gla) in the N-terminus [33], beta hydroxylation of Asp63 [34], N-glycosylation at two sites [35], and cleavage at two sites, Arg139 and Arg142, to give a two-chain molecule [36]. The mature form of Factor X consists of a light chain (139 amino acids) and a heavy chain (303 amino acids) held together by a single disulfide (Figure 2). The Gla residues are responsible for calcium and phospholipid binding and the second EGF domain is thought to mediate binding to Factor VIIIa and Factor Va [37,38]. The heavy chain contains the catalytic domain with the prototypic serine protease active site triad, His226, Asp279, and Ser376. During coagulation, Factor X is converted to the active protease, Factor Xa, by a complex of Factor VIIa/tissue factor or a complex of Factor IXa/Factor, VIIIa/phospholipid, and calcium, both of which cleave a specific Arg-Ile bond to release an activation peptide (Figure 2) [39]. Similar to the activation of chymotrypsin, trypsin, and thrombin, the newly formed N-terminal Ile folds into the interior of the protein to form an ion pair at the active site with Asp]375 [39,40]. In the presence of calcium ions the newly formed Factor Xa associates with Factor Va on a phospholipid membrane surface to form the

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Figure 2 Factor Xa structure. Residues of the catalytic triad (His226, Asp279, Ser376) are circled.

prothrombinase complex that rapidly converts prothrombin (PT) to thrombin by cleavage at two sites in PT, Arg271–Thr272 and Arg320–Ile321 [41]. The x-ray structure of human des(1–45) Factor Xa at 2.2 Å resolution has now been reported [4] (Figure 3). V. Natural Inhibitors of Factor Xa Several small, potent, and naturally occurring Factor Xa inhibitors—tick antico-agulant protein (TAP)[3], tissue factor pathway inhibitor (TFPI) [42], antistasin (ATS) [43], Ecotin [44,45]—have been isolated and characterized (Table 2). All but TAP apparently inhibit the enzyme in the extended substrate conformation referred to as the standard mechanism of inhibition (Figure 4) [47]. In this standard mechanism the inhibitor presents a conformationally constrained binding loop with a partial beta sheet motif to the target enzyme that mimics the required substrate conformation and, after binding to the enzyme, can undergo a reversible proteolytic hydrolysis at the reactive site peptide bond (P1–P1').

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Figure 3 Stereo ribbon diagram of Factor Xa [4]. Residues of the catalytic triad are shown (His57, Asp102, Ser195) as well as residues of specific interest for the binding of small molecules to the active site: Glu192; S4 pocket residues Tyr99, Phe174, Trp215; and S1 pocket residue, Asp189. Residues are designated with the chymotrypsin numbering.

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Page 270 Table 2 Naturally Occurring Inhibitors of Factor Xa Inhibitor

Source

Ki

Structural information

TFPI

Human

3 pM [49] 90 nM [50]

Ecotin

Escherichia coli

50 pM [44]

TAP

Ornithidoros moubata (tick)

Antistasin

Haementeria officinalis (leech)

61 pM [51]

(X-ray in progress) [52]

AcAP5

Ancylostoma caninum

43 pM [19]

homology to Ascaris lumbricoides var.suum [79,80]

135 pM [13]

X-ray; complex with trypsin [46] 2D-NMR [57,58]

Studies of these natural inhibitors can be useful in defining the active site requirements for Factor Xa inhibition, and importantly, can indicate the level of inhibition that may be necessary for an effective Factor Xa inhibitor, recognizing that TAP and antistasin have evolved to yield functional, in vivo antithrombotics. Table 3 shows the reactive-site sequences of these substratelike inhibitors as well as the cleavage site sequences recognized by Factor Xa in the activation of prothrombin (PT), Factor VII, and Factor V. A. Tissue Factor Pathway Inhibitor (TFPI) The mature tissue factor pathway inhibitor (TFPI) is a 276-residue protein consisting of three tandom domains with homology to the Kunitz-like protease

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Figure 4 Extended binding modes for substrates and inhibitors. Sites in the enzyme (S) and in the inhibitor (P) are designated by the Schecter-Berger notation [48].

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Page 271 Table 3 Active Site Sequences for Factor Xa Substrates and Inhibitors Substrates

P4

P3

P2

P1

P'1

P'2

P'3

P'4

PT271,272

Ile

Glu

Gly

Arg

Thr

Ala

Thr

Ser

PT320,321

Ile

Asp

Gly

Arg

Ile

Val

Glu

Gly

FVII

Pro

Gln

Gly

Arg

Ile

Val

Gly

Gly

FV

Lys

Lys

Tyr

Arg

Ser

Leu

His

Leu

Antistasin

Val

Arg

Cys

Arg

Val

His

Cys

Pro

Ecotin

Val

Ser

Thr

Met

Met

Ala

Cys

Pro

TFPI-II

Gly

Ile

Cys

Arg

Gly

Tyr

Ile

Thr

AcAP5

Cys

Arg

Ser

Arg

Gly

Cys

Leu

Leu

AcAP6

Cys

Arg

Ser

Phe

Ser

Cys

Pro

Gly

Inhibitors

inhibitors [42]. A potent inhibitor of both Factor VIIa and Xa as well as trypsin, TFPI does not, however, have significant activity against leukocyte elastase, urokinase, activated Protein C, tissue factor plasminogen activator, thrombin, or kallikrein [53,54]. The second Kunitz domain from the Nterminus of TFPI has been identified as primarily responsible for the Factor Xa inhibition while both the first and second domains contribute to inhibition of Factor VIIa [42] (Figure 5). The proposed mechanism for this Factor-Xa-dependent inhibition of FVIIa/tissue factor involves the formation of a quarternary FXa-TFPI-FVIIa/TF complex [42]. The recombinant, isolated second domain, TFPI-II, has a Ki for Factor Xa of 90 nM [50] compared to 3 pM [49] for the intact protein. The sequence of the P4–P5' region of the Factor Xa inhibitory second Kunitz domain (Table 3) has been incorporated into a prototypic Kunitz inhibitor, bovine pancreatic inhibitor (BPTI), to produce potent and selective Factor Xa inhibitors [75,76]. B. Antistasin (ATS) Antistasin is one of several anticoagulants isolated from the Mexican leech, Haementeria officinalis [14]. It is a 119-amino-acid cysteine-rich protein with a primary structure that shows a two-fold sequence symmetry suggesting the molecule possesses two separate and distinct domains [51]. Mutagenesis studies have shown that ATS binds to Factor Xa in a substratelike manner in the P3(Arg32–P'3(Cys37) regions and is cleaved only in the first domain [55].

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While the x-ray structure of antistasin has not been reported, it is known that a Factor-Xa-induced cleavage occurs between Arg34 and Val35 suggesting this peptide loop conforms to the conformationally rigid substratelike conformation suggested by other known protein inhibitors of serine proteases [55]. A hallmark of this mode of inhibition is the rigid structure around the cleaved

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Figure 5 Predicted secondary structure of tissue factor pathway inhibitor (TFPI) showing the Factor Xa and Factor VIIa inhibitory domains. The arrows point to the P1 sites.

bond, often imposed by cysteine crosslinks constraining the two ends of the cleavage site to be in close proximity even after cleavage [47]. Antistasin has cysteines at the P2(Cys33) and P'3 (Cys37) positions. C. Tick Anticoagulant Peptide (TAP) The tick anticoagulant peptide (TAP) is a 60-amino-acid polypeptide isolated from the soft tick Ornithodorus Moubata and is a potent (Ki = 2–200 pM) and selective inhibitor of Factor Xa, both as the free enzyme and in the prothrombinase complex [13]. The TAP anticoagulant does not inhibit trypsin or other trypsinlike serine proteases and, importantly, is not cleaved by Factor Xa. The mechanism by which TAP inhibits Factor Xa appears to be unique and it apparently does not utilize the substratelike binding modes characteristic of antistasin and the Kunitz inhibitors. Mutagenesis studies have shown that the primary interaction of TAP with Factor Xa occurs at the N-terminous where Arg3 appears to play a key role [56]. The solution structure of TAP has been deter-

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Figure 6 Stereo diagram of the NMR solution structure of the tick anticoagulant peptide. The crucial N-terminus Arg3 is indicated along with the pattern of cysteine bonds.

mined by 2D-NMR studies [57,58] (Figure 6). With the exception of the region neighboring the Cys15—Cys39 bond in TAP, these studies support the originally proposed idea that there are significant structural similarities between TAP and Kunitz proteinase inhibitors [10]. Nevertheless, it is clear that TAP inhibitors FXa by a fundamentally different, and as yet, not fully understood mechanism. D. AcAP's In addition to ticks and leeches, other hematophagous organisms such as hook- worms have also evolved potent and selective Factor Xa inhibitors as anticoagulant strategies. Two such proteins, AcAP5 and AcAP6, have been isolated from the Ancylostoma caninum hookworm [19]. The AcAP5 protein is a 77amino-acid polypeptide with 10 cysteine residues. It inhibits the amidolytic activity of Factor Xa with a Ki of 43 ± 5 pM. Incubation of rAcAP5 with its target enzyme Factor Xa results in partial cleavage of the Arg40–Gly41 peptide bond suggesting the sequence around this cleavage site can adopt the restricted conformational requirements of substrates [47]. The AcAP6 protein is a 75-amino-acid polypeptide, also with cysteines, with a Ki for Factor Xa inhibition of 996 ± 65 pM. Alignment of the sequences of AcAP5 and AcAP6 suggest the P1 residue in AcAP6 is Phe38 and not the basic residue usually associated with Factor Xa specificity [19]. Substitution of Phe38 in AcAP6 with Arg resulted in a mutant that inhibited Factor Xa with a potency similar to rAcAP5. Both AcAP6 and ecotin suggest that an Arg or

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Lys in the P1 position is not an absolute requirement for potent and selective activity. E. Ecotin Ecotin, a protein isolated from Escherichia coli, is a promiscuous protease inhibitor that potently inhibits kallikrein, urokinase, Factor XIIa, granzyme B, trypsin, chymotrypsin, and elastase (reviewed in Reference 46). As with most protein inhibitors (hirudin [59] and TAP [10] being the exceptions) ecotin presents a ‘baitlike’ substrate sequence to the target protease resulting in a reversible cleavage of the P1–P1' peptide bond. However, unlike other Factor Xa inhibitors that require basic residues such as Arg or Lys in the P1 position, ecotin is cleaved between two hydrophobic amino acids, Met84 and Met85 [44]. Three preliminary crystal structures of ecotin with the serine proteases chymotrypsin, trypsin, and fiddler crab collagenase have been described [46]. The conformation of the sequence around the reactive site is similar to the bovine pancreatic trypsin inhibitor (BPTI) but differs in that the Cys at P' (not P2 as in BPTI) provides the rigidifying function for the reactive-loop sequence. Interestingly, antistasin has cysteines at both the P2 and P3' positions. The Pro at P4' is commonly found in FXa inhibitors including antistasin. In its complexes with the serine proteases for which structures are available [46] there is a sub van der Waals contact between Met84-C and the enzyme Ser195-O. The Met84-O faces the oxyanion hole and forms hydrogen bonds with Ser195 and Gly193. In the trypsin structure the Met84 side chain extends into the S1 site in a manner similar to Lys15 in the BPTI-trypsin complex. Ecotin also forms both beta sheet hydrogen bonds to the enzyme Gly216, Ser82-N and O to Gly216 O and N. VI. Small Molecule Inhibitors of Factor Xa While the x-ray structure of native Factor Xa has been reported [4] the nature of its crystal packing, specifically the fact that the active site of one Factor Xa molecule is blocked by the N-terminus of a second resulting in a “continuous polymeric structure,” apparently has precluded diffusing inhibitors into the preformed crystals to obtain complexes. Complexes with inhibitors cocrystallized with Factor Xa also have not been reported [81]. Thus, efforts to do structure-based design with this enzyme have relied on molecular modeling. Since, to date, it has not been possible to directly obtain x-ray structures of inhibitor complexes with Factor Xa, the substantial information available with respect to how serine proteases, particularly thrombin, bind inhibitors can

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be utilized to model known Factor Xa inhibitors using the x-ray coordinates of native Factor Xa. Preliminary modeling of the several inhibitors described below into the Factor Xa active site was accomplished by the author [69] by first superimposing the backbone atoms (N, Ca, C) of the catalytic triads (His57, Asp102, Ser195 in the benzamidine:thrombin and PPACK:thrombin x-ray structures on the corresponding residues in Factor Xa. This allowed an excellent fit of the P1 basic groups, arylbenzamidine in the case of benzamidine and arginine for PPACK, into the S1 pocket of Factor Xa. These groups were then used as templates for positioning the appropriate P1 basic groups of the various synthetic inhibitors [69]. Holding these docked P1 groups fixed, the remaining rotatable bonds were manipulated to allow a reasonable and complementary fit of the inhibitor atoms to the solvent accessible surface of the Factor Xa active site. In those cases where it was possible, hydrogen bonds, particularly to Gly216, were formed. To fit extended peptide sequences such as that for antistasin and the antistasinderived peptides described below, the backbone atoms of the residues around the cleavage site (e.g., P4–P4') were positioned using the corresponding BPTI backbone atoms as a template. This was done after first aligning the trypsin catalytic triad backbone in the BPTI:trypsin x-ray complex (vida infra) to Factor Xa. Where the side chains of the bound peptide segments differed from BPTI, their orientation was either modeled for maximum complementarity to the Factor Xa molecular surface or set by an algorithmic approach [78]. In some cases the structures obtained were energy minimized initially by steepest descent followed by conjugate gradient minimization. Recently, compounds based on a bisamidine motif (e.g., DX-9065a) have been reported as potent and selective Factor Xa inhibitors (1, DX-9065a) [60–63].

The position of the amidino group makes little difference to the Factor Xa potency of these compounds but, interestingly, has a dramatic effect on the selectivity towards thrombin [62]. It was also observed that one carboxylic acid isomer (CX-9065a) was 7 times more potent on Factor Xa than the other. A second set of analogs shows a similar SAR [60].

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In both cases the acids are much more selective for Factor Xa over thrombin. An initial modeling study using a homology-built Factor Xa structure proposed a fit of DX9065a to Factor Xa in which the amidinoary1 group occupies the S1 pocket and the acetimidoyl group is directed out of the S4 pocket [60]. Lin et al. [64] have provided a more systematic study of possible fits of compound 2 and DX9065a using the recently available Factor Xa coordinates [4]. After aligning the His57, Ser195, and Asp102 backbone atoms for Factor Xa and thrombin (in the benzamidine:thrombin x-ray structure [65]) the arylamidino group of 2 was superimposed on the benzamidine template and a systematic conformational search was performed on the rotatable bonds of inhibitor 2. Energetics and complementarity to the Factor Xa surface determined a saved set, about 300 low-energy conformations, for further study. The final result was an optimized structure in which the acetimino group of 2 fits into

Figure 7 Molecular modeling fit of compound 2 with the arylamidino group positioned in the S1 pocket and the acetimino group in the S4 cation-π site [69].

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the S4 pocket formed by the aromatic residues Trp215, Tyr99, and Phe174 (Figure 7). It was proposed that this collection of aryl residues forms the basis for a cation-π interaction that has recently been well documented in other cases [66]. Interestingly, this cation-π site is absent in thrombin's equivalent “aryl binding site” where the corresponding residues are Trp215, Leu99, and Ile174—residues that cannot provide the π-electrons necessary for stabilization of the cation in the inhibitor. The apparent preference for FXa inhibitors to have cations in the P3,P4 position may be directly related to the ability of the Factor Xa S4 site to stabilize these cations via a cation-π interaction. Factor Xa appears to be unique among the coagulation factors in providing this electron-rich S4 pocket (Table 4). The initial discovery of bisamidine structures as potential Factor Xa inhibitors was actually made much earlier with the finding that compounds such as 4 showed an almost 300-fold preference for Factor Xa over thrombin with a Ki of 13 nM (FXa) [67],

As with the DX-9065 analogs and compounds 2 and 3, the Factor Xa potency was relatively insensitive to the positioning of the amidino groups (4,4' versus 3,3') while replacing the 7-membered cycloalkyl ring with the 5 or 6 membered ring analogs reduced potency by about 10 fold. Model building compound 4 and docking into Factor Xa [69], again using the x-ray benzamidine:thrombin complex [65] as a template, shows that the second aryl amidino group can be positioned into the S4 aromatic pocket of Factor Xa in a conformation closely related to the mode of binding proposed for DX9065a (Figure 8) [64]. In an effort to compare the relative efficacy of thrombin versus Factor Xa inhibitors, Markwardt et al. [67] synthesized a set of amidinoaryl compounds with moderate potency as Factor Xa inhibitors (5). Table 4 S4 Residues in Selected Serine Proteases Residue Positiona

Factor Xa

Thrombin

Factor VIIa

Trypsin

99

Tyr

Leu

Thr

Leu

174

Phe

Ile

Pro

Gly

215

Trp

Trp

Trp

Trp

aChymotrypsin

numbering system. Sequence alignments by comparison of x-ray structures (sequence for Factor VIIa).

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The n=3 chain length was optimal while the nature of the tosyl group on the α- nitrogen was relatively nonspecific, with tosylGly and Nα-β-naphthylsulfonyl- Gly being of similar potency. While the phenyl group on the amide nitrogen was best, other groups were also tolerated. Although the authors did not speculate on how these compounds were bound to Factor Xa, it is reasonable to suggest that the amidino phenyl group fits in the S1 pocket similar to the orientation determined by x-ray crystallography for benzamidine in the benzamidine:thrombin x-ray crystal structure. If the aryl amidino group of 5 is matched to that of benzamidine after alignment of the catalytic triad backbone atoms (N,Cα,C) for thrombin and Factor Xa, a proposed fit of 5 to Factor Xa can be made (Figure 9) [69]. This mode of binding is consistent with the structure activity relationships observed but does not suggest the reasons for the observed small preference for Factor Xa over thrombin.

Figure 8 Molecular modeling fit of compound 4 in the Factor Xa active site [69]. The carbonyl of the cycloheptanone makes a hydrogen bond (3.12 Å) to N-Gly216.

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Figure 9 Molecular modeling fit of compound 5 in the Factor Xa active site.

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Figure 10 Proposed model of dansyl-Glu-Gly-Arg-chloromethyl ketone in Factor Xa [4,69].

Tulinsky and coworkers [4] have proposed a model for the complex of dansyl-Glu-Gly-Argchloromethyl ketone using the thrombin-PPACK crystal structure as a template for the fit to Factor Xa (Figure 10). Using antistasin as a starting point, Ohta et al. [68] have synthesized a series of cyclic peptides based on the antistasin sequence. Three of these peptides are shown below and represent the most potent in the series. Ki (FXa) ATS29-47

NH2-Ser-Gly-Val-Arg-Cys*-Arg-Val-His-Cys*-Pro-His-Gly-Phe-Gln-ArgSer-Arg-Tyr-Gly-OH

ATS29-40

NH2-Ser-Gly-Val-Arg-Cys*-Arg-Val-His-Cys*-Pro-His-Gly-OH

11.8 µM

dR-ATS32-38

NH2-dArg-Cys-Arg-Val-His-Cys-Pro-OH

0.96 µM

(The Cys*-Cys* are joined in disulfide bonds to form cyclic structures)

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0.035 µM

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These molecules are cleaved by Factor Xa suggesting they bind in a similar manner to antistasin itself. Assuming the sequence around the cleavage site occupies the FXa active site locally in a manner similar to BPTI in the BPTI:trypsin complex, a modeled structure of the complex can be constructed

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Figure 11 dArg-ATS32-38 modeled into the active site of Factor Xa utilizing the BPTI: trypsin x-ray structure as a template [69].

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using BPTI residues 11–19 as a template [69]. The modeled dR-ATS32-38:FXa complex is shown in Figure 11. Interestingly, these peptides do not inhibit trypsin even at 1000-fold higher concentrations than their FXa inhibitory concentrations. Antistasin, on the other hand, inhibits trypsin with a Ki of 10 nM. Preliminary reports have appeared describing a pentapeptide that is a potent (Ki=3 nM) and selective inhibitor of Factor Xa (SEL2711) [70,71].

Two possible modes of binding can be envisioned for this compound with either the methylpyridinium group occupying the S1 site in a “substratelike mode” or the p-amidinophenyl group in the S1 pocket, which would require a reversed binding reminiscent of the hirudin-thrombin interaction [59]. Figure 12 shows the case for the amidinophenyl group in the Factor Xa S1 pocket [69]. In this mode of binding the methylpyridinium group easily fits the S4-aryl binding site and is well positioned for a π-cation interaction. Of interest from a drug-design viewpoint is the finding that cyclotheonamide, a compound isolated from a marine sponge and originally reported as a thrombin inhibitor, has been found to also inhibit Factor Xa with a Ki of 50 nM [72]. Cyclotheonamide possesses a novel α-ketoamide transition state functionality and x-ray structures of cyclotheonamide with trypsin [73] and thrombin [74] provide templates for modeling this inhibitor into Factor Xa [69]. In the resulting fit (Figure 13) cyclotheonamide does not project functionality into the S4-cation-π site and would not be expected to show Factor Xa selectivity. VII. Defining the Requirements for Factor Xa Inhibition by Mutagenesis of BPTI It has been known for some time that many examples of naturally occurring Kunitz inhibitors exist, both isolated and as domains in larger proteins, which inhibit a variety of serine proteases [47]. This strongly suggests that this molecular framework is compatible with inhibition of this general class. The contact region between these inhibitors and their protease targets is known from a

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Figure 12 Modeled fit of SEL2711 in Factor Xa with the arylamidino group positioned in the S1 pocket and the methylpyridinium group in the cation-π S4 site.

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Figure 13 Cyclotheonamide modeled into Factor Xa utilizing the x-ray structure of cyclotheonamide:trypsin [73] and cyclotheonamide:thrombin [74] as templates [69].

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number of x-ray structures of complexes. Appropriate site-specific and random mutagenesis, particularly of the P3-P4' residues of a prototypic Kunitz inhibitor, bovine pancreatic trypsin inhibitor (BPTI), has been shown to result in potent and selective Factor Xa inhibitors [75,76]. A potent inhibitor of trypsin, kallikrein, and plasmin, BPTI does not inhibit Factor Xa. It binds to serine proteases such as trypsin in an extended substrate mode from residue 13 (P3) through 17 (P'2) [47]. A second loop from BPTI also extends into the active site bringing residues 34, 39, and 46 into contact with the protease-active site. In terms of spatial proximity of residues three clusters can be defined: cluster 1 (13,39); cluster 2 (11,17,19,34); and cluster 3 (16,18,20,46). While residue 39 is approximately in the same region of space as residue 13 (9.4 Å CB-CB) the CA rarrow.gif CB vectors are directed in different directions and substitution at 39 would not be expected to have a cooperative effect with residue 13. Residue 34 on the other hand is in a key position. It is centrally located between residues 11,17, and 19 with CB-CB distances of 5.6, 5.7, and 6.5 Å respectively, and its CA rarrow.gif CB vector converges with the corresponding vectors from these residues to a common point in space. This residue is therefore expected to have a substantial cooperative effect with the other residues of cluster 2. Finally residue 46 is close to residue 20 (CB—CB of 6.5 Å) although the CA rarrow.gif CB vectors are approximately parallel and cooperative effects are expected to be minimal. The BPTI residues 11,12,13,15–20, 34,39, and 46 were therefore the focus of the site-directed and random mutagenesis studies. Residue 14 is Cys in BPTI and was not modified in the mutants since it is required for structural reasons. A. Site Specific Mutagenesis As a starting point for the design of BPTI-based Factor Xa inhibitors, the second domain of TFPI (TFPIII) was used as a template [75,76]. Table 5 shows the results of site-directed mutagenesis of BPTI. Mutant 50cl is a direct analog of TFPI-II with the exception of the Lys at position 46. The finding that 4c2 and 4c10 are essentially equivalent in potency (Ki 2.8 versus 1.8 nM) and are identical Table 5

Site-Directed BPTI Mutants with Factor Xa Inhibition Ki(nM)

12

13

14

15

16

17

18

19

20

34

39

46

r-TFPI-II

90

Gly

Ile

Cys

Arg

Gly

Tyr

Ile

Thr

Arg

Lys

Leu

Glu

50cl

205

Gly

Ile

Cys

Arg

Ala

Tyr

Ile

Thr

Arg

Lys

Leu

Lys

4c2

2.8

Gly

Ile

Cys

Arg

Ala

Tyr

Ile

Thr

Arg

Val

Leu

Glu

4c10

1.8

Gly

Ile

Cys

Arg

Ala

Tyr

Ile

Thr

Arg

Val

Leu

Lys

57c1

1.6

Gly

Ile

Cys

Arg

Ala

Tyr

Ile

Ile

Arg

Val

Leu

Lys

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w.t. BPTI

>1 mM

Gly

Pro

Cys

Lys

Ala

Arg

Ile

Ile

Arg

Val

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Arg

Lys

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cal in sequence with the exception of the Glu46 Lys switch, suggests this residue is of minor importance. Therefore, the potency of 50cl (within a factor of 3) is consistent with r-TFPI-II. On the other hand, the incorporation of Val for Lys at position 34 leads to a dramatic increase in potency (˜100 fold; cf. 4c10 and 4c2 with 50c1). From cluster 2 above this suggests the importance of P5, P'2,P'4 for potency. The changes in wild type BPTI that result in a potent Factor Xa inhibitor are Lys15 rarrow.gif Arg15, Arg17, rarrow.gif Tyr17, and Arg39 rarrow.gif Leu39. B. Random Mutagenesis 11Libraries of mutant BPTI were created by inserting mutagenic cassettes in the BPTI gene of filamentous phage PIII coat proteins [76]. These libraries produced large numbers of mutants (~106) with randomized amino acids in positions 11, 13, 16, 17, 18, 19, 20, 34, and 39. The mutants were panned against Factor Xa, which was affixed to a solid support by a nonneutralizing antibody and the most potent inhibitors were separately expressed as soluble proteins. By this process it was possible to determine consensus sequences at the reactive sites and to define the pharmacophore requirements of inhibitors of Factor Xa in both a functional and conformational sense from the P4 to the P5 positions. Inhibitor amino acid preferences from both site directed and random mutagenesis studies are shown in Table 6. VIII. Positional Requirements of Factor Xa Inhibitors (Table 6) Examination of models of BPTI-mutants bound to Factor Xa show the L-amino acids in the P3 position project into solvent. In the Factor Xa cleavage sites in thrombin these residues are polar and acidic (Glu, Asp); they are polar and basic in antistasin (Arg), and polar and neutral in Ecotin (Ser). The exception is TFPI- II, with this position occupied by Ile. The BPTI random mutant results are consistent with the TFPI-II case and show a preference for aliphatics or aromatics in this position. There is a hydrophobic pocket in the enzyme, formed by Trp215, Tyr99, and Phe174, that would be accessible to a D-residue in this position. The accessibility of the S2 pocket of the enzyme by P2 groups would be expected to be influenced by the orientation of Tyr99. As the x-ray structure shows its position this residue puts severe limitations on the size of the P2 group. Consistent with this is the observation that Gly is the sole residue in the FXa:thrombin cleavage sites. Little information is available from the BPTI mutants, TFPI-II, or antistasin, which all have a structural requirement for Cys at this position. Synthetic compounds show, however, that in potent inhibitors large bulky aromatics are, in fact, allowed at this position, a situation that requires Tyr99 to move out of the way [75].

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Additionally, the binding of BPTI mutants also requires Tyr99 to move because of potential severe interactions with the Cys14–Cys38 bridge of the BPTI mutants. While larger groups can be accommodated, structural requirements are fairly rigid in agreement with the limited mobility expected of the Tyr99. The apparent requirement for a P2-Gly in the larger substrates may suggest that in these cases extended binding occurs that does not permit movement of Tyr99. In the P1 position, as expected, a basic group is preferred. Interestingly, of the two naturally occuring basic amino acids, arginine is preferred over lysine. This is seen in both the BPTI mutants as well as the Arg rarrow.gif Lys switch in antistasin. This may be due to Ala190 in the S1 pocket of FXa, which cannot orient and stabilize the BPTI lysine analog as Ser190 does in trypsin. An interesting exception to the need for a basic group is in Ecotin where a methionine occupies this site. The x-ray of Ecotin with trypsin clearly shows this neutral residue in the P1 pocket, aligned very closely to that seen for lysine in the BPTI:trypsin complex, and proximal to the charged Asp189 [77]. Apparently, extended binding over the rest of the site compensates for this energetically unfavorable situation. In the P'1 position, the natural cleavage sites use Thr and Ile while Ecotin has Met. In contrast to thrombin, FXa lacks the 60-insertion loop and can accommodate large groups at this position. The BPTI mutants, however, are forced to use a small residue (Ala) because of steric hindrance from the Cys58Cys42 group residue 61 in the enzyme. The inhibitor TFPI-II has a Gly at P'1. The BPTI mutants, TFPI-II, and antistasin all show a preference for aromatic groups at P'2. In the BPTI panning experiments Tyr was selected more than 80% of the time at this position. It can be seen from Table 5 that the Arg to Tyr change at position 17 is one of three significant changes that converts wild type BPTI from a non-Factor Xa inhibitor to a ˜1.6 nM inhibitor. There is a possible hydrogen-bond interaction between Tyr17 of the inhibitor and Gln192 of the enzyme, which may explain the strong preference. While models suggest the P'3 residue is directed at solvent and the FXa thrombin cleavage sites have polar residues at this position (Thr, Glu), the BPTI mutant results show a clear preference for a hydrophobic group. It is possible that aromatic groups can pack to Phe41 of the enzyme. Mutant results show Ile is favored over Phe, His, which in turn is selected over Tyr.

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IX. Conclusions Factor Xa is clearly an important component of the coagulation process and inhibition of this enzyme can lead to potent anticoagulant effects. Recently, a number of naturally occurring anti-Factor-Xa polypeptides have been isolated from several hematophagous organisms including ticks, leeches, and hook- worms. With the exception of TAP, these molecules appear to bind to Factor Xa by the standard mechanism of inhibition proposed earlier [49]. The sequence information derived from these inhibitors as well as the natural cleavage sites of substrates of Factor Xa can be used along with the conformational constraints imposed by the proposed substrate-like binding to define the pharmacophore requirements of the active site of Factor Xa. The structurally rigid BPTI mutants, which have been found to be potent Factor Xa inhibitors, also provide important conformational information particularly with regard to the specific binding interactions on the P' side of the Factor Xa active site. A number of small molecule inhibitors have also recently been reported which appear to take advantage of a unique cation-π S4-site available in Factor Xa to achieve good selectivity with moderate potency. The availbility of the X-ray structure of native Factor Xa has allowed molecular modeling approaches to suggest possible fits of these inhibitors to the Factor Xa active site. Note Added in Proof After this review was written, the x-ray structure of Factor Xa with DX-9065a was reported [81]. References 1. Proteinase inhibitors. In: Barrett AJ, Salvesen G, eds. Research Monographs in Cell and Tissue Physiology. Vol 12. New York: Elsevier, 1986. Design of Enzyme Inhibitors as Drugs. Sandler M, Smith HJ, eds. New York: Oxford University Press, 1989. 2. Colman RW, Hirsh J, Marder VJ, Salzman EW. Hemostasis and Thrombosis. Basic Principles and Clinical Practice. Second Edition. Philadelphia: J. B. Lippincott Company, 1987. 3. Hathaway, WE, Goodnight, Jr SH. Disorders of Hemostasis and Thrombosis. New York: McGrawHill, 1993. 4. Padmanabhan K, Padmanabhan KP, Tulinsky A, Park CH, Bode W, Huber R, Blankenship DT, Cardin AD, Kisiel W. Structure of human Des(1–45) factor Xa at 2.2 Å resolution. J Mol Biol 1993; 232:947–966.

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5. Kaiser B, Hauptmann J. Factor Xa inhibitors as novel antithrombotic agents: facts and perspectives. Cardiovascular Drug Reviews 1994; 12 (3):225–236. 6. Lynch JJ, Sitko GR, Mellott MJ, Nutt EM, Lehman ED, Friedman PA, Dunwiddie CT, Vlasuk GP. Maintenance of canine coronary artery patency following thrombolysis with front loaded plus low dose maintenance conjunctive therapy. A comparison of factor Xa versus thrombin inhibition. Cardiovasc Res 1994; 28:78–85. 7. Hollenback S, Sinha U, Lin P-H, Needham K, Frey L, Hancock T, Wong A, Wolf D. A comparative study of prothrombinase and thrombin inhibitors in a novel rabbit model of non-occlusive deep vein thrombosis. Thromb Haemost 1994; 71:357–362. 8. Benedict CR, Ryan J, Todd J, Kuwabara K, Tijburg P, Cartwright Jr J, Stern D. Active site-blocked factor Xa prevents thrombus formation in the coronaryvasculature in parallel with inhibition of extravascular coagulation in a canine thrombosis model. Blood 1993; 81:2059–2066. 9. Schaffer LW, Davidson JT, Vlasuk GP, and Siegl PKS. Antithrombotic efficacy of recombinant tick anticoagulant peptide. A potent inhibitor of coagulation factor Xa in a primate model of arterial thrombosis. Circulation 1991; 84:1741– 1748.

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10. Vlasuk GP. Structural and functional characterization of tick anticoagulant peptide (TAP): a potent and selective inhibitor of blood coagulation factor Xa. Thrombosis and Haemostasis 1993; 70:212–216. 11. Scarborough RM. Anticoagulant strategies targeting thrombin and factor Xa. Ann Reports Med Chem 1995; 30:71–80. 12. Wallis RB. Inhibitors of coagulation factor Xa: from macromolecular beginnings to small molecules. Current Opinion in Therapeutic Patents Aug, 1993. 13. Waxman L, Smith DE, Arcuri KE, Vlasuk GP. Tick anticoagulant peptide (TAP) is a novel inhibitor of blood coagulation factor Xa, Science 1990; 248:593. 14. Hauptmann J, Kaiser B, Vowak G, Struzebecher J, Markwardt F. Comparison of the anticoagulant and antithrombotic effects of synthetic thrombin and factor Xa inhibitors. Throm Haemostas 1990; 63:220–223. 15. Jacobs JW, Cupp EW, Sardana M, Friedman PA. Isolation and characterization of a coagulation factor Xa inhibitor from black fly salivary glands. Thromb Haemost (GERMANY) 1990; 64:235–238. 16. Taylor Jr FB, Chang ACK, Peer GT, Mather T, Blick K, Catlett R, Lockhart MS, Esmon CT Blood 1991; 78:364. 17. Sinha U, Hancock T, Lin P-H, Hollenback S, Wolf D. Expression, purification, and characterization of inactive human coagulation factor Xa (Asn322Ala419). Protein Expr Purif 1992;3:518–524. 18. Dunwiddie CT, Waxman L, Vlasuk GP, Friedman PA. Purification and characterization of inhibitors of blood coagulation factor Xa from hematophagous organisms. Methods in Enzymol 1993; 233:291–312. 19. Stanssens P, Bergum PW, Gansemans Y, Jespers L, LaRoche Y, Huang S, Maki S, Messeno J, Lauwereys M, Cappello M, Hotez PJ, Lasters I, Vlasuk GP. Anticoagulant repertoire of the hookworm ancyclostoma caninum. Proc Natl Acad Sci 1996; 93:2149–2154. 20. Nesheim ME, Kettner C, Shaw E, Mann KG. Cofactor dependence of factor Xa incorporation into the prothrombinase complex. J Biol Chem 1981; 256:6537– 6540.

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21. Davie EW, Fujikawa K, Kisiel W. The coagulation cascade: interaction, maintenance, and regulation. Biochemistry 1991; 30:10363–10370. 22. Leytus SP, Foster DC, Kurachi K, Davie EW. Gene for human factor X: a blood coagulation factor whose gene organization is Essentially identical with that of factor IX and protein C. Biochemistry 1986; 25:5098–5102. 23. Fung MR, Hay CW, MacGillivray RTA. Characterization of an almost full length cDNA coding for human blood coagulation Factor X. Proc Nat Acad Sci USA 1985; 82:3591–3595. 24. Blanchard RA, Faye KLM, Barrett JM, William B. Isolation and characterization of profactor X from the liver of a steer treated with sodium warfarin. Blood 1985; 66(suppl.1):331a. 25. Vlasuk GP, Ramjit D, Fujita T, Dumwiddie CT, Nutt EM, Smith DE, Shebuski RJ. Comparison of the in vivo anticoagulant properties of standard heparin and the highly selective factor Xa inhibitors antistasin and tick anticoagulant peptide (TAP) in a rabbit model of venous thrombosis. Thromb Haemostas 1991; 65:257– 262. 26. Sitko GR, Ramjit DR, Stabilito II, Lehman D, Lynch JJ, Vlasuk GP. Conjunctive enhancement of enzymatic thrombolysis and prevention of thrombotic reocclusion with the selective factor Xa inhibitor, tick anticoagulant peptide. Comparison to hirudin and heparin in a canine model of acute coronary artery thrombosis. Circulation 1992; 85:805–815. 27. Mellot MJ, Strainieri MT, Sitko GR, Stabilito II, Lynch JJ, Vlasuk GP. Enhancement of recombinant tissue plasminogen activator-induced reperfusion by recombinant tick anticoagulant peptide, a selective factor Xa inhibitor, in a canine model of femoral arterial thrombosis. Fibrinolysis 1993; 7:195–202. 28. Mellot JJ, Holahan MA, Lynch JJ, Vlasuk GP, Dunwiddie CT. Acceleration of recombinant tissuetype plasminogen activator-induced reperfusion and prevention of reocclusion by recombinant antistasin, a selective factor Xa inhibitor, in a canine model of femoral arterial thrombosis. Circ Res 1992; 70:1152–1160.

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29. Neeper MP, Waxman L, Smith DE, Schulman CA, Sardana M, Ellis RE, Schaffer LW, Siegel PKS, Valsuk GP. Characterization of recombinant tick anticoagulant peptide. A highly selective inhibitor of blood coagulation factor Xa. J Biol Chem 1990; 265:17746–17752. 30. Dunwiddie CT, Nutt EM, Vlasuk GP, Siegel PDS, Schaffer LW. Anticoagulant efficacy and immunogenicity of the selective factor Xa inhibitor antistasin following subcutaneous administration in the rhesus monkey. Thromb Haemostas 1992; 67:371–376. 31. Schaffer LW, Davidson JT, Vlasuk GP, Dunwiddie CT, Siegel PKS. Selective factor Xa inhibition by recombinant antistasin prevents vascular graft thrombosis in baboons. Arteriosclerosis and Thromb 1992; 12:879–885. 32. Kelly AP, Hanson SR, Dunwiddie CT, Harker LA. Circulation 1992; 86:411. 33. Suttie JW. Vitamin K-dependent carboxylase. Annu Rev Biochem 1985; 54:459– 477. 34. Fernlund P, Stenflo J. β-Hydroxy-aspartic acid in vitamin K-dependent proteins. J Biol Chem 1983; 258:12509–12512. 35. DiScipio RG, Hermodson MA, Davie EW. Activation of human factor X (Stuart factor) by a protease from Russell's viper venom. Biochemistry 1977; 16:5253– 5260.

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36. Leytus SP, Chung DW, Kisiel W, Kurachi K, Davie EW. Characterization of a cDNA coding for human factor X. Proc Nat Acad Sci USA 1984; 81:3699–3702. 37. Skogen WF, Esmon CT, Cox AC. Comparison of coagulation factor Xa and des(1–44) factor Xa in the assembly of prothrombinase. J Biol Chem 1984; 259:2306–2310. 38. Hertzberg MS, Ben-Tal O, Furie BC. Construction, expression, and characterization of a chimera of Factor IX and Factor X: the role of the second epidermal growth factor domain and serine protease domain in factor Xa binding. J Biol Chem 1992; 267:14759–14766. 39. Sigler PB, Blow DM, Matthews BW, Henderson R. Structure of crystalline α- chymotrypsin. II. A preliminary report including a hypothesis for the activation mechanism. J Mol Biol 1968; 35:143–164. 40. Bode W, Turk D, Karshikov A. The refined 1.9 Å X-ray crystal structure of D- Phe-Pro-Arg chloromethylketone-inhibited human α-thrombin. Structure analysis, overall structure, electrostatic properties, detailed active site geometry, structure function relationships. Protein Sci 1992; 1:426–471. 41. Mann KG, Nesheim ME, Church WR, Haley P, Krishnaswamy S. Surface-dependent reactions of the vitamin K-dependent enzyme complexes. Blood 1990; 76:1– 16. 42. Girard TJ, Warren LA, Novotny WF, Likert KM, Brown SG, Miletich JP, Broze Jr GJ. Functional significance of the Kunitz-type inhibitory domains of lipoprotein- associated coagulation inhibitor. Nature 1989; 338:518–520. 43. Nutt E, Gasic T, Rodkey J, Gasic G, Jacobs J, Friedman P, Simpson E. The amino acid sequence of antistasin. J Biol Chem 1988; 263:10162–10167. 44. Lauwereys M, Stanssens P, Lambier AM, Messens J, Dempsey E, Vlasuk GP. Ecotin as a potent factor Xa inhibitor. Thromb Haemostasis 1993; 69:864. 45. Seymour JL, Lindquist RN, Dennis MA, Moffat B, Yansura D, Reilly D, Wessinger ME, Lazurus RA. Ecotin is a potent anticoagulant and reversible tightbinding inhibitor of factor Xa. Biochemistry 1994; 33:3949–3958. 46. McGrath ME, Gillmor SA, Fletterick RJ. Ecotin: lessons on survival in a proteasefilled world. Protein Sci 1995;4:141–148. 47. Laskowski M, Kato I. Protein inhibitors of proteinases. Annu Rev Biochem 1980; 49:593–626. 48. Schechter I, Berger A. On the size of the active site in proteases. I. Papain. Biochem Biophys Res Commun 1967; 27:157–162. 49. Peterson LC. Progress in vascular biology, haemostasis and thrombosis. Abstracts, 1992 Zimmerman Conference. San Diego, California, Feb. 27–29, 1993. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_292.html (1 of 2) [4/5/2004 5:16:15 PM]

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50. Broze Jr GJ. Trends Cardiovasc Med 1994; 2:72. 51. Hofmann KJ, Nutt EM, Dunwiddie CT. Site-directed mutagenesis of the leechderived factor Xa inhibitor antistasin. Biochem J 1992; 287:943. 52. Schreuder H, Arkema A, deBoer B, Kalk K, Dijkema R, Mulders J, Theunissen H, Hol W. Crystallization and preliminary crystallographic analysis of antistasin, a leech-derived inhibitor of blood coagulation factor Xa. J Mol Biol 1993; 231:1137–1138. 53. Broze Jr GJ, Warren LA, Novotny WF, Huguchi DA, Girard JJ, Miletich JP. The lipoproteinassociated coagulation inhibitor that inhibits the factor VII-tissue factor complex also inhibits factor Xa: insight into its possible mechanism of action. Blood 1984; 71:335–343.

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54. Broze Jr GJ, Girard TJ, Novotny WF. Regulation of coagulation by a multivalent Kunitz-type inhibitor. Biochemistry 1990; 29:7539. 55. Dunwiddie CT, Vlasuk GP, Nutt EM. The hydrolysis and resynthesis of a single reactive site peptide bond in recombinant antistasin by coagulation factor Xa. Arch Biochem Biophys 1992; 294:647–653. 56. Dunwiddie CT, Neeper MP, Nutt EM, Waxman L, Smith DE, Hoffman KJ, Lumma PK, Garsky VM, Vlasuk GP. Site-directed analysis of the functional domains in the factor Xa inhibitor tick anticoagulant peptide: identification of two distinct regions that constitute the enzyme recognition sites. Biochemistry 1992; 31:12126–12131. 57. Lim-Wilby MSL, Hallenga K, DeMaeyer M, Lasters I, Vlasuk GP, Brunck TK. NMR structure determination of tick anticoagulant peptide (TAP). Protein Sci 1995; 4:178–186. 58. Antuch W, Guntert P, Billeter M, Hawthorne T, Grossenbacher H, Wuthrich K. NMR solution structure of the recombinant tick anticoagulant protein (rTAP), a factor Xa inhibitor from the tick ornithodoros moubata. FEBS Lett 1994; 325:251–257. 59. Rydel TJ, Tulinsky A, Bode W, Huber R. Refined structure of the hirudin-thrombin complex. J Mol Biol 1991;221:583–601. 60. Katakura S-I, Nagahara T, Hara T, Iwamoto M. A novel Factor Xa inhibitor: structure-activity relationships and selectivity between Factor Xa and thrombin. Biochem Biophys Res Comm 1993; 197:965–972. 61. Hara T, Yokoyama A, Ishihara H, Yokoyama Y, Nagahara T, Iwamoto M. DX- 9065a, a new synthetic, potent anticoagulant and selective inhibitor for Factor Xa. Thrombosis and Haemostasis 1994; 71:314–319. 62. Nagahara T, Yokoyama Y, Inamura K, Katakura S-I, Komoriya S, Yamaguchi H, Hara T, Iwamoto M. J Med Chem 1994; 37:1200–1207. 63. Nagahara T, Kanaya N, Inamura K, Yokoyama Y. Aromatic amidine derivatives and salts thereof. Eur Pat App 0-540-051-A1. 64. Lin Z, Johnson ME. Proposed cation-π mediated binding by Factor Xa: a novel enzymatic mechanism for molecular recognition. FEBS Lett 1995; 370:1–5. 65. Banner DW, Hadvary P. Crystallographic analysis of 3.0 Å resolution of the binding to human thrombin of four active-site directed inhibitors. J Biol Chem 1991; 266:20085–20093. 66. Dougherty DA. Cation-π interactions in chemistry and biology: a new view of benzene, Phe, Tyr, and Trp. Science 1996; 271:163–167.

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67. Sturzebecher J, Sturzebecher U, Vieweg H, Wagner G, Hauptmann J, Markwardt F. Synthetic inhibitors of bovine factor Xa and thrombin comparison of their anticoagulant efficiency. Thrombosis Res 1989; 54:245–252. 68. Ohta N, Brush M, Jacobs JW. Interaction of antistasin-related peptides with factor Xa: identification of a core inhibitory sequence. Thromb Haemostasis (GERMANY) 1994; 72:825–830. 69. The preliminary modeled structures of the synthetic inhibitors described in this review were constructed and energy minimized by the author using HyperChem (1995, Hypercube, Inc., Release 4.5). Docking of these inhibitors to the active site of Factor Xa was accomplished by the author using the x-ray coordinates of native Factor Xa [4] and INSIGHT II (Biosym Technologies, Inc.) and the approach outlined in Section 6.

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70. Seligmann B, Stringer SK, Ostrem JA, Al-Obeidi F, Wildgoose P, Walser A, Safar P, Safarova A, LoCascio A, Spoonamore J, Thorpe DS, Kasireddy P, Ashmore B, Strop P. SEL 2711: a specific, orally available, active-site inhibitor of Factor Xa discovered using synthetic combinatorial chemistry. Abstract, Sixth IBC International Symposium on Advances in Anticoagulants and Antithrombotics. Washington, D. C., Oct. 23–24, 1995. 71. Al-obeidi F, Lebl M, Safar P, Stierandova A, Strop P, Walser A. Factor Xa inhibitors, Patent Appl. WO 95/29189; 1995. 72. Lewis SD, Ng AS, Balwin JJ, Fusetani N, Naylor AM, Shafer JA. Inhibition of thrombin and other trypsin-like serine proteinases by cyclotheonamide A Thrombosis Research 1993; 70:173–190. 73. Lee AY, Hagihara M, Karmacharya R, Albers MW, Schreiber, SL, Clardy J. Atomic structure of the trypsin-cyclotheonamide A complex: lessons for the design of serine protease inhibitors. J Am Chem Soc 1993; 115:12619. 74. Marynoff BE, Qui X, Padmanabhan KP, Tulinsky A, Almond Jr HR, Andrade- Gordon P, Greco MN, Kauffman JA, Nicolaou KC, Liu A, Brungs PH, Fusetani N. Proc Natl Acad Sci USA 1993; 90:8048. 75. Ripka W, Brunck T, Stanssens P, LaRoche Y, Lauwereys M, Lambeir A-M, Lasters I, DeMaeyer M, Vlasuk G, Levy O, Miller T, Webb T, Tamura S, Pearson D. Strategies in the design of inhibitors of serine proteases of the coagulation cascade—factor Xa. Eur J Med Chem 1995; 30 (Suppl):88s–100s. 76. Lasters I, DeMaeyer M, Ripka W. Bovine pancreatic trypsin inhibitor derived inhibitors of Factor Xa. Pat Appl WO 94/01461; 1994. 77. McGrath ME, Erpel T, Bystroff C, Fletterick RJ. Macromolecular chelation as an improved mechanism of protease inhibition: structure of the ecotin-trypsin complex. EMBO 1994; 13:1502–1507. 78. Desmet J, DeMaeyer M, Hazes B, Lasters I. Nature 1992; 356:539. 79. Grasberger BL, Clore AM, Gronenborn GM. Structure 1994; 2:669–678. 80. Huang K, Strynadka NCJ, Bernard VD, Peanasky RJ, James MNG. Structure 1994; 2:679–689.

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81. Brandstetter H, Kuhne A, Bode W, Huber R, von der Saal W, Wirthensohn K, Engh RA. J Biol Chem 1996; 47:29988–29992.

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12 Polypeptide Modulators of Sodium Channel Function as a Basis for the Development of Novel Cardiac Stimulants Raymond S. Norton Biomolecular Research Institute, Parkville, Victoria, Australia I. Introduction Cardiovascular diseases remain one of the major causes of premature death in western societies. Chronic congestive heart failure (CHF) in particular is a common disease with a poor prognosis, median survival times after the onset of heart failure being 1.7 years in men and 3.2 years in women [1]. Current treatment relies on diuretics to reduce fluid volume, vasodilators to decrease the work load of the heart, and positive inotropic agents to increase cardiac contractility [2]. The most commonly prescribed of the positive inotropes is the cardiac glycoside digoxin (Figure 1) [3]. Although this drug has been in therapeutic use for over two hundred years, its efficacy in patients with a sinus rhythm has remained controversial, and evidence for its beneficial effects is quite recent [3–5]. It is also possible that these beneficial effects are not due solely to the positive inotropic activity of digoxin and that its neurohormonal effects may also be important [2, 5–7] Nevertheless, digoxin remains a widely used drug [3] and it follows that a suitable replacement or adjunct would find access to a significant market worldwide. The incentive to develop such a replacement follows from the low therapeutic index of digoxin [8,9] and the relatively common occurrence of side effects due to digitalis toxicity. In the 1960s and 1970s, 20–30% of patients receiving digitalis experienced serious toxicity and about one quarter of this group died [6, 10]. Digitalis toxicity is manifest in CNS side-effects such as

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Figure 1 Structures of the positive inotropes digoxin [3,4], DPI 201-106 [15], and BDF 9148 [15–17]. In digoxin the R group is (O-2,6-dideoxy-β-D-ribo-hexopyranosyl-(1 rarrow.gif 4)-O-2, 6-dideoxy-β-D-ribo-hexopyranosyl-(1 rarrow.gif 4)-2, 6-dideoxy-β-D-ribo-hexopyranosyl)oxy. In DPI 201-106 the configuration at the hydroxyl-bearing carbon influences cardiac activity.

fatigue, visual disturbances, and anorexia, and in cardiac side-effects that depend on the nature and extent of the underlying heart disease [3]. Careful monitoring of digoxin serum levels and bioavailability have reduced the incidence of digitalis toxicity [3] and the recent introduction of digoxin-binding antibodies or antibody fragments has provided an effective means of treating severe digitalis toxicity [3,7]. Nevertheless the quest continues for a substitute for the cardiac glycosides in the treatment of chronic CHF. Positive inotropic compounds can be classified into three groups: cAMP generators, intracellular calcium regulators, and modulators of ion channels or pumps [11]. The cAMP generators such as dopamine, dobutamine, and milrinone (a phosphodiesterase inhibitor) may worsen ischemia, cause arrhythmias, and increase mortality [2,6]. Intracellular calcium modulators have not reached clinical use, possibly because of additional effects such as vasoconstriction,

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whereas, calcium sensitizers such as EMD 57033 may be useful positive inotropic compounds, even in the diseased myocardium [12]. Ion channel modulation represents another approach to positive inotropy [13]. Sodium channel modulators increase Na+ influx and prolong the plateau phase of the action potential; sodium/calcium exchange then leads to an increase in the level of calcium available to the contractile elements, thus increasing the force of cardiac contraction [13,14]. Synthetic compounds such as DPI 201-106 and BDF 9148 (Figure 1) increase the mean open time of the sodium channel by inhibiting channel inactivation [15]. Importantly, BDF 9148 remains an effective positive inotropic compound even in severely failing human myocardium [16] and in rat models of cardiovascular disease [17]. Modulators of calcium and potassium channel activities also function as positive inotropes [13], but in the remainder of this article we shall focus on sodium channel modulators. II. The Anthopleurins Two decades ago “drugs from the sea” were the subject of high expectations and a good deal of effort in various centers around the world. The number of therapeutically useful compounds to have emerged from that effort has been rather limited, but with the advent of high-throughput screening it is likely that useful new leads will be found, even from species investigated previously. Notwithstanding, some valuable leads did emerge from work carried out in the 1970s, amongst which were the polypeptide cardiac stimulants known as the anthopleurins. These were isolated from sea anemones, where they are components of the animal's venom and are believed to have a function in defense and the capture of prey. The work that led to the isolation and characterization of these and related polypeptides from sea anemones is covered in earlier reviews [18,19] and will not be reiterated here. The best characterized of the anthopleurins is anthopleurin-A (AP-A), which was isolated from the northern Pacific sea anemone Anthopleura xan- thogrammica and consists of 49 residues cross-linked by three disulfide bonds [18,20]. It is active as a cardiac stimulant at nanomolar concentrations in vitro, making it some 200 fold more potent on a molar basis that digoxin. Its positive inotropic activity is not associated with any significant effects on heart rate or blood pressure [21], and in conscious dogs its therapeutic index is 7.5, which is about three-fold higher than that of digoxin [8]. Anthopleurin-A is active under conditions of stress and hypocalcaemia [18,22], as well as in ischemic myocardium where many other positive inotropes give equivocal results [23]. The profile of activity for AP-A suggests that it is a potentially valuable lead in the development of an alternative positive inotrope to digoxin

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for use in the treatment of chronic CHF. This chapter describes how this development is being tackled using the approach of structure-based drug design. A. Related Sea Anemone Toxins Anthopleurin-A is a member of a family of sea anemone polypeptides [24] (Figure 2) that is steadily increasing in number. These polypeptides have been classified into two groups, designated Types 1 and 2 [27], which are similar with respect to the locations of their disulfide bridges and a number of residues thought to play a role in biological activity or maintenance of the tertiary structure [24,27], but are distinguishable on the basis of sequence similarity (>>60% within each type but <30% between the two types) and immunological cross-reactivity. Figure 2 shows the amino acid sequences of the Type 1 toxins, which have much stronger affinities for cardiac tissue than the Type 2 toxins characterized to date [24,27]. Apart from AP-A, the best characterized of these polypeptides with respect to its biological activity is Anemonia sulcata toxin II (ATX II) [19]. This molecule is also cardioactive [28], as would be expected from its similarity to AP-A. Renaud et al. [29] have compared the activities of a number of sea anemone and scorpion toxins on isolated rat atria and found that anthopleurin-B (AP-B, also known as Ax II) had the highest potency and the greatest margin between the concentrations necessary for maximal inotropic activity and for provoking arrhythmias (0.3 versus 10 nM). It was also found that sodium channels of rat cardiac cells in culture, which have a low affinity for tetrodotoxin (TTX), have a particularly high affinity for Type 1 anemone toxins [29], whereas Type 2 toxins [30] and scorpion toxins [31] had similar affinities for TTX-sensitive and TTX-insensitive channels in rat neuroblastoma cells and skeletal myotubes, respectively. The polypeptide ATX II has been found to have class III antiarrhythmic activity, indicating its potential in the management of cardiac arrhythmias [32]. A positive inotropic drug that was also effective as an antiarrhythmic might offer significant advantages therapeutically [33,34]. B. Site of Action The sea anemone polypeptides act by delaying inactivation of the myocardial voltage-gated sodium channel [35], thereby prolonging the action potential. Binding sites for a number of different agents, mainly toxins, on the sodium channel have been identified [35–37], the sea anemone toxin binding site being designated site 3. Other polypeptides that also bind to site 3 (or a partially overlapping site) are the short anemone polypeptides [19,24] such as ATX III [38] and PaTX [39], which are neurotoxic to crustacea, the Anemonia sulcata

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Figure 2 Amino acid sequences of Type 1 sea anemone polypeptides. The residue numbering system is based on the sequences of AP-A and AP-B. Literature references are from Norton [24], except in the case of recently published sequences for Bc III [25], Bg II and Bg III [26]. Identical residues are shaded in grey and conserved residues are boxed. The sequences of Bg II and Bg III around residue 28 could also be aligned to bring the Gly-Cys sequence into register with the other toxins if the Arg were treated as an insertion. Toxins are named in this figure according to their genus and species; common names relevant to the text are Ax I = AP-A; Ax II = AP-B; As Ia = ATX Ia.

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polypeptides BDS I and II [40,41], which were claimed to have antihypertensive and antiviral activity, and the α-scorpion toxins [24,27,42]. As discussed below, we may expect to obtain some useful information from comparisons of the molecular surfaces of these different classes of polypeptides, but the utility of this approach depends on the extent to which their binding sites are identical and not just partially overlapping, as well as the issue of channel sub-type specificity of the different toxins. The synthetic agent DPI 201-106 has been evaluated extensively as a potential replacement for digoxin [43]. It is a potent positive inotrope that also acts by delaying inactivation of the sodium channel, but its binding site appears to be distinct from that of ATX II [44]. Moreover, it exerts antihypertensive and local anesthetic effects and may also antagonise the calcium channel [43]. At present we are not aware of any low molecular mass compound that binds to the same site as the anthopleurins. This offers the prospect that a mimetic based on the anthopleurins might have a pharmacological profile distinct from other positive inotropes. The structure of the receptor for the anthopleurins, the α-subunit of the voltage-gated sodium channel, is known only in schematic form [35–37]. As illustrated in Figure 3, it contains four homologous domains, each consisting of six transmembrane regions (assumed to be helices) designated S1 to S6. The S4 segments are thought to act as the voltage sensors of the channel. All four

Figure 3 Schematic of the α-subunit of the voltage-gated sodium channel, based on its amino acid sequence [35–37]. The transmembrane segments S1–S6 in each domain are thought to form helices—with the positively charged S4 segment acting as a voltage sensor—and the S5–S6 loop of each domain is thought to contribute to the transmem-brane pore. Site 3 includes regions of the S5–S6 loops of domains I and IV, and the inactivation gate (h) is located on the intracellular segment linking domains III and IV.

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domains contribute to formation of the transmembrane pore, which is believed to be lined by short segments from the loops linking S5 and S6 in each domain. Binding site 3 is located on the extracellular surface of the channel and involves regions of the loops between S5 and S6 in domains I and IV [37,45]. The binding sites for several modulators of sodium-channel activity, including the blocker tetrodotoxin, have been mapped quite precisely onto the sequence (and thus the structural model of the channel) by combining information derived from comparisons of naturally occurring variants of the channel and from site-directed mutagenesis [37]. A similar approach could be applied to the definition of site 3, but in the absence of a crystal structure for the α-subunit the three- dimensional structure of this site will remain speculative. Therefore current attempts to design a low molecular mass analogue of the anthopleurins must be based on the structure of the ligand rather than the receptor. C. Development of a New Positive Inotrope Being polypeptides, the anthopleurins have limited therapeutic potential in their own right, as they are not active following oral administration and are antigenic in experimental animals [18]. Recent advances in the field of peptide mimetics, however, lend credence to the concept of harnessing the favorable cardiotonic properties of the anthopleurins in a low molecular mass, nonpeptide, synthetic compound. The goal of such a development would be to retain the activity of the parent polypeptides in a molecule that had good bioavailability and was not antigenic. Ideally, it might also be possible to increase the cardiac selectivity of such a compound. In order to achieve this goal, a knowledge of the three-dimensional structures of the anthopleurins and their structure-function relationships is essential. Significant progress has been made towards these goals over the past few years and we are now in a position to commence analogue design and synthesis. The following sections in this chapter summarize our knowledge of the cardioactive pharmacophore of the anthopleurins and the prospects for mimicking this in a nonpeptide moiety. Aspects of the pharmacological profile which such a compound would need to display in order to be useful in CHF therapy are also discussed. III. 3D Structure The structures in aqueous solution of both AP-A [46] and AP-B [47] have been solved using highresolution 1H NMR data. Structures have also been determined for the Type 1 toxin ATX Ia [48] and the Type 2 toxin Sh I [49,50] from NMR data. The main secondary structure element in each of these structures is a

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Figure 4 Richardson-style diagram of the polypeptide backbone of the individual structure of AP-A [46] that is closest to the average over the whole molecule. The locations of the sulfurs in the three disulfide bonds (4–46, 6–36, and 29–47) are shown in CPK format. The locations of reverse turns found in more than half the NMR-derived structures (6–9, 25–28, and 30–33) are indicated by darker backbone shading. The diagram was generated using MOLSCRIPT [51].

four-stranded, antiparallel β-sheet linked by three loops, as illustrated for AP-A in Figure 4. The first of these loops, spanning residues 8–16 in AP-A and 8–17 in AP-B, is the largest and least well defined in solution (Figure 5), although it contains several residues that are essential for activity, as described in the next section. Differences between the structures of AP-A, ATX Ia, and Sh I have been noted [46] but the overall picture that emerges is one of similar backbone folds for all three molecules, making it likely that differences among the potencies and species-specificities of these toxins are due to the presence or absence of particular side chains rather than significant structural differences. Given that ATX Ia and Sh I have weak or negligible activities on mammalian nerve and heart tissue [24,27], we have focused on the anthopleurins in an effort to define the structure of the cardioactive pharmacophore of the sea anemone polypeptides. There are several challenges in this endeavor. One is that the structures are based on NMR data that, because of the paucity of NOE restraints for surface residues compared with those in the core of the structure, do not define the locations of the solvent-exposed side chains very precisely (although it should be borne in mind that this may be a more accurate picture of the actual structure in solution than one in which the side chains are fixed in a single

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Figure 5 Stereo views of 20 structures of AP-B [47] superimposed over the backbone heavy atoms (N, Cα, C) of residues 2–7 and 17–49. The three disulfide bonds are shown in lighter shading. The lower view is related to the upper one by an approximately 180° rotation about the vertical axis.

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location). Another is that both the backbone and the side chains of residues 8– 16 are less well defined than the bulk of the structure due to a lack of medium- and long-range NMR restraints between residues in this loop and the rest of the molecule, as illustrated in Figure 5. These problems are exacerbated in the case of AP-A and AP-B by the presence of multiple conformers in solution, one cause of which is cistrans isomerism about the Gly40-Pro41 peptide bond [52]. The additional peak overlap caused by these conformers limited the number of NOE restraints that could be obtained from the spectra. Finally, the structures available at present are for the free ligand and we have no information on how much these structures might change upon binding to the sodium channel. One way of addressing the issue of a lack of precision in the locations of functionally important side chains is to determine the range of conformational space available to them in different ligands. To this end, we undertook a detailed comparison of the structures of AP-A and AP-B in solution [47]. This proved to be a useful exercise both in terms of defining the positions of side chains known to be important for cardiotonic activity and identifying neighboring residues which might also be involved [47]. Models have been described in the literature for AP-B [53] and Bunodosoma granulifera toxins II (Bg II) [26]. The AP-B model was derived from the structure of Sh I using energy minimization and the Bg II model from that of BDS I using energy minimization and 10 ps of dynamics. In both cases the calculations appear to have been carried out for the molecule in vacuo without the use of a distancedependent dielectric, under which conditions the positions of the charged side chains on the surface are likely to be distorted. Visual comparison of the model of AP-B [53] with the experimentally determined solution structure [47] indicates significant differences in side-chain orientations. IV. Residues Essential For Cardiotonic Activity Information about which residues are essential for the cardiac stimulatory activity of the sea anemone toxins has been obtained from selective chemical modification and proteolysis studies, comparisons among naturally occurring sequences, and, most recently, site-directed mutagenesis. Although there are some discrepancies among the inferences drawn from different studies and different techniques, a consensus is emerging regarding the location of the cardioactive pharmacophore. Ideally, only effects on cardiac tissue should be considered, but doing so would exclude some useful data on activity against mammalian nerve preparations. However, data obtained on the Type 2 toxins or on the activity of Type 1 toxins on nonmammalian tissues will not be discussed in detail.

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A. Chemical Modification A number of chemical modification studies have been carried out on the sea anemone toxins. As discussed previously [24], the results of these studies have to be interpreted with some caution because of less than rigorous characterization of the reaction products in many cases. Nevertheless, in the Type 1 toxins it appears that one or both of the Asp7 and Asp9 carboxylates are required for activity, as well as one or both of the Lys37 and Lys48 ε-ammonium groups [24,27,54]. The C-terminal carboxylate appears not to be essential, whereas the N-terminal ammonium appears to have some role, although the various studies give a confusing view of its importance. There are also conflicting data on the importance of His34 and His39 but it seems that at least one of them might be important. Both are located in the vicinity of other residues found to be necessary for activity, but His39 is in close contact with Asp7 and Lys37 and on this basis may be expected to be the more important. Although the available evidence points to a role for one or both of the Asp7 and Asp9 carboxylates in cardiotonic activity, it has not been established that either residue makes contact with the sodium channel. As indicated above, the carboxylate of Asp9 participates in a hydrogen bond to the backbone amide of Cys6, so its role may be structural. The carboxylate of Asp7 is close to the side chains of Lys37 and His39 and is exposed to the solvent, making it a more likely candidate for direct interactions with the sodium channel. The only evidence for its importance, however, is indirect, coming from the observation that its replacement by Asn in synthetic Sh I abolished toxicity to crabs [55]. Considerable confusion has surrounded the role of Arg14, which is conserved throughout the Type 1 and Type 2 toxins. A recent study has shown, however, that modification of Arg14 in AP-A with 1,2cyclohexanedione under conditions where the positive charge is maintained did not affect positive inotropic activity [54]. This study also showed indirectly that any contact the Arg14 side chain makes with the sodium channel must be relatively loose: although the adduct is active, it is no longer susceptible to tryptic proteolysis, indicating that the modified side chain cannot be accommodated in the substrate binding site of the protease. The conclusion that the positive charge on Arg14, but not its exact spatial location, might be important for activity is consistent with the results of site-directed mutagenesis experiments discussed below. B. Selective Proteolysis When AP-A was treated with trypsin only the Arg14 to Gly15 peptide bond was cleaved [56]. The resulting derivative lacked cardiotonic activity but its binding affinity for the rat brain sodium channel was reduced by less than an order of magnitude (Llewellyn LE et al., unpublished results). Its overall structure, as

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monitored by NMR, was unchanged, although it should be noted that the structure of the loop containing Arg14 was not well defined in either native or trypsinised AP-A. That the backbone structure was largely unaffected was confirmed by the observation that the NH and CαH chemical shifts were unaltered except in the immediate vicinity of the cleavage (which is the site of a reverse turn in AP-A [46]) and the N-terminal regions of the loop and the second strand of the sheet. It is possible that perturbations of functionally important groups near the start of the loop may be responsible for the lack of activity of the trypsinised derivative, and now that a high-resolution structure is available for AP-A a more detailed comparison with the cleavage product would be useful. Other possible explanations for the lack of activity are that the position of the Arg14 side chain relative to other key residues in the molecule is more important than suggested by chemical modification and site-directed mutagenesis data, or that the introduction of additional charges associated with the new termini affects activity. Endoproteinase LysC cleaved AP-A between Lys37 and Ala38 to yield a derivative with cardiotonic activity an order of magnitude lower than that of the parent molecule (Monks SA and Norton RS, unpublished results). This reduction in activity could be a consequence of local conformational perturbations. Treatment of AP-B with carboxypeptidase B removed Lys49, resulting in only a two-fold reduction in cardiotonic activity (Monks SA and Norton RS, unpublished results). C. Sequence Comparisons Among the Type 1 toxins shown in Figure 2, the Bc and Bg toxins (from the genus Bunodosuma) form a subgroup with characteristic differences from the Anemonia and Anthopleura toxins at residues 5, 12–13 and 37–42. In addition, the Bg toxins also have Asp7 rarrow.gif Lys and Gly27 rarrow.gif Arg substitutions. A potent toxin in mice [26], the cardiac stimulatory activity of Bg II has not been reported. The potent activity of Bg II was ascribed to its abundance of positive charges [26]. Ignoring the histidines, which at least in AP-A and ATX II would be predominantly in their neutral forms at physiological pH [57], Bg II has six positively charged side chains and only one negatively charged side chain, whereas, AP-A has three and two respectively, and AP-B has five and two. As discussed below, however, it may be that an abundance of positive charge is associated with a lack of discrimination between the neuronal and cardiac sodium channels, as found for the scorpion α-toxins. Comparison of the activities of Af I and Af II is useful because of their close similarity. Differences between Af I and Af II are Ala3 rarrow.gif Pro, Asn12 rarrow.gif Ser, Thr21 rarrow.gif Ile, and an additional Gly at the N-terminus. Inspection of the sequences in Figures 2 shows that the identity of residue 3 correlates with that of residue 21, Ala or Ser at 3 co-occurring with Thr21, and Pro 3 with Ile21

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(the exception is ATX Ib [19,24], which has Pro 3 and Thr21, but it is not known if this has the same local structure). Residues 3 and 21 are juxtaposed in the β sheet [46–48] and the effect of switching Ser3/Thr21 to Pro3/Ile21 can be assessed by comparing the structures of AP-A [46] and AP-B [47]. It appears that this leads to a distortion of the sheet between the second and third residues but causes no significant perturbations to the overall structure. In the calculated structures of AP-A a Va12 NH rarrow.gif Leu22 CO hydrogen bond was found but this was not present in the AP-B structures, presumably as a result of the local differences. Comparing Af I and Af II, the combination of the Ala3/Thr21 to Pro3/Ile21 switch plus the addition of an extra Gly at the N-terminus of Af II would be expected to alter the local conformation at the N-terminus. The fact that the cardiac stimulatory activities of the two are the same [58] therefore implies that the N-terminus is not important functionally. This inference is conditional on the effect on activity of the only other difference between Af I and Af II, Asn12 rarrow.gif Ser, being minimal. In the Type 1 toxins, Ser is found more often than Asn at position 12, while in the Type 2 toxins, replacement of Asn12 by Tyr increases toxicity in mice by a factor of two [24,27]. Thus, it is possible that the presence of Ser in Af II slightly favors cardiac activity while the changes at the N-terminus might reduce it slightly; the main conclusion to be drawn, however, is that neither change has a significant effect. The lack of effect of changes at the N-terminus is consistent with the results from expression of AP-B [59], where a Gly-Arg extension at the N-terminus had no effect on activity. At seven locations AP-B differs from AP-A. Two of these, Ser3 rarrow.gif Pro and Thr21 rarrow.gif Ile, were discussed above. Two more, Leu24 rarrow.gif Phe and Thr42 rarrow.gif Asn, are conservative changes located in or at the start of loops that are not in the immediate vicinity of the sodium channel binding surface and would not be expected to have a direct effect on activity. This leaves Ser12 rarrow.gif Arg, Val13 rarrow.gif Pro, and Gln49 rarrow.gif Lys, which do have functional significance, as discussed in the following section. It is important to note that there are several residues that are common to all of the long toxins and serve to maintain the biologically active conformations of these molecules. The clearest examples of residues in this category are the six half-cystines, although we believe that the Gly10-Pro11 sequence may influence the structure and flexibility of the Arg14-containing loop [46] (at the other end of this loop Thr17 and/or Ser19-Gly20 may also be important). Also conserved, Trp33 is probably important structurally even though its surroundings are different in the Type 1 and Type 2 toxins. It is also interesting that in ATX Ia, which is a potent crustacean neurotoxin but a poor mammalian cardiac stimulant, Lys37 and both histidines are missing, suggesting that one or more of these side chains may be important in promoting specificity for the mammalian cardiac sodium channel.

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D. Site-Directed Mutagenesis The complete synthesis of AP-A has been reported [60] but so far this approach has not been pursued to generate analogues. More productive has been the analysis of analogues produced by site-directed mutagenesis, following the successful cloning and expression of AP-B [59]. A series of single-site mutations [61,62] showed that R14A, K48A, and K49A had affinities for the cardiac sodium channel that were, respectively, only 3.2-, 2.9-, and 2.4-fold lower than that of native AP-B, while R12A had an 8.5-fold lower affinity. These results suggested that amongst the cationic side chains only Arg12 was significant for activity. Recent data on double mutations [53] indicates that the situation is not quite that simple. For example, the R12S-R14Q double mutant had a 72-fold lower affinity for the neuronal sodium channel whereas the R12S and R14Q mutants individually had 5.9- and 2.4-fold lower affinities, respectively, which should combine to produce only a 14-fold effect. It appears that the presence of one cationic side chain in the Arg14 loop may be sufficient for activity and that its exact location can vary somewhat. It would be interesting to know if the same applies to the C-terminus, where Lys48 and Lys49 may be able to compensate for one another. However, the proposal that the cationic side chains of residues 12, 14, and 49 form a cluster [53] is not consistent with the solution structure of AP-B [47]. A similar situation appears to exist in the ω-conotoxins, which possess 5–7 net positive charges. Substitution of individual cationic groups had a relatively minor effect on affinity for the voltage-gated calcium channel, whereas replacement of several had a significant effect, greater than that expected from the sum of the individual effects [63,64]. A further outcome of analyses of the double mutants was that the cationic side chains at positions 12 and 49 in AP-B seem to favor binding to the neuronal over the cardiac sodium channel. Thus, the R12SR49Q double mutant, in which residues 12 and 49 in AP-A, had a 37-fold lower affinity for the neuronal channel but only a 5-fold lower affinity for the cardiac channel relative to native AP-B [53]. In fact, the affinity of this double mutant for the cardiac channel was lower than that of AP-A, implying that one or more of the other five differences between AP-A and AP-B might decrease affinity for the cardiac channel. It seems, therefore, that while AP-A is slightly less potent than AP-B on cardiac channels, it has greater selectivity for the cardiac channel when measured in sodium flux experiments. In voltageclamp experiments AP-A and AP-B favor the cardiac channel to similar degrees [53]. V. Other Ligands for Site 3 on the Sodium Channel A. Sea Anemone Toxins Apart from the “long” sea anemone polypeptides that are the main focus of our interest, there are two other classes of anemone polypeptides that bind at or near

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site 3 on the sodium channel. The first of these are the short anemone polypeptides [19,24] such as ATX III [38] and PaTX [39], which are neurotoxic to crustacea. The polypeptide ATX III, which consists of 27 residues cross-linked by three disulfide bonds, and PaTX, which has 31 residues and four disulfides, can be aligned such that 16 residues are identical. The three disulfides in ATX III are linked in a 1–5/2–4/3–6 pattern in the same way as in the long polypeptides [65], but the only other similarity at the level of primary structure is a GCPXG sequence corresponding to residues 28–32 of AP-A. Although neurotoxic to crustacea, ATX III is inactive as a positive inotrope [28], suggesting that it possesses an appropriate structural scaffold to interact with site 3 but lacks key side chains required for interaction with the cardiac channel. Nevertheless, the smaller size of ATX III makes it an attractive candidate for further study, with the aim of engineering into it the ability to bind to the cardiac channel. The welldefined solution structure for this toxin [65] provides and essential basis for such an effort. The Anemonia sulcata polypeptides BDS I and II [40], which were claimed to have antihypertensive and antiviral activity, also bind to site 3 on neuronal sodium channels and have weak negative inotropic activity [41]. The points of similarity and difference between the solution structures of BDS I [40] and the long anemone polypeptides have been discussed previously [40,41] and will not be reiterated here; suffice to say that the overall folds are similar but the Arg14 loop in the long polypeptides is truncated in BDS I and the molecule lacks several residues that have been shown to be important for activity. B. Scorpion Toxins The scorpion α-toxins have been shown to bind to site 3 on the voltage-gated sodium channel [24,27,42]. These polypeptides contain up to 70 residues crosslinked by four disulfide bonds, but show no sequence similarity to the anemone polypeptides. Possible structural similarities have been discussed [24], and in a theoretical model of the anemone toxin Bg II, some of the cationic residues were in similar locations to those in the crystal structure of the scorpion toxin Aah II [26]. It is clear that positively charged residues play an important role in the interactions of sea anemone toxins and scorpion toxins with site 3 on the sodium channel (as indeed they do with other polypeptide toxins binding to other ion channels) but this role may be relatively more important for the TTXsensitive sodium channel in nerve and muscle than for the TTX-insensitive channel of the heart. For example, Bg II, which is more positively charged than AP-A and AP-B (see above), has a higher affinity for neuronal sodium channels [26], and replacement of Arg12 and Lys49 in AP-B with uncharged residues favors its binding to the cardiac channel [53]. Similarly, the scorpion α-toxins bind more tightly to the neuronal channel than the anemone toxins but, as with the Type 2

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anemone toxins, do not discriminate between TTX-sensitive and TTX-insensitive channels. In fact the scorpion toxins are less potent cardiac stimulants than the Type 1 anemone toxins [29]. Thus, while it will be interesting to compare the sodium channel binding surfaces of the two classes of toxin as these surfaces become better defined in each case, more useful input to the design of a positive inotrope acting at site 3 is likely to come from direct studies on the anthopleurins. VI. Defining the Cardioactive Pharmacophore Now that high-resolution structures are available for both AP-A [46] and AP-B [47], we can begin to interpret the results described above in terms of an emerging picture of the cardioactive pharmacophore of these molecules. It is encouraging that most of the residues that have been shown hitherto to be important for activity lie on one face of the structures, as shown in Figure 6. Of the residues highlighted in Figure 6a, evidence to support their role in activity on the neuronal or cardiac channels (or both) has come from chemical modification or site-directed mutagenesis studies except in the case of Asn35. The reason for including this residue is that in AP-B it is close to the Asp7/Lys37/His39 region and its side chain is hydrogen bonded to the backbone carbonyl of Lys37 [47]. Moreover, it is conserved throughout the Type 1 toxins (Figure 2). We anticipate that many of the residues highlighted in Figure 6 will participate in interactions with the sodium channel binding site. One residue which may not is Asp9, the side-chain carboxylate of which hydrogen bonds with the amide of Cys6 in AP-A and AP-B. Thus, it is possible that this residue has a “structural” role rather than a “functional” one. Other residues in the vicinity of the pharmacophore that may also have a structural role are Gly10 and Pro11, as discussed above. The side chain of Ser8 is exposed and on the same surface of the molecule, placing it in a position potentially to interact with the sodium channel; it is also conserved throughout the Type 1 toxins.

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Figure 6 (a) CPK representation of the individual structure of AP-B [47], which is closest to the average over the whole molecule, showing residues thought to contribute to the cardioactive pharmacophore. The surfaces of residues 7 and 9 are shaded black, those of 14, 37, 39, and 48 dark grey, and that of 35 lighter grey. As discussed in the text, the primary functions of Asp9 (the side chain of which is hardly visible in this view) and Asn35 may be to maintain the local structure in an active conformation, but it cannot be excluded that they also interact directly with the sodium channel. In AP-B the cationic side chains of Arg12 and Lys49 are also important, but it appears that their roles can be compensated for by nearby cationic side chains (Arg14 and Lys48, respectively) and that they favor binding to the neuronal sodium channel rather than the cardiac channel [53]. (b) Connolly surface of AP-B in the same orientation as in part a, with the charged residues Asp7, Asp9, Arg14, Lys37, and Lys48 highlighted. This figure was generated using Insight (Biosym Technologies).

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In peptide—protein and protein—protein interactions the size of the buried surface area ranges from 400 Å2 to 1400 Å2 [66]. In the potassium channel blocker charybdotoxin, 5–8 residues with surface areas of 530–850 Å2 were found to be essential for binding, depending on the type of potassium channel investigated [67,68]. In the calcium channel blocker ω-conotoxin GVIA, an alanine scan identified only two residues, Lys2 and Tyr13, that were important for activity [69]. It is likely, however, that the number of residues contributing to the binding surface is greater than this, particularly given the high affinity of this toxin for its receptor. If we consider a larger ligand such as human growth hormone, eight of the 31 residues in the growth hormone-receptor interface contribute 85% of the binding energy and more than half make no significant contribution to the affinity [70]. A similar result was found for a growth hormone-monoclonal antibody complex, where only five residues were critical for binding [71]. In fact, the example of growth hormone may be more relevant to the anthopleurins than those of charybdotoxin and conotoxin, which function simply as ion channel blockers. Thus, we expect the essential residues in the anthopleurins to number between five and ten, a total somewhat less than previously anticipated [24]. If we ignore Asp9 and assume that only one of Arg12 or Arg14 and one of Lys48 or Lys49 are necessary, then it is likely that the residues highlighted in Figure 6 make a significant contribution to the sodium channel binding surface of the anthopleurins. They span a larger area on the surface than the essential residues in charybdotoxin, but it is important to note that the conformation of the Arg14 loop in solution is not fixed and that conformations in which the Arg14 side chain is closer to the region around Asp7 could be more representative of the sodium-channel-bound structure. For example, the distance between the Arg14 guanidino group and the Asp7 carboxylate varies from 13 to 26 Å over the family of structures of AP-A and 10 to 22 Å in AP-B. By analogy with other protein—protein interactions, it is likely that the sodium channel binding surface of the anthopleurins will include side chains that mutagenesis studies will not identify as having a significant role in binding or activity. As indicated above, alanine scanning of residues in the human growth hormone—receptor interface indicated that less than a quarter of the contact residues provided most of the binding energy [70]. Thus, we believe that the residues identified above will contribute to the sodium channel binding surface of the anthopleurins by virtue of their location on the protein surface in the immediate vicinity of residues, which clearly are important for activity. Nevertheless, some of them may make only modest contributions to binding affinity and could be altered without destroying activity. Other residues, as yet unidentified, may also make significant contributions. By analogy with other protein—protein interactions characterized to date, it is reasonable to anticipate that some of these residues will be hydrophobic, in contrast to the charged

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residues that have been the main focus of attention hitherto. A key requirement now is to identify those residues that provide the majority of the binding energy and to differentiate these from others whose main role is to stabilize the binding residues in the active conformation. VII. Mimicking The Pharmacophore Significant progress has been made over the past few years in the field of peptide mimetics, although the most successful examples are those where a small peptide ligand or a linear segment of a larger protein has been the target [72–74]. An alternative approach to de novo design is to optimize a lead compound obtained by screening chemical libraries on the basis of a knowledge of the conformation of the polypeptide ligand, as in the case of the endothelin receptor antagonist SB 209670 [75]. The task of mimicking a pharmacophore is simplified where the contributing residues are contiguous in the amino acid sequence. This is not the case in the anthopleurins, with residues from at least four different regions of the sequence contributing to affinity. In charybdotoxin the essential residues come from two or three regions of the sequence, depending on which potassium channel is considered, while in growth hormone, binding site I is comprised of residues from three different regions of the protein and site II from two regions. Mimicking the pharmacophore of the anthopleurins therefore represents a task at least as challenging as those presented by these two examples. Strategies for achieving this goal include de novo design, conformationally directed data base searching and screening chemical libraries (synthetic and naturally occurring) for leads, which could then be optimized on the basis of our knowledge of the structure. Our approach is based on the first two of these. Initial attempts to mimic the pharmacophore of AP-A were based on linear and disulfide-cyclized versions of the Arg14-containing loop [76]. At that time, our level of understanding of the pharmacophore was inadequate and it is clear in retrospect that not enough of the key elements were present. Nevertheless, conformational analysis of these peptides by NMR was useful in showing that they retained several elements of local structure observed in the corresponding region of the native protein, thereby emphasizing the independence of this loop from the rest of the structure in solution. VIII. Conclusions In this chapter I have attempted to summarize the current state of our understanding of the structure and structure-function relationships of the type 1 sea

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anemone toxins and to indicate how the favorable cardiotonic properties of the anthopleurins might be mimicked in a low molecular weight analog. Progress in defining the cardiotonic pharmacophore has been hampered by difficulties in determining high-resolution structures of the anthopleurins in solution due to the presence of multiple conformers, and by uncertainties concerning the exact location of some of the key residues in the Arg14 loop. Both of these problems would have been alleviated by isotopic labeling of the molecules in a high-yield expression system, which would have allowed for better definition of the structures in solution. The question of how the structure in solution might change upon binding to the sodium channel remains open and is particularly relevant to residues in the Arg14-containing loop. Conformationally constrained analogs of the ligand offer one means of addressing this problem. The definitive solution would be provided by a high-resolution structure for the sodium channel, determined by x-ray or electron crystallography, but this will not be available in the near future. In the meantime, the approach of complementary mutagenesis (of both ligand and receptor), which has been very informative in defining the charybdotoxin-potassium channel interface [68,77], can be employed to produce a crude model of the binding site. Once a lead compound is obtained, further development will almost certainly be required to optimize properties such as bioavailability and stability in vivo. A key requirement will also be good selectivity for the cardiac over other sodium channels, but the results of mutagenesis studies carried out so far on the anthopleurins suggest that this should be achievable. Lead compounds will also have to be rigorously evaluated in terms of their effects on cardiac arrhythmias to ensure that they ameliorate rather than exacerbate this problem, especially in the failing heart. Finally, the possibility that the beneficial effects of positive inotropes in vivo may be the result of inotropic and noninotropic activities [6] would need to be evaluated for mimetics of the anthopleurins. These requirements notwithstanding, there is good reason to be optimistic that a mimetic of the anthopleurins can be developed and that it may have significant benefits in the treatment of congestive heart failure. Acknowledgments I am grateful to all the colleagues who have contributed to our sea anemone toxin work over the years, and, in particular, to Steve Monks, Paul Pallaghy, and Jane Tudor for assistance with the figures and for helpful discussions. I also thank Ken Blumenthal for communicating results prior to publication. Note Added in Proof Since completion of this chapter, two additional papers on site-directed mutations of anthopleurin-B have been published. In the first (Khera PK, Blumenthal

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KM. Importance of highly conserved anionic residues and electrostatic interactions in the activity and structure of the cardiotonic polypeptide anthopleurin B. Biochemistry 1996; 35:3503–3507), it was concluded that Asp7 may be important for folding, Asp9 may be important for protein folding and interaction with the sodium channel, and Lys37 for channel interaction. In the second (Dias-Kadambi BL, Drum CL, Hanck DA, Blumenthal KM. Leucine 18, a hydrophobic residue essential for high affinity binding of anthopleurin-B to the voltage-sensitive sodium channel. J Biol Chem 1996; 271:9422–9428), Leu18 was shown to be important for binding, with several hundred fold losses in affinity being associated with its mutation to Val or Ala. This residue is adjacent to the surface highlighted in Figure 6. References 1. Ho KKL, Anderson KM, Kannel WB, Grossman W, Levy D. Survival after the onset of congestive heart failure in Framingham heart study subjects. Circulation 1993; 88:107–115. 2. van Zwieten PA. Pharmacotherapy of congestive heart failure. Currently used and experimental drugs. Pharmacy World and Science 1994; 16:234–242. 3. Lewis RJ. Digitalis: a drug that refuses to die. Critical Care Medicine 1990; 18:S5–S13. 4. DiBianco R, Shabetai R, Kostuk W, Moran J, Schlant RC, Wright R. A comparison of oral milrinone, digoxin, and their combination in the treatment of patients with chronic heart failure. N Engl J Med 1989; 320:677–683.

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5. Packer M, Gheorghiade M, Young JB, Costantini PJ, Adams KF, Cody RJ, Smith LK, Van Voorhees L, Gourley LA, Jolly MK. Withdrawal of digoxin from patients with chronic heart failure treated with angiotensin-converting-enzyme inhibitors. N Engl J Med 1993; 329:1–7. 6. Packer M. The development of positive inotropic agents for chronic heart failure: how have we gone astray? J Am Coll Cardiol 1993; 22(suppA):119A–126A. 7. Lederer WJ, Hadley RW, Kirby MS, Eisner DA. Inotropic mechanisms in heart muscle: cardiotonic steroids—how do they work? In: Gwathmey JK, Briggs GM, Allen PD, eds. Heart failure. Basic science and clinical aspects. New York: Marcel Dekker, 1993; 349–365. 8. Scriabine A, Van Arman CG, Morgan G, Morris AA, Bennett CD, Bohidar NR. Cardiotonic effects of anthopleurin-A, a polypeptide from a sea anemone. J Cardiovasc Pharmacol 1979; 1:571–583. 9. Marsh JD, Smith TW. Epidemiology and general considerations of digitalis toxicity. In: Smith TW, ed. Digitalis glycosides. Orlando, Fla: Grune and Stratton, 1986: 217–225. 10. Beller GA, Smith Tw, Abelmann WH, Haber E, Hood WB Jr. Digitalis intoxication: a prospective clinical study with serum level correlations. N Engl J Med 1971; 284:989–997. 11. Feldman AM. Classification of positive inotropic agents. J Am Coll Cardiol 1993; 22:1223–1227.

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12. Nankervis R, Lues I, Brown L. Calcium sensitization as a positive inotropic mechanism in diseased rat and human heart. J Cardiovasc Pharmacol 1994; 24:612–617. 13. Doggrell S, Hoey A, Brown L. Ion channel modulators as potential positive inotropic compounds for treatment of heart failure. Clin Exp Pharmacol Physiol 1988; 21:833–843. 14. Briggs GM, Gwathmey JK. Role of the sodium channel in the development of force in myocardium. In: Gwathmey JK, Briggs GM, Allen PD, eds. Heart failure. Basic science and clinical aspects. New York: Marcel Dekker, 1993:597–612. 15. Hoey A, Amos GJ, Wettwer E, Ravens U. Differential effects of BDF 9148 and DPI 201-106 on action potential and contractility in rat and guinea-pig myocardium. J Cardiovasc Pharmacol 1994; 23:907–915. 16. Schwinger RHG, Böhm M, Mittmann C, La Rosee K, Erdmann E. Evidence for a sustained effectiveness of sodium-channel activators in failing human myocardium. J Mol Cell Cardiol 1991; 23:461–471. 17. Hoey A, Nankervis R, Brown L. Positive inotropic responses of the sodium channel modulator BDF 9148 in diseased rat myocardium. Clin Exp Pharmacol Physiol 1995; 22:418–422. 18. Norton TR. Cardiotonic polypeptides from Anthopleura xanthogrammica (Brandt) and A. elegantissima (Brandt). Fed Proc 1981; 40:21–25. 19. Beress L. Biologically active compounds from coelenterates. Pure Appl Chem 1982; 54:1981–1994. 20. Tanaka M, Haniu M, Yasunobu KT, Norton TR. Amino acid sequence of the Anthopleura xanthogrammica heart stimulant anthopleurin-A. Biochemistry 1977; 16:204–208. 21. Blair RW, Peterson DF, Bishop VS. The effects of anthopleurin-A on cardiac dynamics in conscious dogs. J Pharmacol Exp Ther 1978; 207:271–276. 22. Kodama I, Toyama J, Shibata S, Norton TR. Electrical and mechanical effects of anthopleurin-A, a polypeptide from a sea anemone, on isolated rabbit ventricular muscle under conditions of hypoxia and glucose free medium. J. Cardiovasc Pharmacol 1981; 3:75–86. 23. Gross GJ, Warltier DC, Hardman HF, Shibata S. Cardiotonic effects of anthopleurin-A (AP-A), a polypeptide from a sea anemone, in dogs with a coronary artery stenosis. Eur J Pharmacol 1985; 110:271–276. 24. Norton RS. Structure and structure-function relationships of sea anemone proteins that interact with the sodium channel. Toxicon 1991; 29:1051–1084.

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25. Malpezzi ELA, De Freitas JC, Muramoto K, Kamiya H. Characterization of peptides in sea anemone venom collected by a novel procedure. Toxicon 1993; 31:853–864. 26. Loret EP, Menendez Soto del Valle R, Mansuelle P, Sampieri F, Rochat H. Positively charged amino acid residues located similarly in sea anemone and scorpion toxins. J Biol Chem 1994; 269:16785–16788. 27. Kem WR. Sea anemone toxins: structure and action. In: Hessinger D, Lenhoff H, eds. The Biology of Nematocysts. New York: Academic Press, 1988:375–405. 28. Alsen C. Biological significance of peptides from Anemonia sulcata. Fed Proc 1983;42:101–108. 29. Renaud J-F, Fosset M, Schweitz H, Lazdunski M. The interaction of polypeptide neurotoxins with tetrodotoxin-resistant Na+ channels in mammalian cardiac cells.

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Correlation with inotropic and arrhythmic effects. Eur J Pharmacol 1986; 120:161–170. 30. Schweitz H, Bidard J-N, Frelin C, Pauron D, Vijverberg HPM, Mahasneh DM, Lazdunski M, Vilbois F, Tsugita A. Purification, sequence and pharmacological properties of sea anemone toxins from Radianthus paumotensis. A new class of sea anemone toxins acting on the sodium channel. Biochemistry 1985; 24:3554–3561. 31. Frelin C, Vigne P, Schweitz H, Lazdunski M. The interaction of sea anemone and scorpion neurotoxins with tetrodotoxin-resistant Na+ channels in rat myoblasts. A comparison with Na+ channels in other excitable and non-excitable cells. Mol Pharmacol 1984;26:70–74. 32. Platou ES, Refsum H, Hotvedt R. Class III antiarrhythmic action linked with positive inotropy: antiarrhythmic, electrophysiological, and hemodynamic effects of the sea anemone polypeptide ATX II in the dog heart in situ. J Cardiovasc Pharmacol 1986; 8:459–465. 33. Vaughan Williams EM. Classification of antidysrhythmic drugs. Pharmac Therap B 1975; 1:115–138. 34. Sasayama S. What do the newer inotropic drugs have to offer? Cardiovasc Drugs Ther 1992; 6:15–18. 35. Catterall WA. Structure and function of voltage-sensitive ion channels. Science 1988; 242:50–61. 36. Wann KT. Neuronal sodium and potassium channels: structure and function. Br J Anaesthes 1993; 71:2–14. 37. Catterall WA. Structure and function of voltage-gated ion channels. Ann Rev Biochem 1995; 64:493–531. 38. Warashina A, Jiang Z-Y, Ogura T. Potential-dependent action of Anemonia sulcata toxins III and IV on sodium channels in crayfish giant axons. Eur J Physiol 1988; 411:88–93. 39. Warashina A, Ogura T, Fujita S. Binding properties of sea anemone toxins to sodium channels in the crayfish giant axon. Comp Biochem Physiol 1988; 90C:351–358. 40. Driscoll PC, Gronenborn AM, Beress L, Clore GM. Determination of the three-dimensional solution structure of the antihypertensive and antiviral protein BDS-1 from the sea anemone Anemonia sulcata: a study using nuclear magnetic resonance and hybrid distance geometry—dynamical simulated annealing. Biochemistry 1989; 28:2188–2198. 41. Llewellyn LE, Norton RS. Binding of the sea anemone polypeptide BDS II to the voltage-gated sodium channel. Biochem Intl 1991; 24:937–946.

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42. Catterall WA, Beress L. Sea anemone toxin and scorpion toxin share a common receptor site associated with the action potential sodium ionophore. J Biol Chem 1978; 253:7393–7396. 43. Scholtysik G. Cardiac Na+ channel activation as a positive inotropic principle. J Cardiovasc Pharmacol 1989; 14 (Suppl 3):S24–S29. 44. Scholtysik G, Quast U, Schaad A. Evidence for different receptor sites for the novel cardiotonic SDPI 201-106, ATX II, and veratridine at the cardiac sodium channel. Eur J Pharmacol 1986; 125:111–118. 45. Thomsen WJ, Catterall WA. Localization of the receptor site for α-scorpion toxins by antibody mapping: implications for sodium channel topology. Proc Natl Acad Sci USA 1989; 86:10161–10165.

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46. Pallaghy PK, Scanlon MJ, Monks SA, Norton RS. Three-dimensional structure in solution of the polypeptide cardiac stimulant anthopleurin-A. Biochemistry 1995; 34:3782–3794. 47. Monks SA, Pallaghy PK, Scanlon MJ, Norton RS. Solution structure of the cardiostimulant polypeptide anthopleurin-B and comparison with anthopleurin-A. Structure 1995; 3:791–803. 48. Widmer H, Billeter M, W¨thrich K. The three-dimensional structure of the neurotoxin ATX Ia from Anemonia sulcata in aqueous solution by nuclear magnetic resonance spectroscopy. Proteins 1989; 6:357–371. 49. Fogh RH, Kem WR, Norton RS. Solution structure of neurotoxin I from the sea anemone Stichodactyla helianthus. A nuclear magnetic resonance, distance geometry and restrained molecular dynamics study. J Biol Chem 1990; 265:13016–13028. 50. Wilcox GR, Fogh RH, Norton RS. Refined structure in solution of the sea anemone neurotoxin Sh I. J Biol Chem 1993; 268:24707–24719. 51. Kraulis P. MOLSCRIPT: a program to produce both detailed and schematic plots of protein structures. J Appl Crystallogr 1991; 24:946–950. 52. Scanlon MJ, Norton RS. Multiple conformations of the sea anemone polypeptide anthopleurin-A in solution. Protein Sci 1994; 3:1121–1124. 53. Khera PK, Benzinger GR, Lipkind G, Drum CL, Hanck DA, Blumenthal KM. Multiple cationic residues of anthopleurin-B that determine high affinity and channel isoform discrimination. Biochemistry 1995; 34:8533–8541. 54. Gould AR, Norton RS. Chemical modification of cationic groups in the polypeptide cardiac stimulant anthopleurin-A. Toxicon 1995; 33:187–199. 55. Pennington MW, Kem WR, Dunn BM. Synthesis and biological activity of six monosubstituted analogs of a sea anemone polypeptide neurotoxin. Peptide Res 1990; 3:228–232. 56. Gould AR, Mabbutt BC, Norton RS. Structure-function relationships in the polypeptide cardiac stimulant, anthopleurin-A. Effects of limited proteolysis by trypsin. Eur J Biochem 1990; 189:145–153. 57. Gooley PR, Blunt JW, Beress L, Norton RS. Effects of pH and temperature on cardioactive polypeptides from sea anemones: a 1H-NMR study. Biopolymers 1988; 27:1143–1157. 58. Sunahara S, Muramoto K, Tenma K, Kamiya H. Amino acid sequence of two sea anemone toxins from Anthopleura fuscoviridis. Toxicon 1987; 25:211–219. 59. Gallagher MJ, Blumenthal KM. Cloning and expression of wild-type and mutant forms of the cardiotonic polypeptide anthopleurin-B. J Biol Chem 1992; 267:13958–13963. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_318.html (1 of 2) [4/5/2004 5:24:15 PM]

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60. Pennington MW, Zadenberg I, Byrnes ME, Norton RS, Kem WR. Synthesis of the cardiac inotropic polypeptide anthopleurin-A. Intl J Pept Prot Res 1994; 43:463–470. 61. Gallagher MJ, Blumenthal KM. Importance of the unique cationic residues arginine-12 and lysine49 in the function of the cardiotonic polypeptide anthopleurin-B. J Biol Chem 1994; 269:254–259. 62. Khera PK, Blumenthal KM. Role of the cationic residues arginine-14 and lysine-48 in the function of the cardiotonic polypeptide anthopleurin-B. J Biol Chem 269:921–925.

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63. Lampe RA, Lo MMS, Keith RA, Horn MB, McLane MW, Herman JL, Spreen RC. Effects of sitespecific acetylation on ω-conotoxin GVIA binding and function. Biochemistry 1993; 32:3255–3260. 64. Basus VJ, Nadasdi L, Ramachandran J, Miljanich GP. Solution structure of ω-conotoxin MVIIA using 2D NMR spectroscopy. FEBS Lett 1995; 370:163–169. 65. Manoleras N, Norton RS. Three-dimensional structure in solution of neurotoxin III from the sea anemone Anemonia sulcata. Biochemistry 1994; 33:11051–11061. 66. Stanfield RN, Wilson IA. Protein-peptide interactions. Curr Opinion Str Biol 1995; 5:103–113. 67. Stampe P, Kolmakova-Partensky L, Miller C. Intimations of K+ channel structure from a complete functional map of the molecular surface of charybdotoxin. Biochemistry 1994; 33:443–450. 68. Goldstein SAN, Pheasant DJ, Miller DJ. The charybdotoxin receptor of a Shaker K+ channel: peptide and channel residues mediating molecular recognition. Neuron 1994; 12:1377–1388. 69. Kim JI, Takahashi M, Ogura A, Kohno T, Kudo Y, Sato K. Hydroxyl group of Tyr13 is essential for the activity of ω-conotoxin GVIA, a peptide toxin for N-type calcium channel. J Biol Chem 1994; 269:23876–23878. 70. Clackson T, Wells JA. A hot spot of binding energy in a hormone-receptor interface. Science 1995;267:383–386. 71. Jin L, Wells JA. Dissecting the energetics of an antibody-antigen interface by alanine shaving and molecular grafting. Protein Sci 1994; 3:2351–2357. 72. Marshall GR. A hierarchical approach to peptidomimetic design. Tetrahedron 1993; 49:3547–3558. 73. Chen S, Chrusciel RA, Nakanishi H, Raktabutr A, Johnson ME, Sato A, Wiener D, Hoxie J, Saragovi HU, Greene MI, Kahn M. Design and synthesis of a CD4 β-turn mimetic that inhibits human immunodeficiency virus envelope glycoprotein gp120 binding and infection of human lymphocytes. Proc Natl Acad Sci USA 1992; 89:5872–5876. 74. Jackson S, De Grado W, Dwivedi A, Parthasarathy A, Higley A, Krywko J, Rock-well, A, Markwalder J, Wells G, Wexler R, Mousa S, Harlow R. Template-constrained cyclic peptides: design of high-affinity ligands for GPIIb/IIIa. J Am Chem Soc 1994; 116:3220–3230. 75. Ohlstein EH, Nambi P, Douglas SA, Edwards RM, Gellai M, Lago A, Leber JD, Cousins RD, Gao A, Frazee JS, Peishoff CE, Bean JW, Eggleston DS, Elshourbagy NA, Kumar C, Lee JA, Yue T-L, Louden C, Brooks DP, Weinstock J, Feuerstein G, Poste G, Ruffolo RR, Gleason JG, Elliot JD. SB 209670, a rationally designed potent nonpeptide endothelin receptor antagonist. Proc Natl Acad Sci USA 1994; 91:8052–8056.

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76. Gould AR, Mabbutt BC, Llewellyn LE, Goss NH, Norton RS. Linear and cyclic peptide analogues of the polypeptide cardiac stimulant anthopleurin-A. 1H-NMR and biological activity studies. Eur J Biochem 1992; 206:641–651. 77. Stocker M, Miller C. Electrostatic distance geometry in a K+ channel vestibule. Proc Natl Acad Sci USA 1994; 91:9509–9513.

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13 Rational Design of Renin Inhibitors V. Dhanaraj* and J.B. Cooper†** Birkbeck College, London, England I. Introduction There has been much interest in the development of therapies for hypertension and associated heart failure, which is a major cause of death in the western world. One of the key mediators in primary hypertension is the plasma octapeptide angiotensin II (AII), which plays a major role by causing vasoconstriction and stimulating aldosterone release, thereby increasing blood volume by its action on the kidneys. Angiotensin II is produced by a proteolytic cascade—known as the renin-angiotensin system—in which the aspartic proteinase renin catalyses the rate-limiting cleavage of angiotensinogen produced by the liver to yield the decapeptide angiotensin I (AI). The subsequent removal of the carboxy-terminal dipeptide from AI by angiotensin-converting enzyme (ACE), yielding AII, is the target for a number of drugs that are effective for treating hypertension, hyperaldosteronism, and congestive heart failure [1]. The development of potent low-molecular-weight orally active ACE inhibitors from natural and synthetic metalloproteinase inhibitors has been rapid, due in part to the relative lack of specificity of this enzyme. In contrast, renin cleaves only its natural substrate or very close analogs and although inhibition of an enzyme more specific than ACE may be desirable for reducing side effects in vivo, the selectivity of renin meant that during the early stages of drug development, potent inhibition required the use of large peptide-based compounds. These were often poorly absorbed and susceptible to gastric proteolysis and biliary excretion. Nevertheless, the commercial and clinical success of ACE inhibitors fueled interest in the search for therapeutic renin drugs. Most inhibitors have been developed by elaboration of the minimal substrate sequence (residues 6–13 of angiotensinogen), which exhibits weak competitive inhibition, and replacement of the scissile bond with various nonhydrolysable surrogates, some of which may be transition state analogues [2]. Current affiliation: University of Cambridge, Cambridge, England. Current affiliation: University of Southhampton, Southhampton, England.

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Figure 1 The three-dimensional structures of human (left) and mouse renins (right) showing oligopeptide inhibitors bound in the active site cleft. The cleft lies between the N- and C-terminal domains of the enzyme and is approximately perpendicular to the plane of the page. It can accommodate 9–10 residues with the substrate/inhibitor bound in an extended conformation. The catalytic aspartic acid residues (not shown) are centrally placed at the base of the cleft.

Renin is a member of the homologous group of enzymes known as aspartic proteinases that includes pepsin and a group of fungal enzymes such as endothiapepsin, penicillopepsin, and rhizopuspepsin. Their sequences all contain two aspartates (at positions 32 and 215 in porcine pepsin) that are essential for catalytic activity. The crystal structures of several aspartic proteinases have been solved by x-ray diffraction at high resolution, revealing a common bilobal structure with a large cleft between the N- and C-terminal domains that can accommodate up to nine residues of a substrate (Figure 1) [3]. The two essential carboxyls of Asp 32 and Asp 215 are within hydrogen-bonding distance and are approximately co-planar due to the constraints of a hydrogen-bonding network involving residues of the two highly conserved loops that contain the essential aspartates. The three-dimensional structures of the two domains are related by a topological two-fold axis passing between the catalytic residues where the pseudosymmetry happens to be strongest. Modeling studies based on the homology with other aspartic proteinases showed that human renin assumes a tertiary structure that is similar to the other enzymes and that the homology is greatest for the binding cleft region [4]. Subsequent x-ray analysis of the structure of renin (described later) revealed the specific interactions made with inhibitors and implicated certain loop regions covering the active site as being important for tight binding of peptides. One enigmatic feature of renin is its extreme substrate specificity, its only known natural substrate being a single Leu-Val peptide bond of angiotensinogen. The minimal synthetic analog is the 6–13 octapeptide that encompasses the scissile peptide bond of angiotensinogen between residues 10 and 11 [5]. It has

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been suggested that specificity of proteinases in general is due to rigidity in the binding pockets [6]; broad specificity results from the ability of the pockets to change shape in response to different ligand side chains. However, renin is known to be inhibited by a wide variety of peptide analogs of different length and sequence indicating either that the active site may be somewhat flexible or that the strength of binding of a substrate does not determine the rate of subsequent turnover. This is emphasised by the existence of substrates that act as competitive inhibitors, e.g., RIP of Haber and Burton [7], which has a Ki that is lower than the Km. Therefore hydrolysis of bound substrate appears to be more specific than the binding step. This may be because only certain substrate sequences allow correct positioning of the scissile bond for hydrolysis. Evidence for this effect was provided by comparison of 21 inhibitor structures of endothiapepsin [8], where it was shown that for inhibitors with different sequences but with the same transition state analog, the scissile bond analog can be disposed somewhat differently with respect to the catalytic carboxyls in each case. The availability of crystal structures of a number of renin inhibitors complexed with fungal aspartic proteinases [8, 9] allowed new compounds to be designed and modeled by such techniques as computer graphics, energy minimization, and molecular dynamics [10]. X-ray crystallographic analysis of aspartic proteinase inhibitor complexes has made a significant contribution to rationalizing the activity data for many of these compounds as well as understanding the catalytic mechanism of this class of proteinase. II. Strategies for Design of Renin Inhibitors Some of the parameters that have been varied in the search for therapeutically active renin inhibitors are outlined below. A. Elaboration of the Transition State Analog There is no evidence that aspartic proteinase catalysis involves a covalently bound intermediate [11] and major advances in the design of nonhydrolysable analogs have stemmed from attempts to mimic an intermediate of the following form.

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This intermediate is derived by nucleophilic attack of a water molecule on the scissile-bond carbonyl. Szelke pioneered the use of reduced-bond analog (-CH2-NH-), which were incorporated into the 6–13 peptide of angiotensinogen [12]. Cocrystallisation with endothiapepsin revealed that the reduced-bond analog associates tightly with aspartate carboxyls (32 and 215) probably via a salt link. The naturally occurring transition-state analogue statine (-CHOH-CH2-CO-NH-) [9] is a closer analogue of the putative intermediate and has been incorporated into many inhibitors [13]. The scissile bond has also been replaced by the ketone analogue (-CO-CH2-) [12] or by a C-terminal aldehyde group in a series of tetrapeptides [14,15]. Although these appear to mimic the substrate more closely than the intermediate, the carbonyl probably binds to the enzyme in the hydrated gem-diol form (-C(OH)2-CH2) [16]. Use of the hydroxyethylene analog (-CHOH-CH2-) has led to exceptionally potent inhibitors [17] as has substitution of fluorines into ketone analogs [16,18] giving, for example, -CO-CF2- which undergoes hydration of the carbonyl to form -C(OH)2-CF2- and exhibits tight binding to renin. The hydrated gemdiol is thought to closely mimic the putative transition state -C(OH)2-NH-. All inhibitors solved in complex with aspartic proteinases by x-ray diffraction are observed to adopt similar main-chain conformations and form a conserved set of hydrogen bonds involving the inhibitor's main-chain groups interacting with enzyme moieties. The inhibitors bind in extended conformations and residues in the P3-P1 region form antiparallel β-sheet-like interactions with residues 217–219 on the enzyme. A β-hairpin turn formed by residues 74–78 lies between the inhibitor and bulk solvent and forms a number of generally conserved hydrogen bonds with the bound peptide. These interactions are indicated in Figure 2. The binding pockets for the inhibitor's side chains are shallow and contiguous with a greater hydrophobic character towards the central region of the active site cleft. The elaboration of several classes of transition state analog are now considered in greater detail. Statine Analogs The natural transition analog statine possesses one less main chain atom than a dipeptide and has been shown by x-ray analysis of inhibitor cocrystals to occupy the S1 and S1' sites of the enzyme [19]. The hydroxyl group of statine binds symmetrically between the catalytic carboxyl groups displacing a solvent molecule bound to the native enzyme. The carboxyl diad, therefore, provides a stereospecific binding site for statine and hydroxyethylene analogues with preference for the S-enantiomer. Replacement of the hydroxyl by an ammonium group might be expected to improve the potency of an inhibitor by introduction of a salt link with the enzyme. The corresponding deoxy-aminostatine (ASTA) analogs have been synthesised [20, 21] and were found to be nearly as potent as

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Figure 2 A schematic diagram of the putative hydrogen bonds formed between oligopeptide inhibitors and the fungal aspartic proteinase endothiapepsin. The latter enzyme provided a useful model system for structural studies of interactions formed by renin inhibitors with the active site cleft of aspartic proteinases prior to the determination of the human renin structure. The inhibitor is shown horizontally with enzyme groups above and below. Intervening hydrogen bonds are indicated by dashed lines. Note the extensive hydrogen-bond interactions made between the transition state analogs and the catalytic apparatus of the enzyme.

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the equivalent statine analogs with the benefit of improved solubility. Preference for the S- versus Renantiomer at the C3 amino position is observed in accordance with the statine inhibition data [22]. Oxahomostatine (-CHOH-CH2-O-CO-) and azahomostatine (-CHOH-CH2-NR-CO-) analogs eliminate the main-chain frameshift that occurs with statine and, in addition, the azahomostatine conveniently reduces the stereochemical complexity of the dipeptide surrogate by introducing a 7-membered urea-like planar group into the S1' binding region. The crystal structure of such a compound complexed with endothiapepsin has been solved at 1.8 Å resolution [23] and reveals that the large planar group is accommodated by the active site and that hydrogen bonds to the P1' CO and P2' NH groups, observed in other complexes, are retained. One shortened analog of statine -CHOH-CO-O-R referred to as norstatine (where R is a C-terminal alkyl group) has been found to be more potent than some equivalent statine analog [24]. The x-ray structure of such an inhibitor complexed with endothiapepsin reveals that the carbonyl oxygen of this analog is held by a hydrogen bond to the active site flap region of the enzyme involving the Gly76 >NH group (pepsin numbering) in much the same way as the P1' >C=O group of other isosteres (Figure 2). Aminoalcohols In principle, a good analog of the putative intermediate would be the aminal –CHOH–NH– group but this would be in equilibrium with the aldehyde and amino fragments. Interposing a methylene group between the hydroxymethyl and amino groups stabilizes the analog and may still allow tight binding to the enzyme. Such aminoalcohols (-CHOH-CH2-NH-) have been synthesized [25] and were shown to be potent inhibitors. Cocrystallisation of two such compounds extending from P1 to P3' with endothiapepsin allowed their bound structures to be solved at high resolution [26]. The bound structures revealed that despite the insertion of a methylene group in the analog a frameshift in the binding mode does not occur since the residue following the aminoalcohol occupies the S1' pocket. In contrast, the single amino acid, statine, replaces two residues of the substrate. The hydroxyl of the aminoalcohol (S-enantiomer) is bound symmetrically to both essential carboxyls as is the case for the hydroxyl of the statine and hydroxyethylene analogs. Glycols Incorporation of glycol or vicinal diol analogs of the peptide bond (-CH(OH)-CH(OH)-) has led to potent inhibitors and the x-ray structure for one such compound complexed with endothiapepsin is available [27]. The first hydroxyl in

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this analog interacts with the catalytic aspartate carboxyls in the same manner as statine or hydroxyethylene moieties; whereas, the second hydroxyl forms a hydrogen bond with the NH group of Gly 76, thereby mimicing the carbonyl oxygen at P1' of other analogs (see Figure 2). Phosphorus-Containing Analogs Aspartic proteinase inhibitors in which the scissile bond is replaced by a phosphinic acid group (shown below) have been reported [28].

These may mimic the tetrahedral intermediate more closely than statine or hydroxyethylene analogs. One of the oxygens binds to the carboxyl diad and the other resides adjacent to Tyr75 (pepsin numbering) forming a hydrogen bond with the outer oxygen of Asp32 [27]. This isostere is very effective against pepsin. However, it ionizes at physiological pH and the resulting anion is ineffective as an inhibitor of renin [29]. Fluoroketone Analogs and Implications for Catalysis Fluoroketone analogs (-CO-CF2-) have been reported [16, 30] and found to be substantially more potent than the unhalogenated statone molecules, presumably due to the ease of hydration and greater complementarity of the resulting hydrated gem-diol with the catalytic site. The structure of a difluorostatone inhibitor complexed with endothiapepsin [31] revealed interactions that indicate how the catalytic intermediate is stabilized by the enzyme (Figure 3). One hydroxyl of the hydrated fluoroketone associates tightly with the aspartate diad in the same position as the statine hydroxyl or the native solvent molecule and the other hydroxyl is positioned such that it hydrogen bonds to the outer carboxyl oxygen of Asp32. It has been suggested that the tetrahedral intermediate is uncharged, because if the carboxyl of Asp32 carries a negative charge instead, the latter can be stabilized by a full complement of hydrogen bonds donated by the gem-diol intermediate and surrounding protein atoms [31]. The current mechanistic proposals are based on the key suggestion by Suguna et al. [32] that, although transition state analogs appear to displace the active-site water molecule located between the two catalytic aspartate carboxyls, the more weakly bound substrate may not. Instead as the substrate binds, the water may be partly displaced to a position appropriate for nucleophilic attack on the scissile bond carbonyl. Details of the proposed mechanism are given in Figure 3.

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Figure 3 The catalytic mechanism proposed by Veerapandian et al. [31] based on the x-ray structure of a difluoroketone (geminal-diol) inhibitor bound to endothiapepsin. A water molecule tightly bound to the aspartates in the native enzyme is proposed to nucleophilically attack the scissile-bond carbonyl. The resulting geminal-diol intermediate is stabilised by hydrogen bonds with the negatively charged carboxyl of aspartate 32. Fission of the scissile C-N bond is accompanied by transfer of a proton from Asp215 to the leaving amino group.

B. Complementarity of the Inhibitor Optimizing the fit of a ligand to its binding site improves the potency by burying lipophilic residues and by maximizing the number of van der Waals contacts, hydrogen bonds, and charge—charge interactions. The principles that apply to ligand binding are similar to those involved in protein folding. Inhibitor

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binding involves displacement of hydrogen-bonded water molecules from both the ligand and the binding cleft. This process is entropy favored since waters in the solvent lattice are more disordered than those bound to protein. The change in enthalpy on forming NH…CO bonds in the complex from >CO…HOH and >NH…OH2 is favorable but relatively small [33]. The hydrophobic effect is therefore thought to play a dominant role in the energetics of binding with hydrogen bonds providing precise alignment of the ligand with respect to the catalytic apparatus. The main chain >CO and >NH groups from P3 to P3' of aspartic proteinase inhibitors are nearly always satisfied by hydrogen-bond interactions on formation of the complex. Therefore, given that polypeptides can form the same hydrogen bonds to the binding cleft regardless of amino acid sequence, differences in affinity for ligands of equal length must be due to other interactions at the specificity pockets, presumably those between the ligand's side chains and the enzyme. One example of optimizing these interactions for renin is the use of the cyclohexylmethyl side chain at P1, which has been shown to improve the potency by two orders of magnitude relative to the equivalent leucine-containing inhibitor [13]. Structure/Activity Relationship (SAR) studies have shown that in many inhibitor types, the cyclohexylmethyl group is optimal for the S1 pocket of human renin; whereas, other analogs such as cyclohexyl, cyclohexylethyl, and the very bulky dicyclohexyl and adamantyl rings generally have significantly reduced potency [10]. The use of a cyclohexylmethyl appears to introduce selectivity for renin versus other human aspartic proteinases. This has been partly rationalized for endothiapepsin where it was shown by x-ray analysis that the cyclohexylmethyl group at P1 can force the Phe at P3 to adopt a less energetically favorable X2 angle. Hence, differences at the S3 pocket in renin may allow the P3 Phe to adopt a more favorable X2 angle in the presence of a cyclohexyl at P1. In contrast the S2 site is able to accommodate a wide variety of side chains depending on inhibitor type, e.g., Phe and His are equipotent in some analogs [34]. The x-ray structures of a number of bound renin inhibitors complexed with endothiapepsin have shown that His at P2 can adopt different X1 angles separated by about 120 degrees [8]. In one conformation the imidazole is lying partly in the S1' pocket, which has a definite hydrophobic character. In the other conformation, the His side chain is in a more polar environment. The ability of aspartic proteinases to accept a variety of both polar and hydrophobic groups at the P2 position may be due to this bifurcation. Many inhibitors possess naphthylalanine side chains at P3 and P4 [14,24,35]. Compounds of this type are potent renin inhibitors with binding constants in the nanomolar range. Cocrystallisation of such an inhibitor with endothiapepsin revealed that one naphthalic ring is accommodated in the S3 pocket by significant conformational changes of local enzyme side chains (Asp77 and Asp114). The other naphthalene lies in the S4 binding region [36].

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C. Rigidification The number of conformations that a peptide can adopt in solution is reduced by cyclization. This can be optimized, at least in theory, to lock the peptide in the conformation that has the highest affinity for the receptor, resulting in a gain of affinity, primarily for entropic reasons. The structures of the fungal aspartic proteinases reveal that the binding cleft is a wide channel with no obvious division between the pockets, e.g., S1 and S3 are contiguous. The bound structures of numerous inhibitors have shown that alternate side chains are in van der Waals contact due to the extended conformation that these ligands adopt. In addition, at certain positions, e.g., P2, the side chains are allowed very different conformations due to the permissiveness of the pocket. Hence, the cross-linking of certain side chains may, at least, not be detrimental to inhibitory potency and may also reduce the susceptibility to degradation in the gut or plasma. It might therefore be expected that oligopeptide renin inhibitors would be suitable' candidates for cross-linking experiments. A similar philosophy of rigidification was pursued in the development of the ACE inhibitor cilazapril [37]. A number of statine-containing inhibitors possessing disulphide links between P2 and P5, and P2 and P4' have been synthesized [13] although the best potencies were slightly less than for the linear peptides. An alkyl cycle of varying length was introduced between the hydroxyl of a serine residue at P1 and the main chain nitrogen of P2 in a series of reduced-bond inhibitors [53]. Potencies similar to the uncrosslinked molecule were achieved but none were greater. This was attributed to the cis isomerisation of the P3—P2 peptide bond giving a conformation that cannot fit the active site of the enzyme. Difficulties in achieving more potent cyclic inhibitors may be due to the tight binding environment provided by some pockets (especially S1 and S3), and the possibility that other unproductive conformations of the inhibitor become favorable. More recently similar findings have been reported for cyclic analogs of pepstatincontaining alkyl crosslinks of variable length between the P1 and P3 side chains [38]. D. In Vivo Stability Peptides, when administered orally, are susceptible to degradation in the stomach by gastric enzymes and the proteinases of the pancreas and brush border of the small intestine. Their lifetimes in the plasma are often short due to rapid proteolysis and other metabolic processes. Early efforts were made to improve the resistance of renin inhibitors to hydrolysis in vivo by the use of blocking groups at the Nand C-terminii [39] and replacement of susceptible peptide bonds other than the renin cleavage site. Studies of SAR have shown that various N- and C-terminal groups, some based on the morpholine nucleus and derivatives of it, have a favorable effect on the duration of inhibition in the

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plasma. This may arise from reduced nonspecific plasma binding due to the relatively polar nature of these blocking groups [24]. Resistance of inhibitors to gut proteolysis has been improved by various methods such as replacement of phenylalanine at P3 with O-methyl tyrosine (or naphthylalanine), which was shown to abolish chymotrypsin cleavage and yet retain high inhibitory potency for renin [40]. III. Structural Studies of Rennin Complexed With Inhibitors The three-dimensional structures of renin-inhibitor complexes had long been sought as an aid to the discovery of clinically effective antihypertensives [41]. X-ray analyses of recombinant human renin [42] and mouse submandibulary renin [43] have given an accurate picture of active-site interactions and largely confirm the predictions of models based on homologous aspartic proteinases [4]. A large number of questions concerning the specificities of renins have been answered by these x-ray analyses. The renin-inhibitor structures also make an important contribution towards the rational design of effective antihypertensive agents. A. X-Ray Analysis of Mouse and Human Renin Complexes For both of these renins multiple copies of the molecules have been independently defined in the x-ray analysis and shown to have very similar structures. These x-ray structures were refined to final agreement factors and correlation coefficients of 0.19 and 0.91 for human renin at 2.8 Å resolution and 0.18 and 0.95 for mouse renin at 1.9 Å resolution. As expected from the high degree of sequence identity of human and mouse renins (approximately 70%), they have very similar three-dimensional structures as shown in Figure 1. The active-site cleft has a less open arrangement in renins than in the other aspartic proteinases. Many loops as well as the helix hc (residues 224–236) belonging to the C-domain (residues 190–302) are significantly closer to the active site in the renin structures compared to those of endothiapepsininhibitor complexes. This is partly due to a difference in relative position of the rigid body comprising the C-domain. For instance, there is a domain rotation of ~4° and translation of ~0.1 Å in the human renin complex with respect to the endothiapepsin-difluorostatone complex. The entrance to the active-site cleft is made even narrower in renins as a consequence of differences in the positions and composition of several well-defined loops and secondary structure elements. Unique to the renins is a cis proline, Pro111, which caps a helix (hN2) and contributes to the subsites S3 and

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S5. This helix is nearer to the active site in renins than in other aspartic proteinases. On an equivalent loop in the C-lobe (related by the intramolecular pseudo 2-fold axis), there is a sequence of three prolines—the Pro292–Pro293– Pro 294 segment. This structure is also unique to the renins among the aspartic proteinases with Pro294 and Pro297 in a cis configuration. Such a proline-rich structure provides an effective means of constructing well-defined pockets from loops that would otherwise be more flexible. This rather rigid poly-proline loop, together with the loop comprised of residues 241–250, lies on either side of the active site “flap” formed by residues 72–81. Hence, in the renins, the cleft is covered by the flaps from both lobes rather than from the N-lobe alone as in other pepsin-like aspartic proteinases. This gives renin a superficial similarity to the dimeric, retroviral proteinases where each subunit provides an equivalent flap that closes down on top of the inhibitor [44,45]. B. The Role of Hydrogen Bonds in Inhibitor Recognition Whereas the mouse renin inhibitor extends from P6 to P4', the human renin inhibitor extends only from P4 to P1'. The cyclohexyl norstatine residue at P1 in the human renin inhibitor mimics a dipeptide analog with its isopropyloxy group occupying the subsite for the side chain of P1'. The mouse renin inhibitor (CH-66) possesses a Leu-Leu hydroxyethylene transition state analog [12]. Both inhibitors are bound in the extended conformation that is found in other aspartic proteinase-inhibitor complexes. Both inhibitors make extensive hydrogen bonds with the enzymes as shown in Figure 4. In general the two renininhibitor complexes described here demonstrate that a similar pattern of hydrogen bonding is probably used in the substrate recognition of all aspartic proteinases although their specificities differ substantially. There is also great similarity between aspartic proteinases in terms of interactions with the transitionstate analog inhibitors at the catalytic center. The catalytic aspartyl side chains and the inhibitor hydroxyl group are essentially superimposable in both renin complexes. The isostere C-OH bonds lie at identical positions when the structures of inhibitor complexes of several aspartic proteinases are superposed, in spite of the differences in the sequence and secondary structure. Most of the complex array of hydrogen bonds found in endothiapepsin complexes are formed in renin with the exception of that to the threonine or serine at 218, which is replaced by alanine in human renin. The similarity can be extended to all other pepsin-like aspartic proteinases and even to the retroviral proteinases [44,45]. This implies that the recognition of the transition state is conserved in evolution, and the mechanisms of this divergent group of proteinases must be very similar.

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Figure 4 The inhibitors complexed with human (top) and mouse (bottom) renin showing the putative hydrogen-bond interactions made with the enzyme moieties.

C. Specificity If the main-chain hydrogen bonding of substrates is conserved among aspartic proteinases, how are the differences in specificities achieved? Table 1 defines the enzyme residues that line the specificity pockets for both mouse and human renin. In modeling exercises (e.g., Reference 4) it was assumed that specificities derive from differences in the sizes of the residues in the specificity pockets (Sn) and their ability to complement the corresponding side chains at positions Pn in the substrate/inhibitor. A detailed analysis now shows that this simple assumption only partly accounts for the steric basis of specificity.

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For example, in the specificity subsite S3 the phenyl rings of Phe P3 occupy almost identical positions in both renin inhibitor complexes. Modeling studies have predicted the specificity subsite S3 to be larger in renins than in other aspartic proteinases [4] due to substitution of smaller residues, Pro 111, Leu114, and Ala115, in place of larger ones in mammalian and fungal proteinases. However, a compensatory movement of a helix (hN2) makes the pocket quite compact and complementary to the aromatic ring as shown in Figure 5. Thus, the positions of an element of secondary structure differ between renin and other aspartic proteinases with a consequent important difference in the specificity pocket. The differing positions of secondary structural elements may also account for the specificities at P2'. Mouse submaxillary and other nonprimate renins do not appreciably cleave human angiotensinogen or its analogs [46], which have an isoleucine at P2', although they do cleave substrates with a valine at this position. In contrast, human renin not only cleaves the human and nonprimate substrates but also the rat angiotensinogen with tyrosine at P2', albeit rather slowly [47]. This can be explained in terms of the threedimensional structures. In the mouse renin complex, the P2' tyrosyl ring is packed parallel to an adjoining helix (h3) in a narrow pocket and there is only limited space available beyond the Cβ methylene group. This appears to be able to accommodate a valine, but not the larger isoleucine at P2', which will suffer greater steric interference from several residues that are conserved in identity and position in the two renins. On the other hand, in human renin differences in the orientation and position of

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Figure 5 The S3 specificity pocket of human renin occupied by phenylalanine in the cyclohexylnorstatine inhibitor.

helix h3 bring it closer by (by ˜0.5Å) to the substrate-binding site than in mouse renin. It is orientated in such a fashion in human renin that, although it can accommodate the isobutyl side chain of isoleucine at P2', aromatic rings on substituents such as phenylalanine and tyrosine will have severe short contacts with the side chain of Ile130 (valine in mouse renin). Thus the reorientation of a helix, coupled with subtle differences in the shapes of the side chains, makes significant changes in the substrate specificity at this subsite. It is interesting to note that in pepsin this helix is in a similar position with respect to the active site as in human renin. This provides a structural rationale for the negative influence of peptides containing phenylalanine [48], tyrosine, or histidine [49] at this subsite (S2') on the rate of proteolytic pepsin cleavage, while isoleucine and valine enhance catalysis. Differences in the specificity subsites at S1' in the human and mouse renins have a more complicated explanation. At first sight the situation appears to be explained by complementarity of the subsites to the valine and leucine at http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_335.html (1 of 2) [4/5/2004 5:26:14 PM]

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P1' in human and mouse angiotensinogens. Accordingly residue 213 is leucine in human renin and valine in mouse renin. The S1' pockets of chymosin, pepsin, and endothiapepsin have an aromatic side chain at residue 189 while the renins have amino acids with smaller side chains (valine in human and serine in mouse renins). This would be expected to make the pocket larger in renins. However the structure of the mouse renin complex shows that the substrate moves closer to the enzyme in renins as a result of the smaller residue at 189 and the pocket is made even more compact due to a compensatory change in the position and composition of the polyproline loop (residues 290–297). Thus, the specificity difference at this site arises not only from a compensatory movement of a secondary structure, in this case a loop region, but also from the substitution of an enzyme residue that allows the substrate to come closer to the body of the enzyme. Elaboration of loops on the periphery of the binding cleft in renins also influences the specificity. This is most marked at P3' and P4', for which it has been particularly difficult to obtain complexes with welldefined conformations for other aspartic proteinases. In endothiapepsin, which has been the subject of the greatest number of studies, different conformations are adopted at P3' and the residue at P4' is generally disordered. In contrast these residues are clearly defined in mouse renin. This is mainly a consequence of the polyproline loop, illustrated in Figure 6, which occurs uniquely in renins. The x-ray analysis of the mouse renin complex shows that the S3' and S4' subsites are formed by the polyproline loop together with residues of the flap, and a similar situation is likely to occur in human renin. The welldefined interactions of P3' described in the mouse renin complex explains the significant affinity when inhibitors have phenylalanine or tyrosine at P3' as well as the importance of a P3' residue for catalytic cleavage of a substrate by renin [50]. Hydrogen bonds between the side chains of the inhibitor and the enzyme do not play a major role in most specificity pockets. However, S2 is an exception. This subsite is large and contiguous with S1', so that in human renin the S-methyl cysteine (SMC) side chain of P2 is oriented towards the S1' pocket, which is only partly filled by the isopropyloxy group of the putative P1' residue. The carbonyl oxygen of P2 accepts a hydrogen from the Oγ of Ser76, which is unique to human renin; residue 76 is a highly conserved glycine in all the other aspartic proteinases, including mouse renin. In mouse renin the P2 histidyl group has a different orientation and forms a hydrogen bond with the Oγ of Ser222. If such a conformation were adopted by the human angiotensinogen in complex with human renin, the two imidazole nitrogens would be hydrogen bonded to the Oγ of both Ser76 and Ser222. The observed reduction in the rate of cleavage of a human angiotensinogen analog containing a 3-methyl histidine substituent at P2 [51] could be explained on the basis of the hydrogen bonding scheme proposed above.

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Figure 6 The P3' tyrosine residue of the mouse renin inhibitor complex showing the unique polyproline loop on the right. Specificity of this and neighboring subsites in renins must derive partly from this rigid loop region.

IV. Rational Drug Design The pioneering work of Burton, Szelke, and others in developing peptide-based renin inhibitors has been followed by a worldwide commercial effort to elaborate such compounds into therapeutically active antihypertensives. The twin problems of insufficient oral bioavailability and rapid clearance has seemingly presented major obstacles to success. In addition, the possible advantages of renin inhibitors compared with ACE inhibitors remain questionable. Never-theless information from human-renin crystallographic studies—such as the more recent high resolution analyses [52] and algorithms for analysing voids in the complexes as potential sites for elaborating the drug molecule (e.g. Figure 7)—may yet provide leads for compounds with suitable therapeutic characteristics. The detailed analyses of renin-inhibitor complexes reported here confirm the general structural features thought to contribute to renin's specificity but

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Figure 7 Schematic illustration of the voids between enzyme and inhibitor in the crystal structure of human renin complexed with a norstatine inhibitor. The figure was produced using the GapE software (Dr. Roman Laskowski). The inhibitor (dark bonds) is enclosed by a net surface and the gaps (where the enzyme and inhibitor are not in contact) are represented by solid surfaces [42].

demonstrate the need for careful, high-resolution x-ray analyses for more confidence in drug design. In particular, they show that even minor alternations in the positions of secondary structural elements can lead to major changes in the disposition of the subsites and thus the recognition of substrates. Since such molecular recognition defines the species specificity and determines the catalytic efficiency of the enzymes, a through understanding is indispensable for the synthesis of suitable inhibitors. The specificity pockets—the molecular recognition sites—are modified by elaboration, particularly of surface loops, which can be disordered in the uncomplexed enzymes and difficult to model with precision from homologous structures. These data establish a new foundation for the rational design of renin inhibitors and have provided a rational base for development of clinically successful HIV proteinase inhibitors. References 1. Ondetti MA, Cushman DW. Enzymes of the renin-angiotensin system and their inhibitors. Annu Rev Biochem 1982; 51:283–308.

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2. Blundell TL, Cooper J, Foundling SI, Jones DM, Atrash B, Szelke M. On the rational design of renin inhibitors: X-ray studies of aspartic proteinases complexed with transition state analogues. Biochemistry 1987; 26:5585–5590.

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3. Pearl LH, Blundell TL. The active sites of aspartic proteinases. FEBS Lett 1984; 174:96–101. 4. Sibanda BL, Blundell TL, Hobart PM, Fogliano M, Bindra JS, Dominy BW, Chirgwin JM. Computer graphics modeling of human renin. FEBS Lett 1984; 174:102–111. 5. Skeggs LT, Dover FE, Levine M, Lentz KE, Kahn JR. In: Johnson JA, Anderson RR, ed. The ReninAngiotensin System. New York: Plenum. 6. Bone R, Silen JL, Agard DA. Structural plasticity broadens the specificity of an engineered protease. Nature 1989; 339:191–195. 7. Haber E, Burton J. Inhibitors of renin and their utility in physiologic studies. Fedn Proc 1979; 38:2768–2773. 8. Cooper JB, Bailey D. A structural comparison of 21 inhibitor complexes of the aspartic proteinase from Endothia parasitica. Protein Science 1994, 3:2129–2143. 9. Foundling SI, Cooper, J, Watson FE, Cleasby A, Pearl LH, Sibanda BL, Hemmings A, Wood SP, Blundell TL, Valler MJ, Norey CG, Kay J, Boger J, Dunn BM, Leckie BJ, Jones DM, Atrash B, Hallett A, Szelke M. High resolution X-ray analyses of renin inhibitor-aspartic proteinase complexes. Nature (London) 1987; 327:349–352. 10. Luly JR, Bolis G, Bamaung N, Soderquist J, Dellaria JF, Stein H, Cohen J, Thomas JP, Greer J, Plattner JJ. New inhibitors of human renin that contain novel replacements. Examination of the P1 site. J Med Chem 1988; 31:532–539. 11. Hofmann T, Fink AL. Cryoenzymology of penicillopepsin. Biochemistry 1984; 23:5249–5256. 12. Szelke M, Leckie B, Hallett A, Jones DM, Sueiras-Diaz J, Atrash B, Lever AF. Potent new inhibitors of human renin. Nature 1982; 299:555–557. 13. Boger J. Renin inhibitors. Design of angiotensin transition state analogues containing statine. In: Kostka V. ed. Aspartic Proteinases and Their Inhibitors. Berlin: Walter de Gruyter, 1985:401–420. 14. Kokubu T, Hiwada K, Murakami E, Imamura Y, Matsueda R, Yabe Y, Koike H, Iijima Y. Highly potent and specific inhibitors of human renin. Hypertension 1985; 7 (suppl.1):8–11. 15. Kokubu T, Hiwada K, Nagae A, Murakami E, Morisawa Y, Yabe Y, Koike H, Iijima Y. Statine containing dipeptide and tripeptide inhibitors of human renin. Hypertension (Suppl.II) 1986; 8:1–5. 16. Gelb MH, Svaren JP, Abeles RH. Fluoroketone inhibitors of hydrolytic enzymes. Biochemistry 1985; 24:1813–1817.

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17. Szelke M. Chemistry of renin inhibitors. In: Kostka V, ed. Aspartic Proteinases and Their Inhibitors. Berlin: Walter de Gruyter, 1985:421–441. 18. Sham HL, Stein HH, Rempel CA, Cohen J and Plattner JJ. Highly potent and specific inhibitors of human renin. FEBS Lett 1987; 220:299–301. 19. Cooper JB, Foundling SI, Blundell TL, Boger J, Jupp R, Kay J. X-ray studies of aspartic proteinasestatine inhibitor complexes. Biochemistry 1989; 28:8596–8603. 20. Arrowsmith RJ, Carter K, Dann JG, Davies DE, Harris CJ, Morton JA, Lister P, Robinson JA, Williams DJ. Novel renin inhibitors: synthesis of aminostatine and comparison with statine-containing analogues. J Chem Soc Chem Commun 1986; 10:755–757. 21. Jones M, Sueiras-Diaz J, Szelke M, Leckie B, Beattie S. Renin inhibitors containing the novel amino-acid 3aminodeoxystatine. In: Deber CM, Hruby VJ, Kopple

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KD eds. Peptides: Structure and Function. Rockford:Pierce Chemical Company, 1985:759–762. 22. Rich DH, Sun ETO, Ulm E. Synthesis of analogues of the carboxyl proteinase inhibitor pepstatin. Effect of structure on the inhibition of pepsin and renin. J Med Chem 1980; 23:27–33. 23. Sali A, Veerapandian B, Cooper JB, Founding SI, Hoover DJ, Blundell TL. High resolution X-ray diffraction study of the complex between endothiapepsin and an oligopeptide inhibitor: the analysis of inhibitor binding and description of the rigid body shifts in the enzyme. EMBO J 1989; 8:2179–2188. 24. Iizuka K, Kamijo T, Kubota T, Akahane K, Umeyama H, Kiso Y. New human renin inhibitors containing an unnatural amino acid, norstatine. J Med Chem 1988; 31:701–704. 25. Dann JG, Stammers DK, Harris, CJ, Arrowsmith RJ, Davies DE, Hardy GW, Morton JA. Human renin: an new class of inhibitors. Biochem Biophys Res Commun 1986; 134:71–77. 26. Cooper JB, Foundling SI, Blundell TL, Arrowsmith RJ, Harris CJ, Champness JN. A rational approach to the design of antihypertensives: X-ray studies of complexes between aspartic proteinases and aminoalcohol inhibitors. In: Leeming PR, ed. Topics in Medicinal Chemistry. London: Royal Society of Chemistry, 1988; 308–313. 27. Lunney EA, Hamilton HW, Hodges JC, Kaltenbrohn JS, Repine JT, Badasso M, Cooper J, Dealwis C, Wallace B, Lowther WT, Dunn BM, Humblet C. Analyses of ligand binding in five endothiapepsin crystal complexes and their use in the design and evaluation of novel renin inhibitors. J Med Chem 1993; 36:3809–3820. 28. Bartlett PA, Kezer WB. Phosphinic acid dipeptide analogues: potent, slowbinding inhibitors of aspartic proteinases. J Amer Chem Soc 1984; 106:4282–4283. 29. Greenlee WJ. Renin inhibitors. Pharm Res 1987; 4(5):364–374. 30. Thaisrivongs S, Pals DT, Harris DW, Kati WM, Turner SR. Design and synthesis of potent and specific renin inhibitors containing difluorostatine, difluorostatone and related analogues. J Med Chem 1986; 29:2088–2093. 31. Veerapandian B, Cooper JB, Sali A, Blundell TL. Direct observation by X-ray analysis of the tetrahedral “intermediate” of aspartic proteinases. Protein Science 1992; 1:322–328. 32. Suguna K, Padlan EA, Smith CW, Carlson WD, Davies DR. Binding of a reduced peptide inhibitor to the aspartic proteinase from Rhizopus chinensis: implications for a mechanism of action. Proc Natl Acad Sci USA 1987; 84:7009–7013. 33. Ptitsyn OB. Pure Appl Chem 1973; 31:227–244.

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34. Rosenberg SH, Plattner JJ, Woods KW, Stein HH, Marcotte PA, Cohen J, Perun TJ. Novel renin inhibitors containing analogues of statine retro-inverted at the C-termini: specificity of the P2 histidine site. J Med Chem 1987; 30:1224–1228. 35. Luly JR, Yi N, Soderquist J, Stein H, Cohen J, Perun TJ, Plattner JJ. New inhibitors of human renin that contain novel Leu-Val replacements. J Med Chem 1987; 30:1609–1616. 36. Cooper J, Quail W, Frazao C, Foundling SI, Blundell TL. X-ray crystallographic analysis of inhibition of endothiapepsin by cyclohexyl renin inhibitors. Biochemistry 1992; 31:8142–8150.

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37. Attwood MR, Hassall CH, Krohn A, Lawton G, Redshaw S. The design and synthesis of angiotensin converting enzyme inhibitor cilazapril and related bicyclic compounds. J Chem Soc Perkin 1986; I:1011–1019. 38. Szewczuk Z, Rebholz KL, Rich DH. Synthesis and biological activity of new conformationally restricted analogues of pepstatin. Int J Pept Res 1992; 40:233–242. 39. Wood JM, Fuhrer W, Buhlmayer P, Riniker B, Hofbauer KG. Protection groups increase the in vivo stability of a statine-containing renin inhibitor. In: Kostka V, ed. Aspartic Proteinases and Their Inhibitors. Berlin: Walter de Gruyter, 1985:463–466. 40. Bolis G, Fung AKL, Greer J, Kleinert HD, Marcotte PA, Perun TJ, Plattner JJ, Stein HH. Renin inhibitors. Dipeptide analogues of angiotensinogen incorporating transition-state, nonpeptidic replacements at the scissile bond. J Med Chem 1987; 30:1729–1737. 41. Greenlee, WJ Renin inhibitors. Med Res Rev 1990; 10:173. 42. Dhanaraj V, Dealwis C, Frazao C, Badasso M, Sibanda BL, Tickle IJ, Cooper JB, Driessen HPC, Newman M, Aguilar C, Wood SP, Blundell TL, Hobart PM, Geoghegan KF, Ammirati MJ, Danley DE, O'Connor BA, Hoover DJ. X-ray analyses of peptide-inhibitor complexes define the structural basis of specificity for human and mouse renins. Nature 1992; 357:466–472. 43. Dealwis CG, Frazao C, Badasso M, Cooper JB, Tickle IJ, Driessen H, Blundell TL, Murakami K, Miyazaki H, Sueiras-Diaz J, Jones DM, Szelke M. X-ray analysis at 2.0 Å resolution of mouse submaxillary renin complexed with a decapeptide inhibitor CH-66, based on the 4–16 fragment of rat angiotensinogen. J Mol Biol 1994; 236:342–360. 44. Wlodawer A, Miller M, Jaskolski M, Sathyanarayana BK, Baldwin E, Weber IT, Selk LM, Clawson L, Schneider J, Kent S. Conserved folding in retroviral proteinases: crystal structure of synthetic HIV-1 proteinase. Science 1989; 245:616–621. 45. Lapatto R, Blundell TL, Hemmings A, Overington J, Wilderspin A, Wood SP, Merson JR, Whittle PJ, Danley DE, Geoghegan KF, Hawrylik SJ, Lee SE, Scheld KG, Hobart PM. X-ray analysis of HIV-1 proteinase at 2.7 Å resolution confirms structural homology among retroviral enzymes. Nature (Lond) 1989; 342:299–302. 46. Poe M, Wu JK, Lin TY, Hoogsteen K, Bull HG, Slater EE. Renin cleavage of a human-kidney renin substrate analogous to human angiotensinogen that is human renin specific and resistant to cathepsin D. Analyt Biochem 1984; 140:459–467. 47. Cumin F, Lenguyen D, Castro B, Menard J, Corvol P. Comparative enzymatic studies of human renin acting on pure natural or synthetic substrates. Biochim Biophys Acta 1987; 913:10–19.

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48. Powers JC, Harley AD, Myers DV. Subsite specificity of porcine pepsin. In: Tang J, ed. Acid Proteases-Structure, Function and Biology. New York: Plenum Press, 1977:141–157. 49. Antonov VK. In: Tang J, ed. Acid Proteases-Structure, Function and Biology. New York: Plenum Press, 1977:179. 50. Skeggs LT, Lentz KE, Kahn JR, Hochstrasser H. Kinetics of the reaction of renin with nine synthetic peptide substrates. J. Exp Med 1968; 120:130–34. 51. Holzman TF, Chung CC, Edalji R, Egan DA, Martin M, Gubbins EJ. Krafft GA, Wang GT, Thomas AM, Rosenberg SH, Hutchins C. Characterisation of recombi-

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nant human renin kinetics, pH-stability, and peptidomimetic inhibitor binding. J Protein Chem 1991; 10:553–563. 52. Tong L, Pav S, Lamarre D, Pilote L, Laplante S, Anderson PC, Jung G. High resolution crystalstructures of recombinant human renin in complex with polyhydroxymonoamide inhibitors. J Mol Biol 1995; 250:211–222. 53. Sham HL, Bolis G, Stein HH, Fesik SW, Marcott PA, Plattner JJ, Rempel CA, Greer J. Renin inhibitors. Design and synthesis of a new class of conformationally restricted analogues of angiotensinogen. J Med Chem 1988; 31:284–295.

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14 Structural Aspects in the Inhibitor Design of Catechol O-Methyltransferase Jukka Vidgren and Martti Ovaska Orion Corporation Orion Pharma, Espoo, Finland I. Introduction Catechol O-methyltransferase (COMT) plays an important role in the catabolic inactivation of catecholamines. It is present both in extracerebral tissues and in the central nervous system. During the last few years there has been a remarkable interest in COMT. Basic biochemical and molecular biology research has given detailed insights into the function and nature of the enzyme. The knowledge of the crystallographic structure has allowed researchers to analyze the molecular mechanism of the catalytic reaction and to accomplish the structure-based design of inhibitors. The development of potent and selective inhibitors has provided effective pharmacological tools to investigate the physiological role of the enzyme. The main clinical interest has been the possible application of COMT inhibitors as adjuncts in the L-dopa therapy of Parkinson's disease. Parkinson's disease is a dopamine deficiency disorder. The dopamine-producing neurons in striatum are destroyed. The medication strategy is to replenish the missing dopamine. L-Dopa, given together with a peripheral inhibitor of dopa decarboxylase (DDC), for example, carbidopa, is a standard therapy in Parkinson's disease. While dopamine does not penetrate into the brain, L-dopa penetrates the blood-brain barrier and is decarboxylated into dopamine in the brain. The half-life of L-dopa is short and in the presence of DDC inhibitor a large amount of the drug is eliminated by COMT. The COMT enzyme produces the metabolite 3-methoxytyrosine (3-OMD), which has no benefit in the treatment of Parkinson's disease, but has a long elimination half-life and may be harmful during chronical treatment. Also a gradual loss of the efficacy of L-dopa occurs during longterm medication. Since the early 1980s active

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Figure 1 The rationale of COMT inhibition as adjunct in the L-dopa therapy of Parkinson's disease (reproduced by permission from COMT News, Issue 1, Orion Corporation, Orion Pharma, 1994).

research in pharmaceutical companies has been carried out to develop new potent, selective, and orally active COMT inhibitors. Some of them (e.g., entacapone), are now in final clinical trials and the results have been promising. The rationale of COMT inhibition can be seen in Figure 1. It can be concluded that COMT inhibition in peripheral tissues improves the brain entry of L-dopa and decreases the formation of 3-OMD. The dose of L-dopa can be lowered and the dose interval prolonged. Also a decrease of the fluctuations of dopamine formation has been observed. The inhibition of COMT seems to be the next step in improving the L-dopa therapy of Parkinson's disease. This paper discusses the structure-based approach for the understanding of the enzyme function and inhibitor design. II. The Enzyme A. Physiological Role of COMT Catechol O-methyltransferase (COMT, EC 2.1.1.6) was originally detected in rat liver extracts [1]. Since then, COMT has been found in many species: http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_344.html (1 of 2) [4/5/2004 5:27:34 PM]

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Figure 2 The reaction catalyzed by catechol O-methyltransferase. Dopamine: R=CH2-CH2-NH2; L-dopa: R=CH2-CH(NH2)-COOH.

animals, plants, and procaryotes [2]. In mammals the highest COMT activities have been found in the liver and kidney, but COMT is common in almost all mammalian tissues [2–4]. The COMT enzyme catalyzes the transfer of the methyl group from the coenzyme S-adenosyl-Lmethionine (AdoMet) to one of the phenolic hydroxyl groups of a catechol or substituted catechol [1] (Figure 2). The presence of magnesium ions is required for the catalysis. The reaction products are Omethylated catechol and S-adenosyl-L-homocysteine (AdoHcy). Physiological substrates of COMT are catecholamine neurotransmitters, dopamine, noradrenaline, and adrenaline, and some of their metabolites. The COMT enzyme inactivates catecholic steroids such as 2-hydroxyestradiol, drugs with a catechol structure such as L-dopa, and a large number of other catechol compounds [1,2,5–7]. The general physiological function of COMT is the inactivation of biologically active or toxic catechols. A schematic view of the major catecholamine pathways in the brain is shown in Figure 3. L-Dopa is the dopamine precursor used in the treatment of Parkinson's disease [8]. B. Primary Structures There are no isoenzymes of COMT known in different mammalian tissues. Two distinct forms of COMT have been found: one is soluble (S-COMT) and the other membrane bound (MB-COMT) [9,10]. Both soluble and membrane-bound COMT have been cloned and characterized [11–16]. The soluble and membrane-bound COMT are coded by one gene using two separate promoters [17]. The soluble COMT contains 221 amino acids, whereas the membrane-bound form has a 50-(human) or 43-(rat) residueslong amino-terminal extension containing the hydrophobic membrane anchor region. The sequences of COMT enzymes from different species are highly similar (see Figure 4). The soluble human protein is 81% identical with the rat enzyme. The 165-amino-acids-long fragment of porcine COMT has 82% homology with the human

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Figure 3 The main metabolic routes of dopamine and noradrenaline in the brain. COMT, Catechol O-methyltransferase; MAO, monoamino oxidase; DDC, dopa decarboxylase; DBH, dopamine β-hydroxylase; 3-OMD, 3-methoxytyrosine; Dopac, dihydroxyphenyl acetic acid.

enzyme [13]. The existence of a thermolabile low-activity and a thermostable high-activity COMT in human population has been reported [18]. Interestingly, the two published sequences of human soluble COMT differ in only one amino acid. Recent kinetic studies have shown that this difference affects unambiguously the thermostability of the enzyme [19]. C. Kinetics of Human COMT The kinetic mechanism of the methylation reaction of human COMT has been studied exhaustively using recombinant enzymes [19]. The mechanism is sequential ordered: AdoMet binding first, then Mg2+ and the catechol substrate as the last ligand. Human S-COMT and MB-COMT have similar kinetic properties. The main difference is the one-order lower Km value of MB-COMT for dopamine as substrate (S-COMT 207 µM and MB-COMT 15 µM). The COMT enzyme is a rather slow enzyme with a low catalytic number. At saturating substrate levels S-COMT has a double efficiency compared with MB-COMT (kcat=37 and kcat =17, respectively). At low substrate concentrations (<10 µM) the MBCOMT seems to methylate catecholamines more rapidly than S-COMT.

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Figure 4 Amino acid sequence comparison of known COMT sequences (rat [11], human [14], pig [13]). Secondary structure elements in the sequence of rat soluble COMT are indicated as well as important active-site residues involved in binding of ligands (a, AdoMet; m, magnesium; s, substrate/inhibitor). The numbering of the residues corresponds to the soluble enzyme. The extension of the MB-COMT consists of the first 50 amino terminal residues. In the human sequence of COMT determined by Bertocci [13], Val108 is replaced by Met108.

Under physiological concentrations of catecholamines in the brain, MB-COMT may play a more important role than S-COMT [19,20]. D. Three-Dimensional Structure of COMT Backbone The crystal structure of rat soluble COMT has been solved at 2.0 Å resolution [21]. The COMT enzyme has a single domain α/β-folded structure, in which eight α-helices are arranged around the the central mixed β sheet. The sheet contains five parallel β strands and one antiparallel β hairpin. An overview of COMT is illustrated in Figure 5.

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The AdoMet binding motif is similar to the Rossmann fold, which is well known from the nucleotide binding proteins [22]. It has been shown that the known crystal structures of methyltransferases are strikingly similar in the AdoMet-binding regions [23], which indicates that all AdoMet-utilizing enzymes may share a common divergent evolution.

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Figure 5 Schematic stereo view of the three-dimensional structure of COMT. The ligands bound to COMT are the methyl-donating coenzyme AdoMet and the magnesium ion. Figures 5–7 and 13 were produced using the program MOLSCRIPT [50].

Figure 6 Stereo view of the AdoMet binding to COMT. The most important amino acid residues are shown as well as the magnesium ion and the inhibitor 3, 5-dinitrocate-chol (DNC).

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Active Site The active site of COMT consists of the AdoMet binding domain and the catalytic site. The structural elements and individual interactions between COMT residues and the enzymatic-action-participating ligands are demonstrated in detail in Figures 5 to 8. AdoMet Binding by COMT. The active-site residues, which have significant interactions with the coenzyme, are shown in Figure 6. The loop region between strand β1 and helix α4 forms the AdoMetbinding consensus sequence (in COMT GAxxG) that is conserved in methyltransferases [23]. In this region the terminal amino and carboxyl groups of AdoMet are bound. The last residue of the strand β2, Glu90, forms a hydrogen bond to the ribose hydroxyls. The residue Met91 has face-to-face van der Waals contacts on one side, and His142 has edge-to-face contacts on the opposite side of the adenine ring. The residue Trp143 closes the adenine of AdoMet into the protein with face-to-edge contact. Furthermore, the N-6 atom of the adenine hydrogen binds to Ser119. The Met40 residue holds the sulphur of AdoMet with the methyl group in right position towards the hydroxyl group of the catechol substrate. As a result of the various hydrogen bonds and van der Waals contacts, AdoMet has a high affinity to COMT with a dissociation constant of 23 µM [19]. Catalytic Site. The catalytic site of COMT is a rather simple environment formed by the metal ion and by the amino acids important for substrate binding and catalysis of the methylation reaction. The magnesium ion plays a crucial role for the catalytic activity of COMT. Figure 7 shows the binding of magnesium to COMT as derived from the

Figure 7 Magnesium binding in COMT. The magnesium ligands are Asp141, Asp169, Asn170, both hydroxyls of 3, 5-dinitrocatechol (DNC) and a water molecule (W).

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Figure 8 The catalytic machinery of COMT. Shown are the COMT residues important for catalysis, Mg2+ binding, the catechol as substrate, and the methyl-donating coenzyme AdoMet. The hydrophobic walls are defined by two tryptophane residues and a proline residue.

crystallographic studies. Magnesium has an octahedral coordination to two aspartic acid residues (Asp141 and Asp169), to an asparagine residue (Asn170), to both catechol hydroxyls of the substrate, and to a water molecule. In addition to the Mg2+ ion, Lys144 and Glu199 participate directly in the methylation reaction as shown in Figure 8. The “gate keeper” residues Trp38 and Pro174 form the hydrophobic “walls” and define the selectivity of the enzyme to different side chains of the substrate. They play a significant role in the binding of the substrates and inhibitors of COMT [19, 21]. III. Mechanism of the Catalytic Action of COMT The catalytic site is a shallow groove with the catalytic machinery at the bottom as illustrated in Figure 8. The two hydroxyl oxygens of a catechol substrate bind directly to the Mg2+ ion. The active methyl group of AdoMet is near one of the hydroxyl groups, on one side of the catechol ring. The amino group of Lys144 is also located near this hydroxyl group, on the other side of the catechol ring from AdoMet. The Glu199 residue is near the other hydroxyl group.

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Figure 9 The energy profile (as calculated at the 3-21G/PM3 level [24]) for the proton transfer step A rarrow.gif B and the methyl transfer step B rarrow.gif C for catechol as a substrate.

The pKa of the catecholic hydroxyl is about 9.8. The role of the Mg2+ ion bound to the enzyme is to make the hydroxyl groups more easily ionizable. It has been shown by quantum mechanical calculations that the hydroxyl protons can be transferred to Lys144 and Glu199 [24]. The proton transfer OH rarrow.gif Lys144 activates the hydroxyl group for the methyl transfer AdoMet rarrow.gif O-. Thus the reaction coordinate for methylation of catechols by COMT consists of a proton transfer from a hydroxyl group to Lys144 and a subsequent methyl transfer from AdoMet to the hydroxyl group (Figure 9). The Lys144 residue acts as a typical catalytic base in a general base-catalysed SN2-like nucleophilic substitution reaction [24]. IV. Inhibitors of COMT A. First-Generation Inhibitors First generation COMT inhibitors such as pyrogallol, U-0521 (3, 4-dihydroxy-2-methylpropiophenone), tropolone, and 8-hydroxyquinoline (Figure 10) were http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_351.html (1 of 2) [4/5/2004 5:28:20 PM]

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Figure 10 Structures of some first generation COMT inhibitors: pyrogallol, U-0521 (3, 4-dihydroxy-2-methylpropiophenone), tropolone, and 8-hydroxyquinoline.

Figure 11 Structures of second-generation COMT inhibitors discussed in this paper: nitecapone (OR-462), entacapone (OR-611), tolcapone (RO-40-7592), and 2-((3, 4-dihy-droxy-2-nitrophenyl) vinyl) phenylketone. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_352.html (1 of 2) [4/5/2004 5:28:27 PM]

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Page 353 Table 1 Inhibitor Constants of Selected COMT Inhibitors Ki (nM) Humanb

Ki (nM)Pigc

Nitecapone

1.0

700

Entacapone

0.3

N.D.

Tolcapone

0.3

N.D.

Vinylphenylketone

4.0a

200

aT.

Lotta, unpublished results

bReference

19.

cReference

41.

N.D.: not determined

used as in vitro tools to investigate COMT inhibition, but because of the lack of potency and selectivity and because they were toxic, they were not clinically useful [2]. These inhibitors have inhibition constants (Ki values) in the micromolar range. Many of them contain the catechol structure and are also substrates of COMT. B. Second-Generation Inhibitors The invention of a new structural family of COMT inhibitors in the late 1980s lead the COMT research into a new active epoch [25,26]. The most potent second-generation inhibitors are nitrocatechol derivatives; some examples are shown in Figure 11. Entacapone (OR-611) and tolcapone (RO-40-7592) are now in clinical trials for the treatment of Parkinson's disease, and have been extensively studied [27–40]. Both compounds are very potent and selective tightbinding inhibitors of human COMT with Ki values of 0.3 nM [19]. They differ mainly in their pharmacokinetic properties. Entacapone acts peripherally while tolcapone inhibits COMT both peripherally and centrally. Recently it was reported that 2-((3,4-dihydroxy-2-nitrophenyl)vinyl) phenylketone is a tight-binding inhibitor of pig COMT [41], and it is also a potent inhibitor of human COMT (Table 1). This inhibitor has a vinylphenylketone group at position 4 of the catechol ring, i.e., in the position ortho to the nitro group. V. Enzyme Inhibitor Interactions A. Background http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_353.html (1 of 2) [4/5/2004 5:28:31 PM]

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From quantitative structure activity relationship studies (QSAR) of COMT inhibitors it became evident that the acidity of the catechol hydroxyl group is the most important factor that influences the inhibitory activity of catechol

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derivatives [26,42]. Structures containing a catechol ring optimally substituted with a nitro group in position 3 and other electron-withdrawing substituents in position 5 showed high-potency inhibition [25,26,42,43]. It was also detected that the side-chain hydrophobicity at position 5 correlates significantly with the inhibitor activity [42]. B. X-Ray Structures of COMT-Drug Complexes The structure of 3,5-dinitrocatechol complexed with COMT has been solved [21]. This inhibitor is a typical nitrocatechol derivative with a high affinity for COMT. The excellent electron density of the inhibitor in the active site of COMT is represented in Figure 12. The planar structure of this compound fits well into the active-site cavity of the enzyme and forms nearly ideal contacts

Figure 12 A portion of the structure model and the 2F0-Fc electron density map contoured at 1.0 standard deviation. The region containing the inhibitor, magnesium, parts of AdoMet, and the residue Glu199 is shown. The sphere marked with “W” represents a water molecule.

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with the tryptophane residues 38 and 143. The hydroxyl groups of the inhibitor are coordinated to the Mg2+ ion. The hydroxyl group in position 1 has an important hydrogen bond to the carboxyl group of Glu199. The other hydroxyl is near the methyl group donated by AdoMet. The pKa of this hydroxyl group is low (about 3.4) [26]. The 3-nitro group of the inhibitor has favourable van der Waals interactions with Trp143. The Trp38 residue is located edge-to-face with the catechol plane, which allows an ideal aromatic hydrophobic contact. Such aromatic hydrophobic interactions have been described to be important in proteins and for the binding of ligands [44,45]. The binding mode of the catechol ring of other crystallographically determined potential nitrocatecholtype inhibitors complexed with COMT is essentially the same as with 3,5- dinitrocatechol (J.Vidgren, unpublished results). C. Differences in the Active Site of Human, Rat, and Pig COMT Models for human and pig COMT are easy to build using the experimental structure of the rat COMT, due to the high degree of homology between the rat, human, and pig COMT enzymes (Figure 4). The active sites are especially well conserved—the few differences in the active-site residues are collected in Table 2. The kinetic data show that the Km values of common substrates for rat and human COMT are very similar. Pig COMT shows, however, a considerably higher Km value for catechol [46]. The same difference is apparent for inhibitors represented by the Ki values in Table 1. The model for the binding of vinylphenylketone to pig and rat COMT is shown in Figure 13. Assuming that the catechol part of the inhibitor adopts the same position as found in the crystal structure with dinitrocatechol, the vinylphenylketone substituent has enough room to bind to both enzymes. The most significant difference between these enzymes lies in residue 38, the hydrophobic tryptophan in rat (and human) COMT and the polar arginine in pig COMT. If Arg38 is directed towards the hydrophobic core of the enzyme in a similar conformation as Trp38 (shown in Figure 13), it causes repulsion with the catechol ring of the inhibitor. However, it is probable that the polar Arg38 is directed towards the solvent. In this case the substrates and inhibitors will lack the favorable contacts that exist with Trp38 in human and rat enzymes. ObviTable 2 Differences in the Active Sites Between Rat, Human, and Pig COMT Position

Rat

Human

Pig

38

Trp

Trp

Arg

173

Val

Cys

Cys

201

Met

Arg

Ser

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Figure 13 The inhibitor 2-((3,4-dihydroxy-2-nitrophenyl)vinyl)phenylketone modeled into the active site of rat (a) and pig (b) COMT.

ously this one amino acid differences in the active site of the isoenzymes can explain the significant differences in the inhibitory potency of vinylphenylketone against these enzymes. From the structural point of view it seems that the position of the side-chain substitution (for example at C5 in entacapone and C4 in vinylphenylketone) is not critical for the inhibitor binding. In both cases the substituent has sufficient space to adapt to the protein structure, and in fact, large substituents reach from the active site cavity to the solvent (Figure 13). The tenfold higher inhibitory activity of entacapone compared with vinylphenylketone against human COMT can be accounted for by the electron- withdrawing effect of the side-chain substitution. In the case of entacapone, the side chain at position C5 has a more beneficial electronic influence on the 2hydroxyl of the inhibitor producing a better inhibition (see Section VI). VI. Mechanism of the COMT Inhibition By Nitrocatechols As described above, catechols with strong electronegative groups are potent inhibitors of COMT. These compounds seem to bind well to the active site, but in spite of that they are very poor substrates. It has been shown, with nitecapone

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Figure 14 The energy profile (as calculated at the 3-21G//PM3 level [24]) for the proton transfer step A rarrow.gif B and the methyl transfer step B rarrow.gif C for 3,4-dinitrocatechol as a substrate.

in a rat COMT assay, that the rate of nitecapone methylation is equivalent to about 1% of the rate of dopamine methylation [47]. The energy profile for the hypothetical methylation of 3,5-dinitrocatechol is shown in Figure 14 [24]. The electronegative nitro groups strongly stabilize the ionized catechol–COMT complex, and the energy barrier for the methylation step is high (see Figure 9 for comparison). This can be understood as decreased nucleophilicity of the hydroxyl oxygen, due to the electron-with- drawing properties of the nitro groups. The electronic effect of the substituents of the catechol ring to the nucleophilicity of the hydroxyl group at the active site can be readily seen from the molecular electrostatic potential (MEP) surfaces of the system. The MEP surfaces were calculated at the PM3 level and plotted at –20 kcal/mol for catechol and 3,5-dinitrocatechol at the active site of COMT [24]. The results are summarized in Figure 15. In the case of catechol the effect of the proton transfer form OH to Lys144 is seen as a remarkable increase in the negative potential between AdoMet and the substrate. 3,5-Dinitro substitution of the catechol ring http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_357.html (1 of 2) [4/5/2004 5:28:58 PM]

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Figure 15 The energy profiles and MEP surfaces (–20 kcal/mol) for catechol (black) and 3,5-dinitrocatechol (grey).

decreases the activating negative potential substantially. As a consequence, the OH group is weakly nucleophilic and 3,5-dinitrocatechol is not a substrate of COMT but a potent inhibitor. VII. Inhibitor Design The drug-design process of COMT inhibitors started long before the structure of the target molecule was available. The most important results were extracted from the QSAR studies of substituted catechols [26,42]. Those investigations clearly indicated the importance of the acidity of one of the two hydroxyl groups in the catechol ring. The ionization of the hydroxyl was greatly influenced by electronwithdrawing substituents in the positions ortho and para to the hydroxyl. The lipophilicity of the side chain was predicted, but after the determination of the enzyme structure it became clear that the active site of COMT is a relatively shallow groove where ligands with longer side chains

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easily reach the surface of the enzyme. The pharmacophore model constructed with biochemical and QSAR knowledge [42] was surprisingly good and realistic in comparison to the experimental structure. The crucial role of the magnesium ion for the binding of substrates and catalysis was not given enough attention. Even without the experimental knowledge of the three-dimensional structure of COMT, the correct decision for the direction of the drug-design process was possible. Many open questions were still waiting for the determination of the molecular structure. The most important drug-design aspects under consideration included the optimization of the pharmacokinetic properties of the molecules, especially the penetration across the blood brain barrier. The structure of COMT clearly showed that with catechol-type inhibitors, one of the positions ortho to the hydroxyls is optimally substituted by a nitro group, while the other ortho position has to be unsubstituted. Sterically, the two remaining sites can be substituted quite freely, so that these substitution positions can be used to modify the physicochemical properties of the inhibitors. The question of the possibility of designing an inhibitor without the catechol structure is important, because the potent inhibitors which rely on the ionization of a catechol hydroxyl, penetrate very poorly into the brain. In clinical trials of the therapeutic use of COMT inhibitors it has become evident that the beneficial effect to L- dopa metabolism is fully reached with peripheral inhibitors such as entacapone [27–29]. As demonstrated above, based on the threedimensional structure of COMT, the design of potent noncatechol type inhibitors may be very tedious. The active site of COMT is a rather simple environment with a few catalytic residues and a magnesium ion defining the structural limits of the catechol ligands. VIII. Clinical Possibilities of Inhibitors of COMT A. Parkinson's Disease Parkinson's disease is a neurological disorder that affects voluntary movement. The symptoms are slowness of movement, rigidity, and tremor. The reason for the disease is unknown. Parkinson's disease is a progressive disorder and involves the deterioration of dopaminergic nerve fibers in substantia nigra, which leads to a striatal deficiency of dopamine. The symptoms of Parkinson's disease are detected only after about 80% of the dopamine-producing neurons are degenerated. There is no known cure but the symptoms are treated with a combination of different drugs. Current and Future Therapy of Parkinson's Disease Dopamine does not penetrate the blood-brain barrier. L-Dopa is actively transported into the brain and then converted to dopamine. L-Dopa is rapidly metabolized and only about 1% of an oral dose reaches the brain. In the current

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therapy of Parkinson's disease L-dopa (precursor of dopamine) is administered together with a peripheral dopadecarboxylase (DDC) inhibitor (such as carbidopa or benserazide). The major peripheral L-dopa metabolizing enzymes are DDC and COMT. After inhibition of DDC, COMT is responsible for the main catabolism of L-dopa. In the central nervous system COMT together with mono- amino oxidase (MAO) participitates in the metabolism of L-dopa and dopamine. Large amounts of orally administered L-dopa are converted by COMT to 3-O-methyldopa (3-OMD). Having a long plasma halflife (approximately 15 h compared with the dopamine 1-h half-life), 3-OMD accumulates in the plasma during the L-dopa treatment. L-Dopa and 3-OMD also compete for the same active transport system into the brain. It has been proposed that 3-OMD could cause some side effects of the L-dopa treatment (dyskenisia, on-off phenomenon). Inhibition of COMT enzyme decreases the 3-OMD formation and improves the brain entry and bioavailability of L-dopa. The use of COMT inhibition should prolong the L-dopa effects and permit a decreased does [27– 29]. Preclinical and clinical results indicate that both entacapone and tolcapone are orally active, nontoxic and well-tolerated drugs. The adjuvant L- dopa therapy with DDC inhibitor + COMT-inhibitor (+ possible MAO inhibitor) may substitute for the present double therapy in the treatment of Parkinson's disease [27–40]. Together with the development of dopamine agonists and MAO inhibitors, the inhibition of COMT will constitute major progress in the treatment of Parkinson's disease in the near future. B. Other Possible Indications of the COMT Inhibition It has been proposed that COMT inhibitors co-administered with L-dopa could have beneficial effects in the treatment of depressive illness [40]. This can be caused by either the better availability of dopamine or by the elevated noradrenaline levels in the brain. Another hypothesis suggests that the increasing level of AdoMet caused by COMT inhibition may cause an antidepressive effect [33]. Dopamine has also natriuretic and diuretic effects in kidney. There has been evidence that abnormalities of the renal dopamine system can lead to salt- sensitive hypertension [48]. In rat kidney, deamination represents the major pathway in the metabolism of dopamine, but when MAO is inhibited, methylation appears to offer an alternative metabolic pathway [49]. Thus COMT inhibition may be important in the regulation of renal sodium excretion. References 1. Axelrod J, Tomchick R. Enzymatic O-methylation of epinephrine and other catechols. Journal of Biological Chemistry 1958; 233:702–705.

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2. Guldberg H, Marsden C. Catechol-O-methyl transferase: pharmacological aspects and Physiological role. Pharmacological Reviews 1975; 27:135–206. 3. Rivett AJ, Francis A, Roth JA. Localization of membrane-bound catechol-O- methyltransferase. Journal of Neurochemistry 1983; 40:1494–1496. 4. Karhunen T, Tilgmann C, Ulmanen I, Julkunen I, Panula P. Distribution of catechol-Omethyltransferase enzyme in rat tissues. Journal of Histochemistry and Cytochemistry 1994; 42:1079–1090. 5. Axelrod J. Methylation reactions in the formation and metabolism of catecholamines and other biogenic amines. Pharmacological Reviews 1966; 18:95– 113. 6. Ball P, Knuppen R, Haupt M, Breuer H. Interactions between estrogens and catechol amines III. Studies on the methylation of catechol estrogens, catechol amines and other catechols by the catechol-Omethyltransferase of human liver. Journal of Clinical Endocrinology 1972; 34:736–746. 7. Borchardt RT. N- and O-methylation. In: Jakoby WB, ed. Enzymatic Basis of Detoxification. Vol. 2. New York: Academic Press, 1980:43–62. 8. Nutt JG, Fellman JH. Pharmacokinetics of levodopa. Clinical Neuropharmacology 1984; 7:35–79. 9. Assicot M, Bohuon C. Presence of two distinct catechol-O-methyltransferase activities in red blood cells. Biochimie 1971; 53:871–874. 10. Nissinen E, Männistõ P. Determination of catechol-O-methyltransferase activity by high performance liquid chromatography with electrochemical detection. Analytical Biochemistry 1984; 137:69–73. 11. Salminen M, Lundstrõm K, Tilgmann C, Savolainen R, Kalkkinen N, Ulmanen I. Molecular cloning and characterization of rat liver catechol-O-methyltransferase. Gene 1990; 93:241–247. 12. Tilgmann C, Kalkkinen N. Purification and partial characterization of rat liver soluble catechol-Omethyltransferase. FEBS Letters 1990; 264:95–99. 13. Bertocci B, Garotta G, Da Prada M, et al. Immunoaffinity purification and partial amino acid sequence analysis of catechol-O-methyltransferase from pig liver. Biochimica et Biophysica Acta 1991; 1080:103–109. 14. Lundström K, Salminen M, Jalanko A, Savolainen R, Ulmanen I. Cloning and characterization of human placental catechol-O-methyltransferase cDNA. DNA and Cell Biology 1991; 10:181–189.

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15. Tilgmann C, Kalkkinen N. Purification and partial sequence analysis of the soluble catechol-Omethyltransferase from human placenta: Comparison to the rat liver enzyme. Biochemical and Biophysical Research Communications 1991; 174:995–1002. 16. Lundström K, Tilgmann C, Peränen J, Kalkkinen N, Ulmanen I. Expression of enzymatically active rat liver and human placental catechol-O-methyltransferase in Escherichia coli; purification and partial characterizaton of the enzyme. Biochimica et Biophysica Acta 1992; 1129:149–154. 17. Tenhunen J, Salminen M, Jalanko A, Ukkonen S, Ulmanen I. Structure of the rat catechol-Omethyltransferase gene: separate promoters are used to produce mRNAs for soluble and membranebound forms of the enzyme. DNA and Cell Biology 1993; 12:253–263. 18. Boudikova B, Szumlanski C, Maidak B, Weinshilboum R. Human liver catechol- Omethyltransferase pharmacogenetics. Clinical Pharmacology and Therapeutics 1990; 48:381–389.

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19. Lotta T, Vidgren J, Tilgmann C, et al. Kinetics of human soluble and membrane- bound catechol-Omethyltransferase: a revised mechanism and description of the thermolabile variant of the enzyme. Biochemistry 1995; 34:4202–4210. 20. Roth JA. Membrane-bound catechol-O-methyltransferase: A reevaluation of its role in the Omethylation of the catecholamine neurotransmitters. Reviews of Physiology, Biochemistry and Pharmacology 1992; 120:1–29. 21. Vidgren J, Svensson LA, Liljas A. Crystal structure of catechol-O-methyltransferase. Nature 1994; 368:354–358. 22. Rossman M, Liljas A, Bränden C I, Banaszak L. Evolutionary and structural relationships among dehydrogenases. In: Boyer PD, ed. Enzymes. New York: Academic Press, 1975:61–102. 23. Schluckebier G, O'Gara M, Saenger W, Cheng X. Universal catalytic domain structure of adometdependent methyltransferases. Journal of Molecular Biology 1995; 247:16–20. 24. Ovaska M. The mechanism of catalysis and inhibition of catechol-O-methyltransferase. Submitted 1996. 25. Bäckström R, Honkanen E, Pippuri A, et al. Synthesis of some novel potent and selective catechol-Omethyltransferase inhibitors. Journal of Medicinal Chemistry 1989; 32:841–846. 26. Borgulya J, Bruderer H, Bernauer K, Zurcher G, Da Prada M. Catechol-O- methyltransferaseinhibiting pyrocatechol derivatives: synthesis and structure- activity studies. Helvetica Chimica Acta 1989; 72:952–968. 27. Nutt JG, Woodward WR, Beckner RM, et al. Effect of peripheral catechol-O- methyltransferase inhibition on the pharmacokinetics and pharmacodynamics of levodopa in parkinsonian patients. Neurology 1994; 44:913–919. 28. Ruottinen H, Rinne UK, Ahtila S, Karlsson M, Kyyrä T, Gordin A. Entacapone increases levodopa response in a one-month double-blind study in parkinsonian patients with fluctuations. Neurology 1995; 45:412S. 29. Ruottinen H, Rinne UK. Entacapone prolongs levodopa response in a one-month double-blind study in parkinsonian patients with levodopa related fluctuations. Journal of Neurology, Neurosurgery, and Psychiatry 1996; 60:36–40. 30. Nutt JG. Effects of catechol-O-methyltransferase (COMT) inhibition on the pharmacokinetics of LDOPA. Advances in Neurobiology. Vol. 69. Philadelphia: Lippincott-Raven Publishers, 1996:493–496.

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31. Törnwall M, Männistö PT. Acute toxicity of three new selective COMT inhibitors in mice with special emphasis on interaction with drugs increasing catecholaminergic neurotransmission. Pharmacology and Toxicology 1991; 69:64–70. 32. Törnwall M, Männistö PT. Effects of three types of catechol-O-methylation inhibitors on 1-3,4dihydroxyphenylalanine-induced circling behaviour in rats. European Journal of Pharmacology 1993; 250:77–84. 33. Da Prada M, Borgulya J, Napolitano A, Zürcher G. Improved theraphy of parkinson's disease with tolcapone, a central and peripheral COMT inhibitor with an S- adenosyl-L-methionine-sparing effect. Clinical Neuropharmacology 1994; 17:26– 37. 34. Kaakkola S, Gordin A, Männistö PT. General properties and clinical possibilities of new selective inhibitors of catechol-O-methyltransferase. General Pharmacology 1994; 25:813–824.

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35. Davis TL, Roznoski M, Burns RS. Effects of tolcapone in parkinson's patients taking Ldihydroxyphenylalanine/carbidopa and selegiline. Movement Disorders 1995; 10:349–351. 36. Deleu D, Sarre S, Ebinger G, Michotte Y. The effect of carbidopa and entacapone pretreatment of the L-dopa pharmacokinetics and metabolism in blood plasma and skeletal muscle in beagle dog: an in vivo microdialysis study. Journal of Pharmacology and Experimental Therapeutics 1995; 273:1323–1331. 37. Napolitano A, Zürcher G, Da Prada M. Effect of tolcapone, a novel catechol-O-methyltransferase inhibitor, on striatal metabolism of L-DOPA and dopamine in rats. European Journal of Pharmacology 1995; 273:215–221. 38. Dingemanse J, Jorga KM, Schmitt M, et al. Integrated pharmacokinetics and pharmacodynamics of the novel catechol-O-methyltransferase inhibitor tolcapone during first administration to humans. Clinical Pharmacology and Therapeutics 1995; 57:508–517. 39. Männistö PT. Clinical potential of catechol-O-methyltransferase (COMT) inhibitors as adjuvants in Parkinson's disease. CNS Drugs 1994; 1:172–179. 40. Männistö PT, Lang A, Rauhala P, Vasar E. Beneficial effects of co-administration of catechol-Omethyltransferase inhibitors and l-dihydroxyphenylalanine in rat models of depression. European Journal of Pharmacology 1995; 274:229–233. 41. Perez RA, Fernandez-Alvarez E, Nieto O, Piedrafita FJ. Kinetics of the reversible tight-binding inhibition of pig liver catechol-O-methyltransferase by [2-(3,4-dihydroxy-2-nitrophenyl)vinyl]phenyl ketone. Journal of Enzyme Inhibition 1994; 8:123–131. 42. Taskinen J, Vidgren J, Ovaska M, Bäckström R, Pippuri A, Nissinen E. QSAR and binding model for inhibition of rat liver catechol-O-methyltransferase by 1,5- Substituted-3,4-Dihydroxybenzenes. Quantitative Structure Activity Relationships 1989; 8:210–213. 43. Lotta T, Taskinen J, Bäckström R, Nissinen E. PLS Modeling of structure-activity relationships of catechol-O-methyltransferase inhibitors. Journal of Computer- Aided Molecular Design 1992; 6:253–272.

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44. Burley SK, Petsko GA. Aromaticaromatic interaction: A mechanism of protein structure stabilization. Science 1985; 229:23–28. 45. Serrano L, Bycroft M, Fersht AR. Aromatic-aromatic interactions and protein stability. Journal of Molecular Biology 1991; 218:465–475. 46. Piedrafita FJ, Elorriaga C, Fernandez-Alvarez E, Nieto O. Inhibition of catechol- Omethyltransferase by N-(3,4-dihydroxyphenyl) maleimide. Journal of Enzyme Inhibition 1990; 4:43–50. 47. Wikberg T. Docotoral Thesis, University of Helsinki, Helsinki, Finland, 1993:29– 30. 48. Aperia A. Dopamine action and metabolism in the kidney. Current Opinion in Nephrology and Hypertension 1994; 3:39–45. 49. Fernandes MH, Soares-da-Silva P. Role of monoamine oxidase and catechol-O- methyltransferase in the metabolism of renal dopamine. Journal of Neural Transmission. Supplementum 1994; 41:101–105. 50. Kraulis PJ. MOLSCRIPT: a program to produce both detailed and schematic plots of protein structures. Journal of Applied Crystallography 1991; 23:946–950.

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15 Antitrypanosomiasis Drug Development Based on Structures of Glycolytic Enzymes Christophe L. M. J. Verlinde, Hidong Kim, Bradley E. Bernstein, Shekhar C. Mande, * and Wim G.J. Hol University of Washington, Seattle Washington I. Trypanosomiasis A. Disease and Treatment Human African trypanosomiasis, also called sleeping sickness, is caused by the parasites Trypanosoma brucei gambiense and Trypanosoma brucei rhodesiense. These unicellular organisms live freely in the bloodstream of the human host and invade the brain during the later stage of the disease. Without treatment the disease is always fatal [1]. The course of the gambiense form may last from months to years, while T. brucei rhodesiense usually kills within weeks. Sleeping sickness occurs in thirty-six subSaharan African countries, putting fifty million people at risk. Each year 25,000 new cases are reported, but the actual number of cases is more likely to be about 250,000 [2]. The current state of antitrypanosomal chemotherapy is dismal; many parasitologists do not want to take the risk of being infected with T. brucei rhodesiense and prefer to study the parasite T. brucei brucei, which is harmless to humans [3]. Not more than four drugs are available to treat the disease: pentamidine, suramin, melarsoprol—all from the first half of this century—and eflornithine, introduced in 1990 (Figure 1). Except for eflornithine, which is an irreversible ornithine decarboxylase inhibitor [4], the mechanisms of the drugs are poorly understood [5]. All four drugs require administration by injection in a hospital setting, which is a major drawback in rural Africa [6]. Pentamidine is useful for treating early stage T. brucei gambiense infection, suramin for both early stage gambiense and rhodesiense infection. The permanent charges on pentamidine *

Current affiliation: Institute of Microbial Technology, Chandigarh, India

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Page 366

Figure 1 Available drugs for the treatment of human African trypanosomiasis.

and suramin explain why they show poor oral absorption and do not cross the blood–brain barrier, making them unsuitable for the treatment of late-stage trypanosomiasis. Melarsoprol, an organoarsenical compound, was until 1990 the only drug effective in the late stage of both forms of trypanosomiasis. Unfortunately, it is also highly toxic, causing reactive encephalopathy in up to 10% of the patients, of which about one half die. This deadly complication is well known to villagers of areas where the disease is endemic and, ironically, discourages people from participating in diagnostic surveys. Eflornithine, heralded as the “resurrection drug” upon its introduction [7], cures patients infected with late-stage T. brucei gambiense but is ineffective against the more virulent rhodesiense form. Additionally, it causes bone marrow suppression in half of the patients and occasionally convulsions [6]. A serious concern is that resistance has been reported against each of the four antitrypanosomal drugs [1].

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Sleeping sickness has been largely ignored by the pharmaceutical industry because the poor socioeconomic situation in the part of the African continent afflicted by this debilitating disease offers little prospect of reasonable financial returns [8]. It is revealing that eflornithine was originally developed as an anticancer drug. It was screened for antitrypanosomal properties only when the biochemistry of the trypanosomal polyamine metabolism was understood [9]. Fortunately, the cell biology of trypanosomes is so extraordinary that they have been the subject of more fundamental research than most other protozoan parasites [5]. Each of these unique features of T. brucei may become a target for new drugs, provided they prove to be essential for the survival of the parasite in the human host. With such a wealth of biochemical information, structure-based drug design provides a tremendous opportunity to arrive at new drugs to cure sleeping sickness. B. Targets for Future Drugs Preventing trypanosomiasis would be a nobler goal than curing it. Unfortunately, trypanosomes are experts in evading our immune system. They achieve this by varying their dense surface coat. It is composed of ten million copies of a single protein, the variant surface glycoprotein (VSG), for which they have no less than a thousand different genes. In this way trypanosomes can change surface antigens more rapidly than the host can produce new antibodies [10]. Clearly, such a mechanism leaves little hope for preventing sleeping sickness by vaccination. In contrast to the small number of drugs available to treat trypanosomiasis, the opportunities for developing new drugs are ample, as can be seen from Table 1. They range from unique RNA processing to reduced metabolism, salvage systems, and different rates of protein turnover. All of these features were the subject of an outstanding review by C.C. Wang [5]. An inventory of the structural information waiting to be exploited by structure-based drug design reveals eight potential target enzymes (Table 2). Since for most trypanosomatid proteins there is a human counterpart it is mandatory that designed inhibitors be selective, i.e., have very little affinity for the equivalent enzymes of the human host. As a consequence, selective design requires pairs of equivalent structures from the parasite and from the host. A complication for selective design is that for three of the mammalian enzymes only the structure of one isoenzyme is known. For example, the structure of human aldolase A has been determined [23] but not the structures of isoenzymes B and C, which share only 69 and 82% sequence identity to isoenzyme A [40–42]. Of course, homology modeling might be a way to overcome this problem. From Table 2 it is evident that only for three proteins, TIM, GAPDH, and trypanothione reductase the structures of the parasite and host enzymes are

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Page 368 Table 1 Trypanosomal Targets, Their Human Equivalents, and Current Leads for Drug Design Human host

T. brucei

Lead

RNA editing by trans-splicing [11]

cis-splicing

none

All energy from fast glycolysis [12]

glycolysis and oxidative phosphorylation

MMBAa[13]

Glucose transporter [14]

human erythrocyte glucose transporter

none

Purine P2 transporter [15]

none

Purine salvage enzymes, e.g., HGPRTb[16]

HGPRT

Slow rate of enzyme turnover [17], e.g., ornithine decarboxylase [18]

fast rate of enzyme turnover

Polyamine metabolism, e.g., S-adenosyl methionine decarboxylase [19]

S-adenosyl methionine decarboxylase

MDL 73811c

Trypanothione reductase [20]

glutathione reductase

mepacrine

VSG anchor: a myristate-containing GPI [21] aMMBA

= 2' -deoxy-2' -(m-methoxybenzamido)-adenosine.

bHGPRT

= hypoxanthine guanine phosphoribosyltransferase.

cMDL

none eflornithine (= drug) [9]

10-(propoxy)-decanoated

73811 = 5' -{[(Z)-4-amino-2-butenyl]methylamino}-5' -deoxyadenosine.

dIt

has not been established whether the trypanocidal effect of this compound is due to its incorporation in the GPI anchor [5].

known. For five more targets the crystal structure of only the mammalian enzyme is available. Efforts to solve the structures of trypanosmatid aldolase, PGK, and PK counterparts are underway in our lab. This review explains how we attempted to arrive at selective inhibitors of three trypanosomal glycolytic enzymes. C. Trypanosomal Glycolysis: Enzyme Inhibition as a Target In the bloodstream of the human host, trypanosomes are metabolically

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“lazy”. Since there is plenty of glucose and oxygen available, they rely solely on glycolysis to the stage of pyruvate for their energy supply [12]. Their glycolysis proceeds at an amazing rate, which is about fifty times faster than in the cells of the human host [43]. This fast rate is a necessity because only two molecules of ATP are generated per molecule of glucose instead of the thirty-six produced by complete oxidation. These findings led to the proposal that inhibitors of trypanosomal glycolysis might be turned into drugs. Support for this idea comes from in vitro experiments where salicylhydroxamic acid (SHAM) was used to bring the parasite under anaerobic conditions, after which glycolysis was

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Page 369 Table 2 Three-Dimensional Structures Available for Trypanosomal Drug Design Trypanosomatid

PDB code

Mammalian

PDB code

(a) Glycolytic: PGIa

—

—

Porcine

1PGI[22]

Aldolase

—

—

Humanb

1ALD[23]

TIM

T. brucei

5TIM[24]

Human

1HTI[25]

GAPDH

T. brucei L.mexicana

1GGA[26,27] [30]

Humanc

3GPD[28,29]

PGK

—

—

Horse Pig

2PGK[31]—[32]

PK

—

—

Catd

1PYK[33]

HumanGR

3GRS[35]

Human

1HMP[38]

(b) Other: TR

T. cruzi

1NDA[34]

C. fasciculata

1TYT[36] 1PPR[37]

—

HGPRT VSG

T. brucei

— 1VSG[39]

No equivalent

aAbbreviations:

PGI = phosphoglucose isomerase, PGK = phosphoglycerate kinase, PK = pyruvate kinase, TR = trypanothione reductase; GR = glutathione reductase, HGPRT = hypoxanthine-guanine phosphoribosyl transferase, VSG = variable surface glycoprotein. bA

isoenzyme, from muscle; other mammalian isoenzymes are B in liver and C in brain.

cMuscle

isoenzyme; a liver isoenzyme exists.

dM1

isoenzyme, from muscle; mammals also have M2 in kidney, adipose tissue and lung, L in liver, and R in rood blood cells.

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blocked by mass action through the addition of glycerol (Figure 2). As a result trypanosomes were lysed within five minutes [44,45]. Treatment of infected rodents with the SHAM/glycerol mixture proved to be effective to clear the blood of the animals from T. brucei [46], although only with sublethal doses was permanent aparasitemia obtained [47]. If glycolysis could be blocked selectively, i.e., without affecting the equivalent enzymes of the host, one might have a promising therapy against trypanosomiasis. D. Beyond Enzyme Inhibition: Protein Routing as a Target In trypanosomes, seven enzymes involved in glycolysis, from hexokinase to phosphoglycerate kinase, are sequestered in specialized organelles, called glycosomes [48]. These microbodies are probably evolutionary relics of an endosymbiont [49] but are devoid of genetic material encoding for the glycosomal enzymes. Instead, these enzymes are encoded in the nucleus and are post-translationally imported into the glycosome. Since the import process likely involves unfolding one might envision blocking import by stabilizing the folded

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Page 370

Figure 2 Glycolysis in bloodstream-form trypanosomes. All net energy production takes place in the cytosol as the result of ATP formation by pyruvate kinase. However, the majority of glycolytic enzymes are sequestered into a specialized organelle, the glycosome. There the net ATP synthesis is zero irrespective of the presence of oxygen. Under aerobic conditions the NADH produced by glyceraldehyde-3-phosphate dehydrogenase is reoxidized via the G-3-P/DHAP shuttle, which couples glycolysis to a mitochondrial glycerophosphate oxidase. Under anaerobic conditions or when the oxidase is blocked by SHAM, equimolar amounts of glycerol and 3-phosphoglycerate are formed. However, the addition of an excess of glycerol to the cytosol prevents the reoxidation of NADH. As a result trypanosomes treated with a mixture of SHAM and glycerol die in a matter of minutes.

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state through the tight binding of ligands. In this way inhibitors might act at two levels, directly by blocking catalysis and indirectly by preventing proper enzyme routing in the parasite. Thus far, we have engaged in a collaborative effort to inhibit three of the glycosomal enzymes. (From Ref. 95. Copyright 1988 by Elsevier.)

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II. Three Glycolytic Enzymes Of T. Brucei: Molecular Biology, Biochemistry, and X-Ray Crystallography A. Triosephosphate Isomerase (TIM) TIM is a homodimeric enzyme that interconverts dihydroxyacetone phosphate and glyceraldehyde-3phosphate. It ensures that both trioses derived from glucose can be used for ATP production in the glycolytic pathway. Triosephosphate isomerase does not require any cofactor. Both the T. brucei and human enzymes have been overexpressed in Escherichia coli [50,25] and their crystal structures were solved in our group [24,25]. In addition, the structures of T. brucei TIM in complex with seven nonselective competitive inhibitors, with inhibition constants of 300 µM or higher were determined: monohydrogen phosphate [51], 2-phosphoglycerate [52], 3-phosphoglycerate [53], 3phosphonopropionate [53], glycerol-3-phosphate [53], 2-(N-formyl-N-hydroxyamino)-ethyl phosphonic acid [54], and N-hydroxy-4-phosphonobutanamide [55]. These studies gave an excellent picture of different ligand binding modes and of the conformational flexibility of the enzyme. All ligands interact with the main features of the catalytic machinery of the enzyme (Figure 3): (1) the phosphate is sequestered by the positive end of a 310-helix and Lys13; (2) polar groups on the carbon framework interact with His95 and Glu167, the catalytic electrophile and the catalytic base of the enzyme, respectively; (3) the entire inhibitor is shielded from the bulk solvent by a flexible loop, which normally closes over the substrate during catalysis to prevent phosphate elimination [56] (Figure 4). The only exception to flexible loop closure is N-hydroxy-4-phosphono-butanamide. It binds to the enzyme with the flexible loop in the open conformation because its size precludes loop closure. Thus, the crystallographic binding studies point out that it should be possible to design two very different classes of selective inhibitors: a class that binds to the enzyme in the closed loop conformation and one that binds to the open loop conformation. Selective inhibitor design in the case of TIM appears to be a formidable task. All residues within 10 Å of the active site are conserved [25]. This is also reflected in the similarity of the kinetic characteristics between trypanosomal and human TIM: for T. brucei TIM, Km (glyceraldehyde-3-phosphate) = 0.25 mM, kcat = 3.7 × 105min-1 [57]; for human TIM, Km = 0.49 mM, kcat = 2.7 × 105 min-1 [25]. There are significant differences in the surface protein of the two enzymes about 15 Å away from the substrate phosphorus atom [58]. In a shallow cleft, T. bruceiTIM has Ala100-Tyr101, while the human counterpart of these residues is His-Val (Figure 5). The cleft is formed by the flexible loop of one subunit of the enzyme and a different loop originating from another subunit. When the flexible loop changes its conformation from the closed to the open form the cleft widens substantially. Moreover, the Ala-Tyr dipeptide becomes then directly accessible from the active site, the distance being about

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Figure 3 Schematic representation of the structure of a TIM monomer. Helices and strands are labeled as H and B, respectively. The view is along the axis of the β barrel, into the active site. Key catalytic residues Lys13, His95, and Glu167 are shown along with the helix that binds the substrate phosphate and the flexible loop that covers the substrate during catalysis. Black dots indicate residues in contact with the second monomer of the enzyme. (From Ref.24. Copyright 1991 by Harcourt Brace.)

10 Å instead of the 15 Å in the closed loop conformation of the enzyme. In any case, selective inhibitor design for TIM appears to require de novo design as there are no leads known that interact with the AlaTyr region of the enzyme. B. Glyceraldehyde-3-Phosphate Dehydrogenase (GAPDH) Glyceraldehyde-3-phosphate dehydrogenase is a homotetramer that carries out the oxidative phosphorylation of glyceraldehyde-3-phosphate into 1,3-bisphos- phoglycerate. During this reaction NADH is formed. Each subunit of the enzyme consists of two domains and has an NAD+ binding site. The N-terminal domain anchors the adenosine portion of the cofactor while the nicotinamide portion is involved in the catalytic reaction at the C-terminal domain. T. brucei

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Figure 4 Stereoview of superimposed TIM monomers, one with an open flexible loop (full black) and another one with a closed flexible loop (open gray). The loop location is marked by an asterisk. Note the proximity of the active site, indicated by the catalytic residues Lys13, His95, and Glu167 (all sterofigures in this paper were drawn with MOLSCRIPT [93]).

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Figure 5 Exploitable structural differences between T.brucei(full) and human (dashed) TIM. The inhibitor 2-phosphoglycolate as observed in the structure of the human enzyme indicates the location of the active site. Drug design targets are the T.brucei Ala100-Tyr101, which are considerably different from their human counterpart His-Val. (From Ref.25. Copyright 1994 by Cambridge University Press.)

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Figure 6 Stereoview of NAD binding by trypanosomal GAPDH (full black). For clarity only 2 of the 4 subunits are shown. A substantial deviation of the protein backbone occurs in human GAPDH (open gray, only one subunit shown) near the adenosine part of the cofactor. The nearby cleft (marked by an asterisk) is important for introducing selectivity in inhibitor binding and has therefore been termed “selectivity cleft.”

glycosomal GAPDH has been overexpressed in E. coli [59] while human erythrocyte GAPDH is available from commercial suppliers. The sequences of the two enzymes are only 55% identical [60]. The crystal structure of the parasite enzyme was solved from Laue data at 3.2 Å resolution in our group (Figure 6) and in a second crystal form at 2.8 Å [26]. The structure of the human muscle enzyme was solved at 3.5 Å resolution in the group of the late Herman Watson [28]. Its resolution was improved to 2.3 Å in our group [26,29]. Both structures were of the holo-enzyme, i.e., the enzyme in presence of the cofactor. The active site and the nicotinamide-binding site of the two enzymes are very well conserved. This is reflected in similar Km (glyceraldehyde-3-phos-phate) values of 0.15 and 0.17 mM for the trypanosomal and human enzyme, respectively [61]. Surprisingly, the Km (NAD+) values differ by a factor ten: 0.45 mM for T. brucei and 0.04 mM for human GAPDH [61]. In view of the conservation of the nicotinamide and pyrophosphate binding sites, the substantial difference in Km (NAD+) has to be ascribed to the adenosine binding environment. Indeed, some of the residues embracing the adenine ring of the cofactor are not identical. In the trypanosomal enzyme the adenine is sandwiched in between a Thr in the back and a Met in the front. This Met is replaced by Pro and Phe in the human enzyme (Figure 7). A second difference involves the residue flanking the C2 atom of the purine ring. It is a Val in the trypanosomal enzyme, and Asn in the human enzyme. These differences apparently account for a ten-fold lower affinity for the cofactor.

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Page 375

Figure 7 Comparison of the binding modes of the adenosine moiety of NAD to GAPDH: (left) in T.brucei,(right) in the human enzyme. Note the identical hydrogen bonds to the purine N6 and the ribosyl hydroxyls. The purine ring is embraced by hydrophobic residues that are not conserved. Also, a unique cleft near O2', which we called the selectivity cleft, is present in the T.brucei enzyme. (From Ref. 13.Copyright 1994 by the American Chemical Society.)

Other differences in the vicinity of the adenosine portion of the NAD+ cofactor are prime targets for selective inhibitor design. Close to C8 of the adenine ring, the trypanosomal GAPDH exhibits a Leu while its human counterpart has a smaller residue, namely Val (Figure 7). Also, the parasite enzyme possesses a hydrophobic cleft near the 2' -hydroxyl of the adenosine ribose. This cleft, termed the “selectivity cleft” is almost absent in the human enzyme due to a different local backbone conformation and the presence of the Ile37 side chain. In conclusion, the adenosine binding region looks like an excellent target for selective inhibitor design. It is exciting that the residues responsible for binding adenosine in T. brucei GAPDH are identical to their counterparts in glycosomal GAPDH of Leishmania mexicana, another trypanosomatid [31]. L. mexicana is one of the most common species of Leishmania throughout Central and South America and the southern United States. In humans it hides as amastigotes in the macrophages and causes hideous skin lesions. Together with about twenty other species of Leishmania these parasites infect about twelve million people annually [63]. Though they are less dependent on glycolysis than T. brucei there is evidence that stibogluconate, a well-known drug for treating leishmaniasis, specifically inhibits glycolysis in these parasites [64]. Therefore, we decided to study GAPDH of L. mexicana in parallel with the T. brucei enzyme. Its structure

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Page 376

was recently solved at 2.8 Å resolution in our lab [30]. The kinetic parameters of the two enzymes are virtually identical: Km (glyceraldehyde-3-phosphate) = 0.13 mM and Km (NAD+) = 0.41 mM for the L. mexicana enzyme [62]. As expected, the structures of the two parasite enzymes are very similar: the rms deviation for all backbone atoms is 0.7 Å and the adenosine binding environment is perfectly superimposable except for Asn39 of T. brucei GAPDH, which is a Ser in the L. mexicana enzyme. In this way drug design for one disease may have implications for another one. C. Phosphoglycerate Kinase (PGK) A monomeric enzyme, PGK transfers the acylated phosphoryl group from 1,3- bisphosphoglycerate to ADP, thus forming 3-phosphoglycerate and ATP. The enzyme uses a metal ion as a cofactor, namely Mg2+ [65]. The PGK enzyme from T. brucei has been overexpressed in E. coli [66]. Human PGK is not

Figure 8 Steroview of B.stearothermophilus PGK [69] in complex with ADP bound to the C-terminal domain. To illustrate the sugar binding site, the 3-phosphoglycerate has been added to the figure based on the crystal structure of pig muscle PGK [32].From the distance between the two substrates it is obvious that during catalysis a hinge-bending motion between the domains of the protein has to occur to bring the substrates together.

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Page 377 Table 3 Residues Involved in ADP Binding in Glycosomal and Human PGK T.brucei

Human_1a

Human_2b

In contact with ADP moiety

Ala 242

Gly 238

Gly 238

Adenine

Tyr 245

Phe 241

Tyr 241

Adenine

Lys 259

Leu 256

Leu 256

Adenine

Ala 314

Gly 309

Gly 309

Adenine

Ser 378

Thr 375

Thr 375

β-phosphate

aSomatic bPGK

PGK [67].

in spermatogenic cells [68].

commercially available. However, its sequence has been determined [67] and appears to be 97% identical to horse and pig PGK. For completeness it should be mentioned that there is a second human PGK in testis tissue that is 87% identical to the somatic enzyme [68]. The crystal structures of the apo-enzyme from horse [31] and of the binary complex between pig PGK and its substrate [32] (Figure 8) are available from the Protein Databank. The substrate was found to bind to the N-terminal domain of the enzyme. The binding site for ADP is known from the structure of its binary complex with PGK from B. stearothermophilus [69]. It resides in the Cterminal domain. Since the substrate and ADP binding sites are 10 Å apart, a hinge-bending motion between the two domains has been postulated to occur during catalysis [70]. Kinetically glycosomal PGK from T. brucei and mammalian PGK are very similar: the Km values for ATP are 0.29 and 0.46 and mM, respectively; the Km values for 3-phosphoglycerate are 1.62 and 0.62 mM, respectively (due to the unavailability of the human enzyme the rabbit muscle enzyme was used as a substitute) [71]. The residues responsible for binding the substrate [32] are identical between human and glycosomal PGK [72]. However, five of the residues involved in the binding of ADP differ between the two enzymes (Table 3). Apparently, the biggest difference between human PGK and the parasite enzyme is the charged residue Lys259, which has the apolar Leu256 as a human counterpart. Molecules that bind at the ADP binding site and specifically recognize Lys259 might therefore be good starting points for drug design. III. Search For New Leads A. Triosephosphate Isomerase: The Crystallographic Cocktail Soak Approach

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Because there are no known leads that bind to the selectivity region of TIM, the design of selective inhibitors is an exercise in de novo ligand design. We tried to

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design such molecules on the basis of the trypanosomal TIM structure by a linked-fragment approach [57]. In that strategy small building blocks are designed to be complementary to the targeted surface of a protein. Such fragments can then be synthesized or purchased, tested for their effect on enzyme kinetics and for their binding mode by crystallography. Promising fragments are then linked together into larger molecules. The idea behind this stepwise approach is to obtain early experimental feedback in the drugdesign cycle. The crystallographic follow-up of our linked-fragment approach design for trypanosomal TIM was disappointing. Two designed fragments were soaked into a crystal of trypanosomal TIM, namely 4hydroxy-2-butanone and D-asparagine. Despite high concentrations of these molecules in the mother liquor, 220mM and 30 mM respectively, no convincing electron density could be seen in difference Fourier maps calculated between 10.0-2.8 Å [72]. Common to both molecules is that they are fairly polar, rather flexible, and were expected to displace crystallographically observed water molecules. Apparently, de novo design of tightly binding small ligands is far from trivial. We also tried to find new leads by a completely experimental approach. For that purpose the crystallographic cocktail soak (CCS) approach was developed. In this method cocktails of fine chemicals are soaked into a crystal in the hope of finding crystallographic evidence of binding for one of the molecules from the cocktail. The identification of such a molecule might not be clear immediately because several molecules in the cocktail might be compatible with the shape of the electron density, especially if the resolution is not very high. An outcome would be provided by a dichotomic approach (Figure 9), in which the crystallographic soaking experiment is repeated with ever smaller subcocktails of the original one. For example, if a ligand shows up from a cocktail soak of 32 compounds, a second experiment should be done with only half of the compounds. If the ligand fails to show up, one knows that it is one of the alternative 16 compounds. After at most six experiments the identity of the ligand is known. One might like to think of the CCS approach as the experimental analog of the computational methods in programs like GRID [73] or MCSS [74], but with thirty-two compounds at a time.

Figure 9 Dichotomic search for unknown ligand from cocktail 1. The number of compounds in the sub cocktail is indicated by n, and the interpretation about the presence of compound X in the subcocktail is given as Y/?/N.

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For trypanosomal TIM we experimented with three different cocktails of 32 compounds (Table 4). Molecules were chosen in such a way that they would be compatible, soluble, cheap, and as varied as possible. Each compound was present at a concentration of 1 mM. The final cocktail solutions were clear and devoid of precipitate. Since this was a pilot experiment both subcocktails were checked at each stage of the dichotomic strategy. Only the soak with cocktail 1 revealed electron density that could not be accounted for by water molecules, hereafter called peak X. The soaks with cocktails 2 and 3 led to featureless difference Fourier maps. The quality of the data and refinement can be inspected from Table 5, while Figure 9 illustrates the dichotomic search to identify peak X. An oxidized molecule of DTT, identified in the high-resolution structure of the native TIM crystals [24], served as an internal reference to judge the quality of the data and the noise level in the final difference Fourier maps. Peak X was found near His95 of the second subunit of the enzyme, i.e., the subunit where the flexible loop adopts the closed conformation in this crystal form. Its signal was somewhat weaker than that of DTT. The same density showed up when crystals were soaked with subcocktail 1B but not with 1A, narrowing down the list of potential ligands to sixteen compounds. However, the next round of the dichotomic search led to a problem that has not been solved thus far. Peaks of roughly the same shape as the original peak X appeared with both subcocktails 1BA and 1BB. Several strategies were followed to improved the quality of the maps. First, the model was further refined with all data while a bulk solvent scattering correction [75] was incorporated. Second, a variety of maps were calculated: (|Fo|-|Fc|) eiαc, (2|Fo|-|Fc|) eiαc, (3|Fo|-2|Fc|) eiαc and (|Fo|-|Fo,native|)eiαc. Third, all maps were SIGMAA-weighted [76]. Since the shape of peak X varied substantially between the different maps it can be tentatively concluded that peak X did not originate from the presence of a compound but was noise. The lesson of this experiment seems to be that the crystallographic cocktail soaking approach should only be tried when high- resolution data can be obtained, probably better than 1.8 Å resolution. B. Glyceraldehyde-3-Phosphate Dehydrogenase: Docking In order to discover new ligands that would block GAPDH of T. brucei by occupying the adenosine binding region we used the program DOCK [77], version 3.5. This program characterizes a binding site by filling it with a set of overlapping spheres. The centers of these generated spheres constitute an irregular grid, called a “graph” by mathematicians. Docking of a ligand then consists of matching subsets of ligand interatomic distances onto subsets of the receptor graph. Finally, the quality of the fit between a docked ligand and the receptor is evaluated. Within DOCK 3.5 three methods are available for this evaluation: contact scoring, which measures shape complementarity; force-field scoring, which is an estimate of the enthalpy of the intermolecular interaction; and elec-

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Page 381 Table 5 Crystallographic Cocktail Soaking Experiments of Trypanosomal TIM Crystalsa res

DTT

X

C

a(Å)

b(Å)

c(Å)

m(°)

(Å)

R-sym

Compl

R

(σ)

(σ)

1

112.7(1)

97.7(1)

46.7(1)

0.7

2.85

0.044

0.97

0.136

5.0

3.5

1A

112.6(1)

97.6(1)

46.8(1)

0.9

2.50

0.099

0.96

0.173

7.0

2.0

1B

112.7(2)

97.5(3)

46.8(2)

0.7

2.30

0.044

0.88

0.172

5.0

4.0

1BA

112.6(2)

97.9(2)

46.7(2)

0.7

2.40

0.051

0.96

0.202

7.8

3.0

1BB

112.6(1)

97.8(1)

46.7(1)

0.7

2.30

0.034

0.90

0.203

5.0

3.0

2

112.9(1)

97.8(1)

46.7(1)

0.9

2.40

0.060

0.92

0.172

5.0

-

3

112.9(1)

97.7(3)

46.7(2)

0.7

2.40

0.057

0.93

0.170

4.0

-

aThe

following data are tabulated column-wise: C = cocktail; a, b, c = cell parameters of the P212121 crystals; m = mosaicity; res = resolution; R-sym = agreement between symmetry-related reflections; Compl = completeness; R = agreement between data and model; DTT = signal of oxidized DTT in the final difference Fourier map. DTT is present in the mother liquor but not incorporated in the model. X = signal of peak X in the final difference Fourier map. Maps were calculated with data between 10.0 Å and the high resolution limit.

trostatic scoring, where the linearized Poisson-Boltzmann equation is solved [78]. We report here on docking experiments to identify GAPDH inhibitors from the Available Chemicals Directory-3D 93 (ACD) [80]. Force-field scoring and electrostatic scoring require the assignment of partial atomic charges. Unfortunately, such charges are not available in the ACD, mainly because there is no consensus on a method to calculate them. Since the number of molecules in the ACD is very large, about 73,000 in 1993, we opted for the charge-equilibration algorithm developed by Rappé and Goddard [81] as implemented in the BIOGRAF program [82]. This method leads to charges that are in excellent agreement with experimental dipole moments and with atomic charges obtained from electrostatic potentials of accurate ab initio calculations. A script was written that processed the entire ACD automatically on an R4000 processor in about two days. Charges of the protein atoms were assigned from the table of AMBER-derived charges provided with DOCK 3.5.

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Because there are tens of parameters that can be varied in the program there is no such a thing as the DOCK run for a given protein target. We chose to perform two parallel runs that differ in receptor description. In run 1 the DOCK sphere center description was used while in run 2 the atomic coordinates of 2'- deoxy-2'-(3-methoxybenzamido)adenosine, a designed selective inhibitor of T. brucei GAPDH (see Section IV), were picked to describe the receptor site. For each run the same program parameters were chosen (Table 6) and the ACD database was split up in batches of 10,000 molecules. The computation was done on an Indigo2 workstation with an R4400 processor operating at 175 MHz. It took 3 h 19 min of CPU time for run 1 and 34 h 48 min for run 2. The ten-fold time difference between the two runs originates from the different number of

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Page 382 Table 6 Parameters Chosen for the DOCK Runs with T. brucei GAPDHa Program

Variable

SPHGEN

DISTMAP

aVariables

Value

Program

dentag

X

DOCK

dotlim

Variable

Value

distance_tolerance

1.0

0.0

nodes_maximum

4

radmax

5.0

nodes_minimum

4

radmin

1.0

ligand_binsize

0.4

polcon

2.3

ligand_overlap

0.1

ccon

2.8

receptor_binsize

0.8

discut

4.5

receptor_overlap

0.2

perang

3

atom_minimum

5

as defined in the DOCK 3.5 manual and discussed in References 79 and 83.

centers to describe the receptor, namely 20 for run 1 and 34 for run 2. According to theory [83], the difference should scale as 344/204 = 8.3, which fits our observation. Since it is well known that scores exhibit poor correlation with real affinities [84] we decided to subject the 200 best-scoring ligands of each batch to inspection on the graphics. Scanning through the 3200 compounds required about ten days. Eventually, sixteen compounds were selected for purchase on Table 7 Parasite GAPDH Inhibitors Discovered with DOCK Contact Inhibitor

IC50 (mM)

Delphi

Score

Rank

Score

Rank

Run 1 2-Guanidinobezimidazole

1.2

97

1432

1.01

1427

2-Benzimidazoylurea

1.8

102

958

0.27

1241

4-Nitrophenyl sulfone

2.8

106

608

-0.46

370

Tryptophan

6.0

128

32

-0.48

357

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5-Methoxytryptamine

19.0

92

1495

-0.32

488

Ephedrine

25.0

99

1315

-0.22

590

Epinephrine

>7a

102

989

-3.09

27

Aspartic acid dimethyl ester

>10

87

1551

-1.66

146

3-Amino-L-tyrosine

>11a

102

969

-0.14

703

1,3-Diphenylguanidine

>11a

112

301

-1.28

209

Octopamine

>50a

106

660

-2.03

83

4.4

132

900

0.11

1112

12.0

143

241

-2.02

156

Norepinephrine

>5.4a

151

84

-2.09

137

4'-Amino-N-methyl acetanilide

>7.8a

131

1027

0.04

1021

>9.3

142

300

0.13

1156

Run 2 2-Nitrobenzoic acid hydrazide Dopamine

L-histidinol aCould

not be tested at higher concentrations due to solubility problems.

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Page 383

Figure 10 Binding mode of 2-guanidinobenzimidazole to T.brucei GAPDH as predicted by DOCK. The benzimidazole moiety occupies roughly the position occupied by adenine in the holo-enzyme, whereas the guanidino group a salt bridge to Asp37.

the basis of structural rigidity, chemical stability, solubility, and electrostatic complementarity. The latter property was evaluated with the program DELPHI [85]. Each of the sixteen compounds was tested for GAPDH inhibition (Table 7). Half of them were inactive while the other ones showed inhibition in a range between 1.2 and 25 mM. Unfortunately, there appears to be no correlation between the DOCK scores and the IC50 values. For examples, norepinephrine and 1,3-diphenylguanidine are inactive while they have a better score than 2- guanidinobenzimidazole (Figure 10), the compound with the best IC50. Also, it appeared that the two different receptor descriptions used led to almost completely different lists of compounds. Only 156 molecules occurred in both lists of top-scoring molecules. In the modeled binding mode all of the inhibitors occupy roughly the same position as the purine ring of NAD in the crystal structure of GAPDH. While the values obtained for IC50 are indicative of poor inhibition, one has to keep in mind that adenosine exhibits an IC50 of 50 mM [13]. By using the program DOCK we were able to discover ligands that have a substantially higher affinity for GAPDH than the natural ligand. C. Phosphoglycerate Kinase: Leads from the Past The starting point for drug design in the case of PGK is quite different from TIM or GAPDH because a number of nonsubstrate-like inhibitors have been

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Figure 11 Two-dimensional structure of SPADNS, a micromolar inhibitor of T.brucei PGK.

discussed in the literature. Suramin inhibits glycosomal PGK of T. brucei with a Ki of 8.0 µM [69]. In addition a number of yeast PGK inhibitors are known: gallic acid with a Ki = 0.4 mM [86], hydroxyethylidene biphosphate with a Ki = 24 mM [87]. 1,3,6-naphthalenetrisulphonic acid with a Ki = 5.5 mM, and 2-(p- sulphophenylazo)-1,8-dihydroxy-3,6-disulphonic acid, also known as SPADNS (Figure 11), with a Ki = 126 µM [87]. None of the four yeast PGK inhibitors are potent, but, for SPADNS, the binding mode has been further characterized. Studies by Williams et al. [87] have demonstrated that SPADNS is directly competitive with both enzyme substrates, 3-phosphoglycerate and ATP. Moreover, by 600 MHz 1H-NMR it was shown that SPADNS interacts with the nucleotide binding site while the conformation of the enzyme changes substantially [87]. Since the four yeast PGK inhibitors are commercially available it was logical to test them for T. brucei PGK inhibition. The first three compounds were active in the millimolar range. However, SPADNS exhibited a Ki of 10.0 µM in these in preliminary tests [88]. Moreover, when assayed against a commercially available rabbit muscle PGK, SPADNS had no influence on the enzyme kinetics up to a concentration of 250 µM [88]. In conclusion, SPADNS appears to be an excellent lead because of its potency and selectivity. Crystallographic experiments to determine its binding mode to T. brucei PGK are underway. IV. Lead Optimization: Glycosomal Gapdh From the selectivity point of view the adenosine binding site of GAPDH is attractive for drug design, as we explained in Section II.B. Unfortunately, inhibition studies on T. brucei and L. mexicana GAPDH revealed the poor affinity of our natural lead adenosine with IC50 values of 100 mM and 50 mM, respectively. Moreover, adenosine is an “antiselective” lead because the IC50

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for human GAPDH is better than for the parasite enzymes, namely 35 mM [13]. Despite the fact that these IC50 values are about ten thousand times higher than what would be considered a lead in the pharmaceutical industry we decided to optimize the affinity and selectivity of adenosine. Each of the three areas where differences occur between the parasite and the human enzyme are hydrophobic. Therefore, we modeled hydrophobic substituents at positions C2, C8, and O2' of adenosine under the constraint that they were conformationally compatible with the C2'-endo pucker of the ribose sugar. Designing derivatives at O2' was a problem, however. Each of the two ribosyl hydroxyls forms a hydrogen bond with the carboxylate of Asp37. Since making direct derivatives of the hydroxyl, such as ethers or esters, would deprive the Asp of a hydrogen-bond partner while burying the carboxylate, resulting molecules would have a dramatically reduced affinity. Moreover, an alignment of 47 GAPDH sequences made it clear that the Asp is highly conserved [89]. An elegant way to overcome this problem was to replace the 2' -hydroxyl by a 2'- amino function. Moreover, coupling with carboxylic acids was appealing from a synthetic point of view while the conformational properties of the amido-substituted system would ensure the correct orientation of substituents into the selectivity cleft. The modeled inhibitors were evaluated for the quality of their fit to the protein surface and subsequently synthesized. From Table 8 it can be seen that our predictions were successful. The addition of a methyl group at C2 of the adenine ring, which is close to Val36, increased the affinity for parasite GAPDH by an order of magnitude. The effect of a thienyl substitution on C8, targeted to Leu112, was even bigger, namely two orders of magnitude. However, both substitutions are only mildly selective (Table 8). As expected, the greatest gain in selectivity was obtained by modifying the 2'-position of the ribose, so that the selectivity cleft is filled up (Figure 12). The 2'-deoxy-2'-(3-methoxybenzamido) adenosine compound (Figure 13) bound at least 48 times better to L. mexicana GAPDH than to the human enzyme. The selectivity versus T. brucei GAPDH appeared to be smaller. This has to be ascribed to a difference in residues contacting the 3-methoxy moeity. The residue Asn39 of T. brucei GAPDH has a Ser equivalent in the L. mexicana Table 8 Inhibition Gains of Designed GAPDH Inhibitors with Respect to Adenosine C2-subst

C8-subst

CH3

H

H

T. brucei

L. mexicana

human

OH

12.5

6.25

<3.5a

thien-2-yl

OH

167

100

27

H

H

NHCO-phenyl

16.7

NDb

3.5

H

H

NHCO-(3-OCH3-phenyl)

45

167

<3.5a

aUpper bNot

C2'-subst

limit because solubility problems did not allow for an IC 50 determination.

determined

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Figure 12 Predicted binding mode of 2' -deoxy-2'-(3-methoxybenzamido)adenosine to T.brucei GAPDH. (From Ref. 13. Copyright 1994 by the American Chemical Society.)

enzyme. In conclusion, the strategy of burying hydrophobic residues with lipophilic substituents paid off. Despite the rather poor IC50 values of our optimized inhibitors, an evaluation of their effect on live trypanosomes appeared to be useful. Enzyme inhibitors that are not taken up by the parasites would be of no use as a drug. Therefore, the effect of 2-methyl-adenosine, 8-(thien-2-yl)-adenosine and 2'- deoxy2'-(3-methoxybenzamido)adenosine on the growth of T. brucei in cultures, as described by Baltz et al. [90], was monitored. At 0.1 mM all compounds inhibited the growth completely, unlike adenosine derivatives that were without inhibitory effect against T. brucei GAPDH [91]. Experiments are underway to confirm that the growth inhibition is due to blockage of the glycolytic pathway. Also, te mechanism of uptake of te inhibitors will be examined because it is now well established that trypanosomes possess a unique P2 purine transporter that they use for uptake of purines from the host [15]. The experimental antitrypanosomal drug 5'-{[(Z)-4-amino-2-butenyl}methylamino}-5'deoxyadenosine(MDL 73811) Figure 13), which is an irreversible S- adenosyl-L-methionine decarboxylase inhibitor, is actively taken up through the P2 transporter. Moreover, MDL 73811 is not actively transported in the human host, which presumably contributes to the drug's selectivity [19]. It is not unthinkable that our inhibitors might use the same transporter because of the nature of their scaffold, adenosine.

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Figure 13 Two-dimensional structures of 2'-deoxy-2'-(3-methoxybenzamido) adenosine (MMBA), a selective T.brucei GAPDH inhibitor and MDL 73811, an irreversible inhibitor of trypanosomal S-adenosyl methionine decarboxylase.

Finally, we want to point out that pessimism about adenosine derivatives as drugs is not necessarily warranted. This doubt stems from the argument that many proteins recognize NAD(P), adenosine, or ATP. Cross-reactivity of adenosine with these different proteins and, therefore, toxicity may be expected. That this is not necessarily so is evident from the use of adenosine derivatives as antileukemia agents. For example, fludarabine, a C2'-epimer of adenosine, exhibits relatively low toxicity [92]. The much bigger changes to the adenosine scaffold in our inhibitors may hence lead to a surprisingly high overall selectivity. V. Conclusions Our goal is to discover and design selective inhibitors of trypanosomal glycolsysis. Thusfar, three enzymes have been targeted. Whereas little success was obtained with TIM, substantial progress is being made with GAPDH and PGK. Obstacles encountered during this project were the need for selective inhibitors and the absence of potent inhibitors as lead compounds. From our studies to inhibit TIM it appears that it is difficult to come up with inhibitors for areas on the protein surface for which no known inhibitors exist. On the other hand, lead optimization for GAPDH by over two orders of magnitude in just one cycle of drug design was straightforward. Selectivity was obtained by using part of the cofactor as a lead and exploiting the hydrophobic patches at the surface of the parasite enzyme. In particular, 2'-deoxy-2' (3methoxy-benzamido) adehttp://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_387.html (1 of 2) [4/5/2004 5:42:15 PM]

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nosine proved to inhibit the parasite more than fifty times better than the human enzyme. Additionally, by means of the program DOCK, eight new leads for GAPDH inhibition were found. None of them were micromolar inhibitors but all of them were more potent than the natural lead, adenosine. For trypanosomal PGK a potent lead compound, SPADNS, was discovered by testing inhibitors that had been described as weak inhibitors of yeast PGK. Moreover, this lead had no effect on mammalian PGK at concentrations up to twenty-five times higher than that needed for T. brucei inhibition. At present, none of our inhibitors is potent enough to consider clinical tests. There is hope, however, since our GAPDH inhibitors inhibit the growth of trypanosomes in cultures. Acknowledgments It is a pleasure to thank the many colleagues and collaborators who have contributed to this project: Paul Michels, Veronique Hannaert, Sylvie Allert, and Linda Kohl (Institute for Cellular Pathology in Brussels) for cloning and overexpressing trypanosomatid enzymes; Phil Petra (University of Washington, Seattle) for helping us out with protein purification protocols; Mia Callens and Fred Opperdoes (Institute for Cellular Pathology in Brussels) for enzymology and parasitology; Rik Wierenga, Martin Noble, Fred Vellieux, Randy Read, Risto Lapatto, Hillie Groendijk, Tjaard Pijning, and Kor Kalk for laying the structural foundation of the project in Groningen and Heidelberg; Cees Witmans and the late Alan Horn (University of Groningen), Michèle Willson and Jacques Perié (University of Toulouse), Serge Van Calenbergh, Arthur Van Aerschot, and Piet Herdewijn (University of Leuven) for synthesizing TIM and GAPDH inhibitors; Kim Simons (University of Washington) for going after completely new GAPDH inhibitors; Véronique Mainfroid and Joseph Martial (University of Liège) for providing human TIM; Klaus Muml;uller and Klaus Gubernator (Hoffmann-La Roche, Basel) for valued modeling advice; and Mike Gelb (University of Washington) for valued discussions. Financial support for these investigations has been provided by the World Health Organization, Hoffmann-LaRoche in Basel, the Dutch Organization for the Advancement of Science (NWO), the STD program of the European Community, the School of Medicine of the University of Washington, and the Murdock Charitable Trust. Note Added in Proof We just solved the ternary structure of PGK from Trypanosoma brucei in complex with ADP and 3phosphoglycerate. The enzyme is in the closed conformation that has eluded crystallographers for 20 years.

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90. Baltz T, Baltz D, Giroud C, Crockett J. Cultivation in a semi-defined medium of animal infective forms of Trypanosoma brucei, T. equiperdum, T. evansi, T. rhodesiense and T. gambiense. EMBO J 1985; 4:1273–1277. 91. Opperdoes FR, personal communication. 92. Whelan JS, Davis CL, Rule S, Ransom M, Smith OP, Metha AB, Catovsky D, Rohatiner AZ, Lister TA. Fludarabine phosphate for the treatment of low grade lymphoid malignancy. Br J Cancer 1991; 64:120–123. 93. Kraulis PJ. MOLSCRIPT: a program to produce both detailed and schematic plots of protein structures. J Appl Crystallography 1991; 24:946–950. 94. Bernstein BE, Michels PAM, Hol WGJ. Synergistic effects of substrate-induced conformational changes in phosphoglycerate kinase activation. Nature 1997; 385:275–278. 95. Michels PAM. Biology of the Cell 1988; 64:157–164.

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16 Progress in the Design of Immunomodulators Based on the Structure of Interleukin-1 Glen Spraggon University of California, San Diego, La Jolla, California Pandi Veerapandian Axiom Biotechnologies, Inc., San Diego, California, and La Jolla Institute for Experimental Medicine, La Jolla, California I. Introduction The interleukin-1 (IL-1) family of cytokines exhibit both normal and pathological effects in almost every tissue and organ system and, as such, have been associated with cells engaged in the immune response, inflammatory cells, and cells engaged in development, differentiation, and repair processes [1–4]. Interleukin-1 can produce either a direct response on one specific target cell or act as an indirect effector molecule, inducing the expression of a variety of genes and synthesis of several proteins such as IL-2–IL-8, tumor necrosis factor (TNF), colony-stimulating factors (CSF), platelet-derived factors (PDFs), and other cytokines. Pathologically uncontrolled, IL-1 activity can induce disease states and has been linked to septic shock, the growth of acute and chronic myelogenous leukemia cells, inflammation associated with arthritis and colitis, development of atherosclerotic plaques, insulin-dependent diabetes, osteoporosis, parsitemia, and cancer [1,5,6]. Strategies to treat such diseases are being developed and usually involve the inhibition of IL-1 synthesis or the blocking of its activity [7]. The therapeutic advantage of reducing the activity of IL-1 resides in preventing its deleterious biological effects without interfering with homeostasis. In order to achieve such a goal, an understanding of the structure-function relationship of the IL-1 family of proteins at the molecular level is important. Such a knowledge could lead to the design and development of

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synthetic agents with the ability to modulate IL-1 responses. Here we review the present knowledge of the IL-1 system and the attempts toward the development of its modulators. A. Cytokines and Physiological Responses Cytokines are multifunctional hormones produced by a variety of cells to carry out a wide range of overlapping biological actions including communication within the immune system and between other cell types. They are produced by macrophages, endothelial cells, fibroblasts, keratinocytes, T cells, B cells, and natural killer cells in response to injury and infection [7]. Upon production they act either locally or as systemic intercellular signaling factors and induce the production of other cytokines for continuing their communications via cell surface receptors. The cytokines network also has associated with it a complementary set of soluble or membrane-bound antagonist or mediator molecules that are capable of shutting down these effects [8]. During infection and injury, the leukocytes adhere themselves to the endothelium and migrate into tissues. Cells like T-lymphocytes, monocytes, macrophages, and neutrophils collect themselves as inflammatory infiltrate. Once within the tissues, these cells neutralize the microorganisms or infected cells, secreting other cytokines that can modulate the adhesion molecule expression on the endothelium. In addition, chemotactic signals are delivered to cells passing in the circulation. Cytokines are also involved in tissue repair, i.e., remodeling of connective tissues and revascularization of damaged areas. Soluble mediators are released from the site of damage into the circulation to act in an endocrine fashion. The cytokines then control hepatic responses to tissue damage. During the regulation of hematopoiesis, cytokines induce the production and release of a number of colony-stimulating factors and other interleukins. Thus induced factors are responsible for the replacement of leukocytes and erythrocytes that were lost after trauma. A large number of cytokine molecules have been characterized [7]. Structurally it appears that these cytokines can be classified into the following main groups based on their folding pattern: 4-helix bundles, beta-trefoil, beta sandwich, EGF-like, beta cysteine knot, and alpha/beta. Thus far the structures of many representatives of each family have been solved by both x-ray crystal- lography and NMR (Table 1). Extensive experimentation has indicated that despite a large degree of structural diversity, there is a large degeneracy in the cytokine network and the molecules have a wide range of overlapping biological functions. For example, IL-1, TNF, PDGF, and TGFB all exhibit similar biological behavior. The structures of the receptors for these molecules have not been forthcoming and only three examples of the extracellular domains of the receptors have been reported. They are growth hormone receptor in complex

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Page 397 Table 1 Known Three-Dimensional Structures of Cytokines

Molecule

Brookhaven

Structure

Code

Determined by

Type

Interleukin-1 alpha (IL-1α)

Beta trefoil

2ILA

X-ray crystallography

Interleukin-1 beta (IL-1β)

Beta trefoil

1ILB, 2ILB, 4ILB, 5ILB, 6ILB

X-ray crystallography and NMR

Interleukin-1 receptor antagonist (IL-1ra)

Beta trefoil

1ILR, 1IRP

X-ray crystallography and NMR

Fibroblast growth factor (acidic) (aFGF)

Beta trefoil

1AFC

X-ray crystallography

Fibroblast growth factor (basic) (bFGF)

Beta trefoil

1BFG

X-ray crystallography

Granulocyte-colony stimulating factor (G-CSF)

Long-Chain 4 helix bundle

1RHG, 1GNC

X-ray crystallography

Growth hormone

Long-Chain 4 helix bundle

1HGU

X-ray crystallography

Leukemia inhibitory factor (LIF)

Long-Chain 4 helix bundle

1LKI

X-ray crystallography

Granulocyte-macrophage colony stimulating factor (GM-CSF)

Short-Chain 4 helix bundle

1CSG, 1GMF

X-ray crystallography

Interleukin-2 (IL-2)

Short-Chain 4 helix bundle

3INK

X-ray crystallography

Interleukin 4 (IL-4)

Short-Chain 4 helix bundle

1BBN, 1ITM, 2CYK

NMR

Interferon gamma (IFN-γ)

Dimeric 4 helix bundle

1IKI

X-ray crystallography

Interleukin 10 (IL-10)

Dimeric 4 helix bundle

1ILK

X-ray crystallography

Nerve growth factor (NGF)

Beta cysteine knot

1BET, 1BTG

X-ray crystallography

Platelet derived growth factor (PDGF)

Beta cysteine knot

1PDG

X-ray crystallography

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Beta cysteine knot

1TFG, 2TGI

X-ray crystallography

Transforming growth factor alpha (TGFA)

Beta EGF-like

2TGF

NMR

Tumour necrosis factor (TNF)

Beta sandwich

1TNF

X-ray crystallography

Macrophage inflammatory protein 1-beta

Alpha/Beta

1HUM

NMR

Interleukin 8 (IL-8)

Alpha/Beta

1IL8

NMR

Melanoma growth stimulating activator (MGSA)

Alpha/Beta

1MGG

NMR

Platelet factor 4

Alpha/Beta

1RHP

X-ray crystallography

Transforming growth factor beta2 (TGFB2)

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with growth hormone, prolactin receptor (PRLR), and tumor necrosis factor with a ligand. In the case of the human growth hormone-receptor complex and tumor necrosis factor-receptor complex the interaction between the ligand and the receptor is intricate and takes place over a large area. It is likely that this will be a characteristic feature of all cytokine-receptor complexes. B. Cytokine-Based Therapy Certain disease states can occur due to a disregulation of the cytokine network. This can lead to chronic stimulation of the immune and inflammatory response and ultimately to disease. Many cytokines have been implicated in autoimmune diseases like myasthenia gravis, insulin-dependent diabetes, atherosclerosis, systemic lupus erythematosus, and rheumatoid arthritis [1,7,9]. The intimate relationship between cytokines and the pathology of disease development and progress can be exploited to provide therapeutic benefit. By subtly manipulating the cytokine communication network one can modulate the disease process. Understanding of the functional roles of cytokines that mediate the communication between them and the factors involved in the immune system's cell-cell interactions forms the foundation for cytokine therapies. Such therapeutic agents are now a possibility due to modern techniques such as molecular biology, structural biology, high-power computation, combinatorial chemistry, and functional screening. The findings from biological techniques in conjunction with the structural insights as obtained through protein crystallography and nuclear magnetic resonance form the foundation for rational drug design [10–13]. Discovery of IL-1-based therapeutic agents have led to promising applications in the treatment of the above mentioned diseases. Design of synthetic adjuvants, for exogenous immunomodulation, is also an important factor to be considered in the field of vaccine development. In this review we will discuss the available literature on interleukin-1 and the recent attempts toward the design of immunomodulators. II. The Interleukin-1 Family The three molecules of the IL-1 family, interleukin-1α (IL-1α), interleukin-1β (IL-1β), and interleukin-1 receptor antagonist (IL-1Ra) map to the long arm of chromosome two in the human genome. It appears that the family arose via a gene duplication event some 350 million years ago, and the molecules possess between 27.5 and 36% sequence identity with each other (Table 2) [1,14,15]. In addition, the genes for the two IL-1 receptors IL-1R1 and IL-1RII [16,17], and an IL-1R accessory protein (IL-1RacP), which binds to the IL-1, IL-1 receptor complex [18], have been identified. Together, these molecules via their differential activity serve primarily to modulate the host defense mechanism.

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Page 399 Table 2 RMS Distance of Aligned Cα Moieties of the Three Interleukin-1 Structures IL-1β

IL-1Ra

IL-1α

IL-1β

0.0

1.8 Å (36.3%)

1.42 Å (30.7%)

IL-1α

1.8 Å (36.3%)

0.0

1.8 Å (27.5%)

IL-1Ra

1.42 Å (30.7%)

1.8 Å (27.5%)

0.0

Number in parentheses referred to sequence identities between the aligned structures.

A. Expression and Processing of IL-1 All three IL-1 molecules, IL-1α, IL-1β, and IL-1Ra are synthesized as 31 kDa precursor molecules produced primarily by mononuclear phagocytes. These precursor proteins can be subsequently processed to mature 17 kDa molecules. The means whereby this is achieved appears to be different for each type of molecule and may help to explain the roles of the three. Both agonist molecules IL-1α and IL-1β lack a classical hydrophobic leader sequence and thus must be processed in an alternative way. The two can be separated by their biological activity in the precursor form, IL-1α being active in both precursor and mature forms while IL-1β produces a biological response only in its processed form. In addition, a specific enzyme, Interleukin-1 beta converting enzyme (ICE), cleaves IL-1β to its mature form. This enzyme is a cysteine protease whose only known substrate is proIL-1β, which it cleaves at Ala 117 [19,20]. The means whereby pro IL-1α is processed is still largely a mystery although it has been postulated that a calpain or related protein may perform the task. Naturally occurring IL-1Ra is a 22 kDa glycosylated protein [14,15,21– 24] that possesses a signal sequence. It is likely that it is processed in the conventional manner as can be concluded by the glycosylation states of the molecule not present in the agonists [25]. The LPS-stimulated human blood monocytes initially express the gene for IL-1Ra [25]. An alternatively spliced form of IL-1Ra also exists (intracellular IL-1Ra), which remains inside the cell presumably to block intracellular IL-1 action [26]. Both soluble and intracellular forms of IL-1Ra block IL-1R but do not trigger any biological response. B. Receptors and Responses

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All effects of the IL-1 family are exerted via their receptors, which are present on a variety of different cell types. There are two known IL-1 receptors. The first, type I (IL-1RI) is an 80-kDa formation found mainly on T cells and fibroblasts; the second, type II (IL-1RII) is present on B cells, monocytes, neutrophils, and heptoma cells and is approximately 60 kDa in size. From sequence analysis these molecules are believed to belong to the Ig-superfamily, the extracellular portion of the receptors consisting of three immunoglobulin-like domains of approximately 100 residues each [27,28]. The IL1RI receptor

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Figure 1 Schematic representations of the members of the IL-1 family.

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has a single membrane-spanning region and a cytoplasmic region of 213 amino acids. The IL-1RII receptor has a single membrane-spanning region and a cytoplasmic region of 29 amino acids. All three IL-1 molecules bind with a high affinity to the receptor IL-1RI. However only IL-1α and IL1β produce a detectable response [29]. The IL-1Ra molecule completely blocks the binding of the former two molecules without inducing any signal transduction events. The second IL-1 receptor, IL1RII, similarly binds IL-1 tightly but does not produce any signal [30]. The soluble portion of IL-1RI binds to IL-1Ra, IL-1α, and IL-1β in decreasing order of affinity whereas the soluble portion of IL-1RII binds IL-1β, proIL-1β, IL-1α, and IL-1Ra, in that order. It therefore appears that both IL-1Ra and IL1RII take part in modulation of the agonist molecules, an argument supported by the finding that soluble IL-1RII binds IL-1β with a similar affinity to the cell-associated receptor; whereas, the affinity of IL1Ra to such a soluble receptor is some 2000 times less [31,32,33]. Greenfeder, et al., [18] have identified a new molecule, the IL-1 accessory protein IL-1RacP. The complex of IL-1 and IL-1 receptor binds to IL-1RacP and together they induce the signal [18]. Greenfeder also observed that antibodies to IL-1RI and to IL-1RacP block IL-1 binding and activity. From sequence analysis it appears that the cytoplasmic domains of IL-1R and IL-1RacP contain the same amino acid domains commonly found in the members of the GTPase family of proteins [34]. It has been proposed that such a complexation may lead to a closer proximity of these cytoplasmic domains and thus facilitate signal transduction. A scheme showing functional relationships between molecules that make up the IL-1 system—IL-1α, IL-1β, IL1Ra, IL-1RI, IL-1RII, and related receptors—is shown in Figure 1. C. Autoantibodies of IL-1 In addition to these molecules, naturally occurring neutralizing autoantibodies of IgG type to IL-1α have been identified. These have been detected in serum isolated from human donors. [35,36]. These antibodies bind to both proIL-1α and 17-kDa IL-1α [37] and completely prevent the binding of IL-1α to type-I cell surface receptors [38]. Patients with autoimmune diseases have higher populations of these antibodies [39]. III. Three-Dimensional Structural Information In an effort to understand the natural and interactions of the IL-1 family, various laboratories have undertaken the task of elucidating the three-dimensional structure of the molecules. To the present this has resulted in the structures of all three members of the IL-1 family being solved independently by both x-ray

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crystallography and heteronuclear NMR spectroscopy [40–49]. In addition to these, the structure of IL1β converting enzyme has been determined [50]. The molecule is an oligomeric cysteine protease consisting of 10- and 20-kDa subunits. To date proIL-1β is its only physiological substrate. As processing is essential for IL-1β extracellular transport and activity, inhibitors of ICE could provide a method to block the production of the active form of IL-1β. Although the elucidation of these structures have been important in determining various differences among the individual proteins, perhaps a more important result would be the elucidation of the structures of the two interleukin-1 receptor molecules, either individually or in complex with their substrates. This information could provide a means to draw together the present structural and biochemical knowledge into a coherent picture of the agonistic activity of IL-1α and IL-1β as opposed to the antagonist effect of IL-1Ra. The structures should also provide a foundation upon which structure-based drug design could proceed. To date no crystals or structural reports for the receptors have been published. A. The β-Trefoil Fold Structurally IL-1 exhibits a unique fold, known as a β-trefoil fold. Each IL-1 molecule show the characteristic β-trefoil fold and small deviations of the back-bone despite the relatively low sequence identity (Table 2). Figure 2 shows the structural alignment of the IL-1 molecules. The β-trefoil fold was first observed in Kunitz-type soybean trypsin inhibitor [51]. The fold has since been identified numerous times in the Kunitz family of protease inhibitors, the interleukin-1 system molecules, and the acidic and basic fibroblast growth factors [52]. Although these proteins have diverse biological function and low sequence identity to each other, all appear to have a common structural core. Each one, however, binds to its specific receptor with high affinity. The present known examples of proteins with such a fold are composed of between 125 and 170 amino acids. The overall fold itself is an antiparallel β barrel consisting of six two-stranded hairpins. Three of these form a barrel structure (strands denoted β1, β4, β5, β8, β9, and β12) while the other three are in a triangular array that caps the barrel. The arrangement of these moieties is such as to give the molecule a pseudo-three-fold axis [52]. Each fragment contributes one pair of antiparallel beta strands to the barrel and one pair to the cap of the barrel, thus forming a so called “open” and “closed” end to the structure [42]. B. Three-Dimensional Structure of IL-1β The structure of IL-1β was the first of the IL-1 family to be solved and has been solved independently five times: four times by x-ray crystallography [40–

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Figure 2 Structural alignment of IL-1α, IL-1β, and IL-1Ra. Residue numbering is taken from IL-1β. Residues bordered in black are conserved over the three molecules while those in gray constitute a conservative substitution. Arrows indicate sheet region, and the cylinders indicate helix. Figure produced by Stamp [109] and Alscript [110].

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Figure3 (a) Stereo diagram of the secondary structural elements of IL-1β. Produced by Molscript [106]. (b) Stereo plot of all the atoms of IL-1β viewed parallel to the axis of the barrel.

42,44], and once by NMR [43]. The four crystal structures, each to 2.0 Å, were all solved in the same space group, P43, each structure being in relatively good agreement with the others [53]. As pointed out previously, the molecule adopts a β-trefoil fold with about 65% of the molecule in beta sheet and 35% in random coil/turn (Figure 3a). The IL-1β molecule resembles a conical barrel with a shallow open face on one end and a closed face on the other. The length of the long tubular core of the molecule is about 23 Å. Twelve antiparallel β

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strands constitute the secondary structural elements of IL-1β. Three pairs of β strands (one pair from each of the fragments) form the six-stranded barrel and the other three pairs cover one end of the barrel, referred to as the “closed end”. The amino and carboxy termini are close to each other at the “open end” of the barrel (Figure 3b). The overall structure of the molecule consists of three similar fragments (F1, F2, F3) related by a pseudo-3-fold symmetry, with each fragment having a βββLβ motif. Residues 1 to 52 (F1), 53 to 107 (F2), and 108 to 153 (F3) form these fragments. These βββLβ motifs can be superposed to display structural similarities. The core of the molecule consists entirely of hydrophobic residues, two-thirds of which are leucines and phenylalanines—a feature that seems to be essential to maintain the structural integrity of the molecule. Most of the aromatic groups have their planes aligned along the barrel axis and both ends of the barrel have concentrations of exposed polar residues that may be involved in binding interactions with the receptor (see below). No alpha-helical structure is observed in IL-1β although one short region of 310 helix is observed between residues Gln34 and Gln38. Two of the β hairpins are in the open end and three are at the closed end. Residues 18 to 28, 69 to 82, and 122 to 135 form the three β hairpins at the closed end where three strands (residues 24–28, 78–82, and 129–135) are in close proximity and form three sides of a triangle, covering this end of the barrel. C. Open End of the Barrel Analysis of the surface of IL-1 reveals an epitope consisting of many polar residues widely spaced, approximately in an annular fashion, around the open end of the barrel (Figure 3b). It has a broad surface area containing many polar residues [42]. Residues from six β-turns [type I, β turn 1 (T1=1117); type III, β turn 3 (T3=33-36); type I', β turn 6 (T6=52-55); type I, β turn 7 (T7=62-65); type I, β turn 9 (T9=86-89); type I', β turn 10 (T10=106-109)] are present in this open end. There is a β bulge between strands β4 and β5 in one of the hairpins; this bulge may play a key role in the formation of the putative binding surface. The β bulge in IL-1 is of the “wide” type stabilized by multiple interactions, whose conformation is such as to place the side chains of polar residues (Glu51, Asn53, Asp54) in the proposed binding surface, fanning out in the direction of the open end of the barrel. Multiple interactions serve to stabilize the strained conformation of the loop, between residue 86 and 94, presenting the side chains of Asp86, Lys88, Asn89, and Lys93 at the open end of the barrel. In well-defined residues having abnormal φ, ψ values, strained conformations often may be related to functional properties and are observed either in the active site or in the region responsible for its activity. The reason for having such strains in the loop at this open end may be to place these charged side chains in the putative receptor binding surface. There is a short 310-helix in the region between β3 and

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β4 (residue Gly33 and Gln39). By such a conformation, residues Gln32, Gln34, and Glu37 have their side chains in the proposed binding surface. A β turn between 62 and 65 is folded in such a way that the charged groups of the residues Lys63, Glu64, and Lys65 present themselves on the proposed binding surface. Conformational arrangement via saltbridge scaffolds and other multiple hydrogen-bonding interactions stabilize the residue 102 to 113 loop, and orient the side chains of Lys103, Glu105, Asn107, and Asn108 in the proposed surface at the open end of the barrel. Similar arrangement places the side chains Arg11 and Gln15 in the proposed epitope. The amino and carboxy termini form a part of the proposed epitope, contributing the polar side chains of Arg4, Ser5, Asn7, Gln149, Ser152, and Ser153. Based on extensive analysis, we found that the structural elements like electrostatic and hydrogenbonding interactions in the loops between strands allow the polypeptide to adopt a conformation that enables an unusual concentration of polar and charged groups to be presented at the open end of the barrel. Such a cluster of charged residues around an area that is almost perpendicular to the barrel axis forms a hydrophilic patch with which IL-1 might bind to the receptor [42]. The core of the barrel, though probably not involved in binding, must nevertheless be important to the function of IL-1 because mutations within it reduce activity but not binding. We hypothesized that binding and cell proliferation through signal transduction involve separate regions of the IL-1 molecule; the surface polar loops are required for the binding and the core of the barrel is required for the physiological response. D. Three-Dimensional Structure of IL-1α The structure of IL-1α has been determined by x-ray crystallography [45] to a resolution of 2.7 Å in space group P21. Its general fold is very similar to that of IL-1β, having the same central β barrel along with the adjoining loops (Figure 4a, b). The overall rms distance between 133 aligned Cα positions of IL1β and IL-1α is 1.8 Å (Figure 5). The major difference between the two molecules is an N-terminal extension of 8 residues beyond the N-terminus of IL-1β. This projection forms a short β strand (residues 6-10), the presence of which positions the N-terminus of IL-1α in an alternative conformation. It has been postulated that this conformation is responsible for the differences in binding of precursor IL-1α and β: while precursor and mature IL-1α bind to IL-1RI and elicit response, only processed IL-1β binds with the receptor. This suggests that the N-terminal region of IL-1 probably plays a role in receptor binding. Extra residues in the alternate conformation of immature IL-1β serve to inhibit the receptor binding and thereby the biological activity. In contrast to IL-1β, IL-1α incorporates two other secondary structural elements: a short strand (residues 97–99) and about two turns of a 310 helix (residues 101–105), neither have been shown to be important in the function of the molecule.

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Figure 4 (a) Stereo diagram of the secondary structural elements of IL-1α. Produced by Molscript [106]. (b) Stereo plot of all the atoms of IL-1α viewed parallel to the axis of the barrel.

E. Three-Dimensional Structure of IL-1Ra The x-ray structure of IL-1Ra was first reported in 1994 [47] and has since been solved by x-ray crystallography and NMR by several different laboratories [46–49]. Again, this molecule possesses the characteristic trefoil fold (Figure 6a) and similar tertiary structures (Figure 6b). The structure is similar to that of IL-1α and IL-1β with an rms deviation of 1.8 and 1.42 Å, respectively, when considering their Cα positions (Figure 7a). Structural superposition of these three molecules shows a common trefoil core (Figure 7b). In the absence of receptor substrate complexes, the comparison of all three structures combined with mutational studies (see below) is invaluable in obtaining a model of the regions

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Figure5 Superposition of the Cα atoms of IL-1α and β. Thinner line represents IL-1α and the thicker line is that of IL-1β.

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Figure 6 (a) Stereo diagram of the secondary structural elements of IL-1Ra. Produced by Molscript [106]. (b) Stereo plot of all the atoms of IL-1Ra viewed parallel to the axis of the barrel.

involved in binding and exertion of biological effect in the IL-1 system. Similarity between the three molecules has been observed mainly in the β strands. One notable region of

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difference between IL-1Ra and IL-1 agonists has been observed on the side of the beta barrel on site B of the receptor binding site (see below). The Cα positions in this region (residues 84–94 in loop β7–β8) differ by 9.1 Å [26] in comparison with IL-1β. This region also contains Asn84, which is the site for at least two forms of N-linked glycosylation that occur in IL-1Ra in addition to the nonglycosylated form. Mutagenesis experiments have also

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Figure 7 (a) Superposition of the Cα atoms of IL-1α, IL-1β, and IL-1Ra. (b) A common trefoil core based on the structural superposition of all the three IL-1 molecules.

identified this region and the surrounding area in IL-1β with a large receptor binding epitope encompassing the N and C termini of the protein as well as loops β4–β5 and β9–β10. The other postulated binding epitope (epitope A) is structurally conserved in IL-1Ra. Other differences in structure not related to any implicated biological structure are an extended 310 helix in residues 92–99 in IL-1Ra as opposed to a type-1 β turn found in IL-1β. F. Three-Dimensional Structure of Interleukin-1 Beta Converting Enzyme (ICE) As mentioned above, the processing of the different members of the IL-1 family is important to discovering their function and may also provide clues in the

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design of inhibitors. The structure of ICE has been solved by x-ray crystallography to 2.6 Å in complex with an acetyl-Tyr-Val-Asp-H tetra-peptide [50]. Figure 8a illustrates the tertiary structure of the heterodimer with a dimension of about 45 Å × 35 Å × 25 Å. The molecule itself is oligomeric and contains two subunits, p20 (residues 120–297) and p10 (317–404) of relative molecular weight 20 kDa and 10 kDa, respectively. The two subunits form an intimately connected heterodimer. The core of ICE is a six-stranded β sheet containing 5 parallel strands and one antiparallel strand. The core is bounded by six alpha helices that lie parallel to the beta sheets. Physiologically ICE occurs as a (p20)2(p10)2 tetramer. The molecule is a cysteine protease, which has a unique preference among the mammalian proteases for cleaving bonds. It has only one known with aspartic acid adjacent and N-terminal to the P1 scissile bond. It has only one known physiological substrate, proIL-1β, which it cleaves to active IL-1β. The struc-

Figure 8 (a) Stereo diagrams of the heterodimer structure of Interleukin-1 Converting Enzyme (ICE). (b) The tetrapeptide inhibitor (Asp-Ala-Val-Tyr) covalently bound to Cys285 in the active site. Tetrapeptied shown in black, p20 subunit in dark gray, and p10 subunit in light gray. Produced by Molscript [106].

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ture is unlike other cysteine proteases such as papain and appears to have only one known possible homologue, CED-3 protein [54]. The crystal structure with the tetrapeptide (acetyl-Tyr-Val-Ala-Asp-H) substrate in the binding pocket (Figure 8b), coupled with the results obtained by the mutational experiments, has provided a detailed understanding of the enzyme mechanism and substrate bonding of the molecule. Structural analysis has pinpointed a catalytic diad of residues Cys285 and His237 responsible for the catalytic activity of the molecule: Cys285 acts as an active-site nucleophile while the imidazole ring of His237 participates in destabilizing the cysteine Oγ hydrogen. Although both of above these residues are contained in the p20 subunit, the p10 subunit is essential for the maintenance of a binding pocket (and thus the specificity of the molecule), providing binding sites S2 to S4 (residues 338–341) and jointly contributing, with subunit p20, the S1 (Arg179–Arg341) site. The uniqueness and substrate specificity of ICE make it an ideal target for small molecules to block the production of mature, active IL-1β. Since the substrate peptide sequence is known (Tyr-Val-Ala-Asp), it has been a starting point to develop peptidomimetics to inhibit ICE. IV. Therapeutic Strategies A. Manipulating the IL-1 System Since IL-1 is an important mediator of human disease processes, modulating its activity or completely inhibiting its synthesis may be of therapeutic benefit. Blake and Henderson [7] reviewed and portrayed a general strategy to interfere with the cytokine at any one of the following stages: induction or initiation of gene expression–transcription, RNA processing and translation in IL-1 production, folding, release and secretion of extracellular protein, IL-1 in circulation, IL-1 binding to its cell surface receptors, signal transduction and resulting activities (Figure 9). All of the above approaches hold promise for the future. The wealth of structural information on IL-1 combined with extensive site-directed mutagenesis studies on the molecules have helped to build up a picture of the regions involved in biological function. A schematic representation of the present-day efforts to design efficient antagonists of IL-1 its shown in Figure 10. B. Site-Directed Mutants Extensive mutagenesis studies have provided information related to the structural integrity, receptor binding region, and residues that are important for IL-1 function. Site-directed mutagenesis (SDM) can create a single-site mutant and its receptor binding and bioactivity values can be calculated. The results

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Figure 9 Points of intervention for IL-1-based immunomodulation.

obtained from such studies can be imported into the available 3-dimensional structures. A combination of such structural insights coupled with the bioassay results provide clues leading to IL-1 functional information. In a previous structural report on IL-1β, we summarized the SDM results and identified a plausible receptor-binding epitope of interleukin-1 [42]. As mentioned before, electrostatic and hydrogen-bonding interactions in the loops between strands allow the polypeptide to adopt a conformation that enables an unusual concentration of polar and charged groups to be presented at the open end of the barrel. This cluster of charged residues forms an epitope with which IL-1 might bind to the receptor. Following our proposal, many groups have supported this hypothesis by employing mutagenesis studies. For a list of mutants and their activity results refer to References 42, 56, 59, and 61. Ju and coworkers are employing SDM to characterize interleukin-1 [55,56,58,61]. Substitution of Lys for the Asp145 of IL-β (D145K) greatly reduced agonist activity, while retaining 100% binding to the IL1RI [56]. Based on the sequence alignment of IL-1β with IL-1Ra, they selected Lys145 of IL-1Ra for mutagenesis and converted it to an aspartic acid. This mutant analog (IL-1Ra K145D) maintained receptor binding and gained partial agonist activity [56]. Following this study, Ju and coworkers selected five other amino acid residues in IL-1Ra for further analysis because the side chains of these residues appear to be in close proximity to Lys145 in IL-1Ra [61]. Mutations were made at Val18, Thr108, Cys116, Cys122, and Tyr147, usually by a replacement with the corresponding amino acid of IL-1β at each position. None of these muta-

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Figure 10 Schematic diagram showing the IL-1-based therapeutic molecules.

tions provided enough information related to the structure-activity relationship. The mutant IL-1Ra K145D was mutated further to V18S, T108K, C116F, C122S, C122A, Y147T, Y147G, H54P, and a H54I. Of these, K145D + T108K showed a 2-fold decrease in IL-1RI binding and a 3-fold decrease in bioactivity compared to the IL-1Ra K145D analog. The K145D + C116F combination resulted in the complete loss of bioactivity, whereas full receptor-binding activity was maintained. The observation that receptor-binding activity is preserved indicates that the binding site is not altered that much. Structural alignment

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indicates that Cys116 in IL-1Ra is in a homologous position with Phe117 in IL- 1β. The K145D + Y147T analog lost all detectabled activity (both binding and bioactivity), whereas the Y147G analog lost all bioactivity but retained 100% binding. These data suggest that Tyr147 is important for bioactivity of IL-1Ra K145D. C. Insertion of β Bulge A region of charged amino acids (Gln48 to Asn53, a β bulge) positioned between β strands 4 and 5 has been implicated in IL-1β binding to its receptor and its immunostimulatory properties [42, 64–67]. The β-bulge residues form a protrusion on the edge of the open end of the β barrel. Evidence for this patch of amino acids involved in the function of this molecule comes from mutagenesis studies in which deletions or substitutions of residues in this region reduced IL- 1β agonist activity without affecting receptor binding [62,63]. Simoncsits et al. have shown that deletion of amino acids 52–54 (SND) in IL1β reduces IL-1RI binding by 10 fold and biological activity by 1000 fold [63]. Also, studies indicate that a synthetic peptide derived from IL-1β (VQGEESNDK), which contains these six β bulge amino acids, has immunostimulatory but no inflammatory effects normally associated with IL-1 [64–66]. The insertion of VQGEESNDK into recombinant human ferritin H chain and recombinant flagellin from Salmonella muenchen increased the immunogenicity of these antigens in mice [67]. Greenfeder et al. inserted this β-bulge region into IL-1Ra K145D either after Ile51 of after Pro53 [61]. The insertion of the β bulge (QGEESN) after either position 51 or 53 of IL-1Ra K145D resulted in analogs that retained full IL-1RI binding and increased bioactivity by 3–4 fold. Aslo they tried to obtain the analogs of the QGEESN insertion in the absence of the K145D mutation either after amino acid 51 or 53 of IL-1Ra. None of the plasmid clones with the insertion at position 51 of IL-1Ra produced the appropriate protein, whereas they were able to isolate clones with the insertion at position 53. Based on these results, Greenfeder et al. suggest that in the first case the insertion interfered in the proper folding of the protein, whereas the second mutant folded properly and exhibited only 10 to 20% of the IL-1RI binding activity. The mutants with K145D + QGEESN insertion after Ile51 of IL-1Ra showed an increase in bioactivity in the range of 3–8 fold. The triple mutant IL-1Ra K145D/H54P/QGEESN showed a higher bioactivity, and based on their mutagenesis study, Greenfeder et al. suggest that this increase in activity may be due to the introduction of Pro and not the removal of His at position 54 in the K145D mutant of IL-1Ra. The cumulative effects of these three mutations are also interesting since their positions on the IL-1Ra protein appear to be spatially separated. The residues Ile51 and His54 are located on the open face of the β barrel of IL-1Ra, whereas Lys145 is located away from the open barrel end. In IL-1β, the same relative

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positions of the β bulge and Asp145 are observed [42] with the two regions separated by the known IL1RI binding site. We have compiled the available site-directed mutants of IL-1β and sorted them according to the following four categories: (1) mutants that shows significantly higher agonistic activity, A1T, P2M, S5R, N89G, K92R, and K93R; (2) mutants that show significantly higher antagonistic activity, R4E, C8S, T9G, T9Q, T9E, L10T,C,S,A, C71X, R11G, K93M, M95R, E96Q, K103Q, and D145K;, (3) mutants that show significantly higher binding, A1T + P2M, S5R, T9L, T9W, P87S, P87H, K88V,G,L, N89R, E96Q, K103Q, G, C and M148A; and (4) mutants that show significantly lower binding, R4A,K,D, L6A, T9E, L10N,T,C,S,A, H30R, M44S, F46D, A, 156A, V58A, K92E, K93L,A,F,S,E,Q,L, K103S, and E105S,K. These four types of mutants are shown in Figure 11a–d. Taken together, this information supports our earlier proposal of the receptor-binding epitope. This was further characterized as functional sites A and B of interleukin-1. The first site, Area A is structurally conserved in all three molecules and contains residues Arg11, His30, and Asp145 in IL-1β Asn17, Ala36, and Asp147 in IL-1α; and Trp16, Tyr34, and Lys145 in IL-1Ra. Present in both active IL-1 molecules, Asp145 has been recognized as an important residue in IL-1 binding. Area B has also been identified in both IL-1α and IL-1β it contains a large hydrophilic ridge around solvent-accessible hydrophobic residues. In-IL-1β, this region contains the 7-residue hydrophilic ridge (Arg4, Gln48, Glu51, Asn53, Lys93, Glu105, Asn108) around 5 hydrophobic solvent-accessible residues (Leu6, Val47, Ile56, Leu110, Val151). Figure 12 shows a surface presentation of the proposed receptor-binding epitope. In IL-1α, the 7-residue hydrophilic ridge has been identified with residues (Arg12, Ile14, Asp60, Asp61, Ile64, Lys96, and Trp109, which when mutated resulted in significant loss of binding to the receptor. Area B is structurally conserved in IL-1β and in IL-1α, but is lacking in IL-1Ra. For this reason Area A has been proposed as the binding region while Area B has been proposed as the triggering region. D. Peptide Fragments Peptide-based IL-1 antagonists have been derived from the primary sequence of IL-1α or IL-1β. Stepwise synthesis of a series of peptides from amino-terminal to carboxy-terminal regions did not provide any satisfactory results [68]. Polypeptide fragments of IL-1β, termed somnogeneic peptides, induced sleep in mammals [69], 61.5% NREMS at a dose of 20 ng. The numbering of these peptides in proIL-1 are 178–207, 199–225, 208–240 and that of mature IL-1β are 62–91, 83–109, and 92–124 (seq 92–124=KKKMEKRFVFNKIENNKLEFESAQFPNWYIST). Monsanto company has identified a peptide fragment of IL-1β (41–70) exhibiting inhibitory effects for IL-1β and IL-1β, but not TNF-α, at 5 nM (5 ng/mL). Peptide 56–70 is a

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Figure 11 Functional residues of IL-1β, identified by the site-directed mutagenesis results. Produced by Molscript [106].

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Figure 12 The open end of IL-1β, shown as a surface. The positions of some of the residues that have been subjected to site-directed mutagenesis studies have been marked. Produced using the program GRASP [108].

weak agonist. Peptide 47–55 (VQGEESNDK) has been identified as an activator of T cells. It also stimulates glycosaminoglycan synthesis, excites antitumor activity in vivo, and lacks proinflammatory and pyrogenic activities [70]. Later a shorter segment (49–53 = GEESN) with higher activity than 47–55 has been identified. Peptides based on IL-1α and IL-1β sequences were claimed to induce production of prostaglandin E2 but actually maintain other biological activities. Few other peptides or peptidecontaining epitopes were identified by using neutralizing antibodies. Labriola-Tompkins and his colleagues have reported obtaining epitopes for neutralizing antibodies by fractionation of a goat polyclonal antiserum over columns containing individual immobilized synthetic

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Figure 13 The identified peptide fragments. Peptide fragments are shown as striped coil, with residues at the start and end of the peptides are numbered. Produced by Molscript [106], modified by R. Esneuf.

peptides derived from an IL-1α sequence [58]. Their work resulted in 4 peptide regions with residue numbers 4–12, 44–63, 64–88, and 89–105. Peptides 47–55, 81–99, 92–124, 121–153 in the 3dimensional structure of IL-1β are shown in Figure 13. E. Other Strategies Biochemical and structural knowledge has opened many pathways for the development of novel therapeutics. Such strategies include inducer blockers, nucleotide intercalators, antisense RNAs, and other novel molecular mimics. It is known that potent inducers such as lipopolysaccharides, c5a, and integrins bind to IL-1-producing cells and induce the over expression of IL-1. Such an induction can be interrupted by raising the level of neutralizing monoclonal antibodies against the inducers. Alternatively, one can design ligands that can bind to the inducer receptors, which leads to the inhibition of the IL-1 synthesis process. Transcription factors bind to specific DNA sequences and stimulate gene transcription. Controlling such a specific gene transcription can be achieved by a number of means. Intercalators or fragments of the same DNA sequences as those bound by the transcription factors can selectively inhibit transcription. Double-stranded oligonucleotides, having the same consensus sequence, could complete with the transcription factor for binding to the promoter region. Antisense RNA, which when introduced into eukaryotic cells induces sequence-specific inhibition of target gene expression, can be used. The

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antisense strand hybridizes the complementary mRNA to form a double-stranded helix thereby achieving the inhibition of the gene products. Antisense oligo-deoxynucleotide derivatives have been shown to inhibit species-specific fibroblast PGE2 synthesis stimulated by IL-1. These approaches generated much interest and many laboratories are pursuing the design of compounds based on antisense nucleotides, triple-helical transcription inhibitors, aptamers, and other novel nucleic acid derivatives. V. Neutralizing Soluble IL-1 in Circulation Once IL-1 is released into the extracellular fluid, the following molecules can be employed to manipulate its activity: soluble receptors, IL-1Ra, IL-1 neutralizing antibodies, IL-1-specific binding proteins, high-affinity small molecules (Figure 10). A. Soluble IL-1 Receptors The most effective neutralizer of a cytokine is likely to be its receptor. Some viruses have been reported to use an IL-1 receptor mimic to evade the immune system [72]. Natural shedding of cytokine receptors is a common occurrence and may form part of a normal homeostatic regulatory system, and there is a high potential for the use of such soluble forms as therapeutic agents. The binding affinity of the mature forms of the interleukin-1 molecules and their receptors have been reported [8,31–33,71]. These studies show that the affinity of both the membrane-bound and soluble forms of human IL-1RI for the mature forms of human IL-1α, IL-1β, and IL-1Ra are approximately the same. In contrast to IL-1RI, IL-1RII binds IL-1βpreferentially. Based on the binding-affinity studies one can infer that soluble IL-1RI is a better inhibitor of IL-1α than IL-1β and soluble IL-1RII is a better inhibitor of IL-1β. Both soluble receptors, at sufficiently high concentrations, will completely block the binding of both IL-1 forms to cells. Dower and coworkers [8] have studied the real-time binding of human IL-1α, IL-1β, and IL-1Ra to human soluble IL-1RI and IL- 1RII. It seems that the binding of IL-1Ra to IL-1RI is essentially irreversible, whereas its binding to IL-1RII is rapidly reversible. In contrast, IL-1RII binds IL-1β irreversibly [8,1]. This indicates that the IL-1RII can be used as a high-affinity antagonist of IL-1β. Dower further suggest in his studies that the soluble receptors have several potential advantages over anticytokine antibodies due to the fact that they have much higher affinities (100 to 1000 fold) and should not be recognized by the immune system. Even though soluble receptors have high-binding affinity they are difficult to synthesize in large quantities and they have a low half-life.

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B. Interleukin-1 Receptor Antagonist Over the past 5–10 years a number of inhibitors of IL-1 and TNF have been found in biological fluids and cell-culture supernatants. Interleukin-1 receptor antagonist was the first protein receptor antagonist to be described. Recombinant IL-1Ra has been shown to block the activity of IL-1α and IL-1β both in vitro and in vivo in animal models by binding to both type I and type II IL-1 receptors without demonstrable agonist activity. These effects have been extensively studied. However clinical trails carries out by Synergen on human subjects provided a negative support for the antagonist behavior of IL1Ra for sepsis. It may be that the system took an alternative route by inducing TNF in excessively high levels thus achieving signal transduction. Therefore neutralizing TNF in combination with IL-1 inhibition could be an alternate procedure. Even though IL-1Ra exhibits very high binding affinity it is rather a poor inhibitor of IL-1 action in vivo. It must be present at greater than a 100-fold molar excess over either agonist form to block action, and large doses are required to block IL-1-mediated effects in vivo [1]. Burger and Dayer suggest that the simultaneous use of IL-1Ra and IL-1RII might be beneficial, since this mixture—contrary to the use of IL-1Ra alone—completely abolished the production of interstitial collagenase in the inflammatory pathway [73]. C. Monoclonal Antibodies Chimarized or humanized neutralizing monoclonal antibodies for IL-1 can be used as IL-1 can be used as IL-1 antagonists. Otsuka Pharmaceuticals has developed a monoclonal antibody (IgG1 kappa) against IL-1β. It can be used in the immunoassay method for the selective detection of human IL-1β and also to determine the biological activity of IL-1β. Their patent (EP 0-364-778) also covers the use of such an antibody against IL-1β. when it is abnormally produced in disease states. Another patent of Otsuka Pharmaceuticals (EP 0- 408-859-A2) relates to an antibody and its application to inflammatory processes. This antibody is directed to a specific antigen on activated human endothelial cells (1E7/2G7) and blocks the binding of white blood cells causing inflammatory responses. Based on the antibody complementarity-determining region low-molecular-weight nonpeptide mimetics can be developed. This attempt might result in low-molecular-weight, orally active compounds. D. Low-Molecular-Weight Antagonists Low-molecular-weight antagonists are attractive due to their low cost and bioavailability. Available literature indicates that many laboratories are attempting to create immunomodulators of small synthetic molecules, peptidomimetics, bacterial cell-wall components, macrocycles, corticosteriods, and others [74].

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Figure 14 2-Dimensional structures of small molecules that exhibit IL-1 modulating activities. They are tenidap, ciprofloxacin, 3-Deazaadenosine, (SK&F 86002), E5110, DMARDs (Chloroquine, Auranofin, Sodium aurothiomalate and Dexamethasone) tiaprofenic acid, dexamethasone, tricyclic-ylidene-acetic acid and its derivative, Probucol, eicosapentenoic acid + docosahexenoic, pentoxifylline, Denbufylline, and Romazarit (Ro-31-3948).

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American Home Products has patented molecules ranging from substituted quinoline, piperidine, and naphthpyridine compounds as immunomodulators. Ciprofloxacin, a quinolone antibiotic, reduced the extracellular IL-1 activity in human monocytes and delayed the peak production of IL-1α IL-β by 24 h and decreased total IL-1β production, but did not change total IL-1α production [75,76]. Tenidap, an antiarthritic drug, has shown efficacy both in rheumatoid arthritis and osteoarthritis [77,78]. It is a known inhibitor of 5-lipoxygenase and cyclooxygenase (5-LO/CO) and in vitro studies indicate that it also inhibits the synthesis of mature IL-1 and pro IL-1 [77,79]. Kadin reports analogs of Tenidap as antiinflammatory agents and analgesics [80]. An antiarthritic molecule, 3- Deazaadenosine, has been demonstrated to inhibit IL-1 production by LPS- stimulated human PBMCs acting at the level of RNA synthesis and also by blocking the effects of IL-1α on EL4 cells and induction of PGE2 release by human fibroblast [81,82]. Another 5-LO/CO inhibitor (SK&F 86002) has been reported to inhibit the synthesis of IL-1 in human monocytes and human synovial cells in a dose-dependent manner [83,84]. Analogs of this compound

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Figure 14 (Continued)

have been patented as IL-1 inhibitors [85–87]. In human monocytes, E-5110 is a dual 5-LO/CO inhibitor found to reduce extra- and intracellular IL-1 activity induced by LPS in a dose-dependent manner. This compound also inhibits the IL-1 generation induced by antigen-antibody complexes, zymosan, and silica particles [88].

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Antinflammatory DMARDs such as chloroquine, auranofin, sodium aurothiomalate, and dexamethasone have been shown to inhibit IL-1 synthesis [89]. Analogs of these compounds have exhibited potent inhibition of IL-1α- induced cartilage resorption [90]. Elevated collagenase and proteoglycanase levels caused by IL-1 in human cartilage were found to be reduced by tiapro-

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Figure 14 (Continued)

fenic acid and dexamethasone [91]. A patent report from the National Institutes of Health describes a method of treating diseases associated with elevated levels of interleukin-1. Rosenthal, as the inventor of this patent, describes a method for inhibiting the release of IL-1 from IL-1-producing cells by administering a therapeutically effective amount of an aromatic diamidine (WO9115201-A).

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These aromatic diamidines inhibit IL-1 production and also block IL-6 and TNF. Imidazoline blocks IL1 and TNF and is less toxic to the cell with an in vitro LD50 >>10-4. Examples of these compounds are 1,5-bis(4-amidophenoxy) pentane (pentamidine), in the form of pentamidine isothionate, and an imidazoline in the form of 1,5-di(4-imidazolinophenoxy)pentane. Tricycli- cylidene-acetic acid [92] and its 2-chloro derivative [93] were found to be inhibitors of IL-1 release, claiming clinical improvement in patients with psoriasis, periodontal disease, and Alzheimer's disease. In vitro this compound blocks the synthesis of prostaglandins and inhibits the release of IL-1α and IL-1β from human monocytes and murine macrophages. Probucol, a hypocholesterolemic drug that possesses antioxidant activity, inhibits the ex vivo release of IL-1 from LPS-stimulated macrophages of mice pretreated orally with 100 mg/kg/day of this compound [94,95]. This compound has been shown to inhibit LPS-induced zinc-lowering effect, is cited as direct evidence for the inhibition of IL-1 release, and may be useful candidate for the treatment of atherosclerosis [95,96]. An amino-dithiol-one derivative (RP 54745) blocked the proliferative action of IL-1β on murine thymocytes in vitro and also inhibited the production of IL-1 in mouse peritoneal macrophages in vitro and in vivo. The compound RP 54745 selectively inhibited the expression of IL1α and IL-1β mRNA while TNFα mRNA was unaffected [97, 98]. Administration of a cocktail containing eicosapentenoic acid and docosahexenoic acid to volunteers for up to 6 weeks, resulted in a significant depression in IL-1β (61%), IL-1α (39%), and TNF (40%) synthesis. These levels returned to normal after a few weeks [99]. In vitro studies indicate that Pentoxifylline can block the effects of IL-1 and TNF on neutrophils [100]. It is a phosphodiesterase (PDE) inhibitor that causes increased capillary blood flow by decreasing blood viscocity and is used clinically in chronic occlusive arterial disease of the limbs with intermittent claudication. Denbufylline, a closely related xanthine, has been patented as a functional inhibitor of cytokines and exhibits a similar profile to Pentoxifylline [101]. Romazarit (Ro-31-3948) derived from oxazole and isoxazole propionic acids has been shown to block IL- 1-induced activation of human fibroblasts in vitro and in animal models reduces inflammation [102,103,104]. By using a spontaneous autoimmune MRL/lpr mouse model, a significant efficacy was shown [105]. Two-dimensional structures of some of these molecules are shown in Figure 14. Even though the above mentioned small molecules exhibit IL-1 inhibition none of them were discovered based on defined functional or structural aspects. An understanding of the three-dimensional structure of IL-1s and their receptors, by themselves or in complexes, will form a very strong foundation for structure-based design of more specific and potent IL-1-based immunomodulators.

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VI. Conclusion The design of novel compounds to inhibit or manipulate the IL-1 system remains a daunting task. At this time, the design of immunomodulators for the IL-1 system is still in its infancy and has largely been confined to the use of whole or fragmented proteins or the identification of nonspecific small molecules. In addition, newer approaches have also been initiated and these include the use of antisense oligonucleotides, small molecules designed to compete with IL-1's binding to its receptor, ICE inhibitors, and intracellular signaling inhibitors. All such strategies show promise. Structure-based design has not been explicitly used in the design of agonists and antagonists of IL-1. But as of now we have the structures of IL-1α, IL-1β, and IL-1Ra. A new insight may be forthcoming once the complex crystallographic structure of one of the interleukin-1 molecules and its corresponding receptor molecule is available. This structural information, coupled with the anticipated IL-1 + IL-1R complex structure, will form the foundation for rational design of inhibitors with improved selectivity for the treatment of various IL-1-mediated diseases. Acknowledgements Our special thanks to Professor Russell Doolittle for his encouragement and support, and also to Dr. Mitch Lewis for providing us with the very high-resolution coordinates of IL-1α. We gratefully acknowledge San Diego Super Computing Center for their assistance and support in providing valuable software and high-power computing time. We thank Dr. Donald Kyle for his valuable comments. We also thank Dr. Per Kraulis for providing us the latest version of MOLSCRIPT, Dr. Anthony Nicholls for the program GRASP, Dr. Rob Russell for the structural alignment, and Professor Lynn Ten Eyck and Dr. Jerry Greenberg for their help. References 1. Dinarello CA. Biological basis for interleukin-1 in disease. Blood 1996; 87- 6:2095–2147. 2. Dinarello CA. Interleukin-1 is produced in response to infection and injury. Rev infect Dis 1984;6:51–56. 3. Oppenheim JJ, Kovacs EJ, Matsushima K, Durum SK. There is more than one interleukin 1. Immunol Today 1986;7:45–56. 4. Durum KS, Oppenheim JJ, Neta R. Immunophysiologic role of interleukin-1. In: Oppenheim JJ, Shevach EM, eds. Immunophysiology. New York: Oxford University Press, 1990:210–225.

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5. Dinarello CA. On the Biology of Interleukin-1. Adv Immunology 1989; 44:153– 205. 6. Meager A. Cytokines. 1990; Open University Press. 7. Henderson B, Blake S. Therapeutic Advantage of Cytokine Manipulation. TIPS 1992; 13:145–152. 8. Dower SK, Fanslow W, Jacobs C, Waugh S, Sims JE, Widmer MB. Interleukin-1 antagonists. Therapeutic Immunol 1994; 1:113. 9. Burger DD, J. Inhibitory Cytokines and Cytokine Inhibitors. Neurology 1995; 45(supp 16):s39–43. 10. Abraham DJ. X-ray crystallography and drug design. In: Perun TJ, Propst CL, eds. Computer Aided Drug Design. New York: Marcel Dekker, Inc., 1989:93–132. 11. Fesik SW. Approaches to drug design using nuclear magnetic resonance spectroscopy. In: Perun TJ, Propst CL, eds. Computer-Aided Drug Design. New York: Marcel Dekker, Inc., 1989:133–184. 12. Veerapandian B. Structure aided drug design. In: Wolf M, ed. Buerger's Drug Discovery and Medicinal Chemistry. New York: John Wiley and Sons, 1995:303– 348. 13. Whittle PJ, Blundell TL. Protein Structure based drug design. Ann Rev Biophy and Biomol Structure 1994; 23:349–375. 14. Carter DB, Deibel MRJ, Dunn CJ, Tomich C-SC, Laborde AL, Slightom JL. Berger AE, Bienkowski MJ, Sun FF, McEwan RN, Hams PKW, Yem AW, Waszak GA, Chosay JG, Sieu LC, Hardee MM, Zucher-Neely HA, Reardon IM, Heinnckson RL, Truesdell SE, Shelly JA, Eessalu TE, Taylor BM, Tracey DE. Purification, cloning, expression, and biological characterization of an interleukin-1 receptor antagonist protein. Nature 1990; 344:633–638. 15. Eisenberg SP, Evans RJ, Arend WP, Verderber E, Brewer MT, Hannum CH, Thompson RC. Primary structure and functional expression from complementary DNA of a human interleukin-1 receptor antagonist. Nature, 1990; 343:341–346. 16. Sims JE, March CJ, Cosman D, Widmer MB, MacDonald HR, McMahan CJ, Grubin CE, Wignall JM, Jackson JL, Call SM, et al. cDNA expression cloning of the IL-1 receptor, a member of the immunoglobulin superfamily. Science 1988; 241:585. 17. McMahon CJ, Slack JL, Mosley B, Cosman D, Lupton SD, Brunton LL, Grubin CE, Wignall JM, Jenkins NA, Brannan C1, Copeland NG, Huebner K, Croce CM, Cannizzaro LA, Benjamin D, Dower S, Spriggs MK, Sims JE. A novel IL-1 receptor cloned from B cells by mammalian expression is expressed in many cell types. EMBO J 1991; 10:2821.

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35. Bendtzen K, Svenson M, Jonsson V, Hippe E. Autoantibodies to cytokines-Friends or foes? Immunol Today 1990; 11:167. 36. Svenson M, Hensen MB, Bendtzen K. Distribution and characterization of autoantibodies to interleukin-1α in normal human sera. Scand J Immunol 1990; 32:695. 37. Svenson M, Hensen MB, Kayser L, Rasmussen AK, Reimert CM, Bendtzen K. Effects of human anti-IL-1α autoantibodies on receptor binding and biological activities of IL-1. Cytokine 1992; 4:125. 38. Satoh H, Chizzonite R, Ostrowski C, Ni-Wu G, Kim H, Fayer B, Mae N, Nadeau R, Liberato DJ. Characterization of antiIL-1α autoantibodies in the sera from healthy humans. Immunopharmacology 1994; 27:107.

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39. Saurat JH, Schfferli J, Steiger G, Dayer J-M, Didierjean L. Anti-interleukin-1α auto antibodies in humans. J Allergy Clin Immunol 1991; 88:244. 40. Preistle JP, Schar H-P, Grutter MG. Crystallographic Refinement of Interleukin 1β at 2.0 Å Resolution. Proc Natl Acad Sci USA 1989; 86:9667–9671. 41. Finzel BC, Clancy LL, Holland DR, Muchmore SW, Watenpaugh K, Einspahr HM. Crystal structure of recombinant human interleukin-1β at 2.0 Å resolution. J Mol Biol, 1989; 209:779–791. 42. Veerapandian B, Gilliland GL, Raag R, Svensson AL, Masui Y, Hirai Y, Poulos TL. Functional implications of interleukin-1β bases on the three-dimensional structure. Proteins: Struct, Funct Genet 1992; 12:10–23. 43. Clore GM, Wingfield PT, Gronenborn AM. High resolution three dimensional structure of interleukin-1β in solution by three and 4 dimensional nuclear magnetic spectroscopy. Biochemistry 1991; 30:2315–2319. 44. Ohlendorf DHT, A, Weber PC, Wondolski JJ, Salemme FR, Lischwe M, Newton RC. A comparison of the high resolution structures of human and marine interleukin-1β to be published. 45. Graves BJ, Hatada MH, Hendnckson WA, Miller JK, Madison VS, Satow Y. Structure of interleukin1α at 2.7 A resolution. Biochemistry 1990; 29:2679– 2684. 46. Stockman BJS, TA, Strakalaitis NA, Brunner DP, Yem AW, Deibel MR Jr. Solution structure of human interleukin-1 receptor antagonist protein. FEBS Letters 1994; 349:79–83. 47. Vigers GPA, Caffes P, Evans RJ, Thompson RC, Eisenberg SP, Brandhuber BJ. X-ray structure of interleukin-1 receptor antagonist at 2.0 Å resolution. J of Biol Chem 1994; 269:12874–12879. 48. Schreuder HAR, J-M, Tardif C, Soffientini A, Sarubbi E, Akeson A, Bowlin TL, Yanofsky S, Barrett RW. Crystal structure of the interleukin-1 receptor antagonist. To be published. 49. Spraggon G, Singh O, Stuart DI, Jones EY. The crystal structure of intact interleukin-1 receptor antagonist. To be published. 50. Wilson KPB, JF, Thomson JA, Kim EE, Griffith JP, NAvia MA, Murcko MA, Chambers SP, Aldape RA, Raybuck SA, Livingston DJ. Structure and mechanism of interleukin-1β converting enzyme. Nature 1994; 370:270–275. 51. McLachlan AD. Three-fold structural pattern in the soybean trypsin inhibitor (Kunitz). J Mol Biol 1979; 133:557–563.

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52. Murzin AG, Lesk AM, Chothia C. b- Trefoil Fold: patterns of structure and sequence in the kunitz inhibitors interleukins-1β and 1α and fibroblast growth factors. J Mol Biol 1992; 223:531–543. 53. Ohelndorf DH. Accuracy of refined proteins structures II. Comparison of four independantly refined models of interleukin-1β. Acta Cryst 1994; D50:808–812. 54. Yuan J, Shaam S, Ledoux S, Ellis HM, Horvitz HR. The C. elegans cell death gene ced-3 encodes a protein similar to mammalian interleukin-1β converting enzyme. Cell 1993; 75:641–652. 55. Labriola-Tompkins E, Chandran C, Kaffka K L, Biondi D, Graves BJ, Hatada M, Madison VS, Karas J, Klian PL, Ju G. Identification of the discontinuous binding site in human interleukin-1β for the type I interleukin-1 receptor. Proc Natl Acad Sci USA 1991; 88:11182–11186. 56. Ju G, Labriola-Tompkins E, Campen CA, Benjamin WR, Karas J, Plocinski J, Biondi D, Kaffka KL, Klian PL, Eisenberg SP, Evans RJ. Conversion of the IL-1

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receptor antagonist into an agonist by a single amino acid substitution. Proc Natl Acad Sci USA 1991; 88:2658–2662. 57. Kawashima H, Yamagishi J, Yamayoshi M, Ohue M, Fukwi T, Kotani H, Yamada M. Structureactivity relationships in human interleukin-1α: identification of key residues for expression of biological activities. Protein Eng 1992; 5–2:171–176. 58. Labriola-Tompkins E, Chandran C, Varnell TA, Madison VS, Ju G. Structure-function analysis of human IL-1α: Identification of residues required for binding to the human type I IL-1 receptor. Protein Eng 1993; 6–5:535–539. 59. Guinet F, Guitton J, Gault N, Folliard F, Touchet N, Cherel J, Crespo A, Destourbe A, Bertrand P, Denefle P, Mayaux J, Bousseau A, Duchesne M, Terlain B, Cartwright T. Interleukin-1β specific partial agonists defined by site-directed mutagenesis studies. Eur J Biochem 1993; 211:583–590. 60. Grutter MG, van Oostrum J, Pnestle JP, Edelmann E, Joss U, Feige U, Vosbeck K, Schmitz A. Protein Eng, 1994; 7:663–671. 61. Greenfeder SA, Varnell T, Powers G, Lombard-Gillooly K, Shuster D, McIntyre KW, Ryan DE, Leven W, Madison V, Ju G. Insertion of a structural domain of interleukin-1β confers agonist activity to the IL-1 receptor antagonist. J Biol Chem 1995; 270:22460–22466. 62. Wolfson AJ, Kanaoka M, Lau F, Ringe D, Young P, Lee J, and Blumenthal J. Biochemistry 1993; 32:5327–5331. 63. Simoncsits A, Bnstulf J, Tjornhammar ML, Cserzo M, Pongor S, Rybakina E, Gatti S, Bartfai T. Cytokine 1994; 6:206–214. 64. Antoni G, Presentini R, Penn F, Tagliabue A, Ghiara P, Censini S, Volpini G, Villa L, Boraschu D. J Immunol 1986; 137:3201. 65. Boraschi D, Nencioni L, Villa L, Censini S, Bossu P, Ghiara P, Presentini R, Perin F, Frasca D, Dona G, Forni G, Musso T, Giovarelli M, Ghezzi R, Bertini R, Besedovsky H, Del Rey A, Sipe J, Anotoni G, Silvestn S, Tagliabue A. J Exp Med 1988; 168:675–686. 66. Frasca D, Boraschi D, Baschien S, Bossu P, Tagliabue A, Adorini L, Dona GJ Immunol 1988; 141:2651–2655. 67. Beckers W, Villa L, Gonfloni S, Castagnoli L, Newton SMC, Cesareni, Ghiara P. J Immunol 1993; 151:1757–1764. 68. Joss UR, Schmidli I, Vosbeck K. Mapping the receptor binding domain of interleukin-1β by means of binding studies using overlapping fragments: Why did it fail? J Recept Res 1991; 11:275–282.

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69. Obal F, Opp M, Cady AB, Johannsen L, Postlethwaite AE, Poppleton HM, Seyer JM, Krueger JM. Interleukin-1β and an interleukin-1β fragment are somnogenic. Am J Physiol, 1990; 259:R439. 70. Antoni G, Presentini R, Perin F, Tagliabue A, Ghiara P, Censini S, Volpini G, Villa L, Boraschi D. Peptide analogues of IL-1 and biochemical assay of their binding to its receptors. J Immunol 1986; 137:3201–3204. 71. Slack J, McMahan CJ, Waugh S, Schooley K, Spriggs MK, Sims JE, Dower SK. Independent binding of interleukin-1α and interleukin-1β to type I and type II interleukin-1 receptors. J Biol Chem 1993; 268:2513. 72. Alcami AS, Smith GL. A soluable receptor for interleukin-1β encoded by Vaccinia virus: A novel mechanism of virus modulation of the host response to infection. Cell 1992; 71:153–167. 73. Burger D, Dayer JM. Inhibitory cytokines and cytokine inhibitors. Neurology 1995; 45 (suppl 6):S39–S43.

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74. Bender PE, Lee JC., eds. Pharmacological modulation of interleukin-1 (1989). Annual reports in medicinal Chemistry-25. Johns. Section IV-Metabolic diseases and endocrine function. Chapter 20. 75. Roche Y, Fay M, Gougerot-Pocidalo MA. Antimicrob Chemother 1988; 21:597. 76. Bailly S, Mahe Y, Ferrua B, Fay M, Wakasugi H, Tursz T, Gougerot-Pocidalo MA. Cytokine 1989; 1:303. 77. Otterness IG. Abstracts, 3rd Interscience World Conference on Inflammation. Monte-Carlo, 1989:371. 78. Otterness IG, Bliven ML, Downs JT, Manson DC. Arthritis Rheum. Abstracts, 1988; 31–4:S90, C55. 79. McDonald B, Loose L, Rosenwasser LJ. Arthritis Rheum. Abstracts, 1988; 31– 4:S52, A88. 80. Kadin, U.S. Patent 4,730,004 (1988). 81. Jurgensen CH, Wolberg G, Zimmerman TP. Agents Actions 1989; 27:398. 82. Schmidt JA, Bomford R, Gao XM, Rhodes J. Int J Immunopharmacol 1990; 12:89. 83. Lee JC, Griswold DE, Votta B, Hanna N. Int J Immunopharmacol, 1988; 10:835. 84. Lee JC, Votta B, Griswold DE, Hanna N. Agents Actions 1989; 27:280. 85. Bender PE, Griswold DE, Hanna N, Lee JC. 1988; U.S. Patent 4,794,114. 86. Bender PE, Griswold DE, Hanna N, Lee JC. 1988; U.S. Patent 4,780,470. 87. Bender PE, Griswold DE, Hanna N, Lee JC. 1988; U.S. Patent 4,778,806. 88. Shirota H, Goto M, Hashida R, Yamatsu I, Katayama D. Agents Actions 1989; 27:322. 89. Goodacre J, Carson WD. Allison in Immunopathogenetic Mechanisms of Arthritis. Boston: MTP Press, 1988:211. 90. Rainford KD. J Pharm Pharmacology 1989; 41:112. 91. Shinmei M, Kikuchi T, Masuda K, Shimomura Y. Drugs, 1988; 35 (Suppl. 1):33. 92. Seibel MJ, Bruckle W, Respondek M, Beveridge T. Schnyder J, Muller W, Rheumatol Z. 1989; 48:147. 93. Bollinger P, Gubler HU, Schnyder J. 1989; Derwent 89–138880–B2; DE 38 36 329 Al.

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94. Ku G, Doherty N. 1988; Derwent 88-314770; AU-A-13160/88. 95. Ku G, Doherty NS, Wobs JA, Jackson RL. A, J Cardiol 1988; 62:778. 96. Marx JL. Science 1988; 239:257. 97. Folliard F, Terlain B. Abstracts, 3rd Inter-science World Conference on Inflammation. MonteCarlo; 1989; 415. 98. Folliard F, Bousseau A, Terlain B. Cytokine 1989; 1:108. 99. Endres S, Ghorbani R, Keliey VE, Georgilis K, Lonnemann G, van der Meer JWM, Cannon JG, Rogers TS, Klempner MS, Weber PC, Schaefer EJ, Woldf SM, Dinarello CA. N Engl J Med 1989; 320:265. 100. Sullivan GW, Carper HT, Novick Jr. WJ, Mandell GL. Infect Immun 1988; 56:1722. 101. Mandell GL, Sullivan GW, Novick Jr. WJ, 1989; Derwent 89–191 551–B2; WO 89 05145.

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102. Machin PJ, Osbond JM, Sqix CR, Smithen CE, Tong BP. U.S. Patent 4,774,253 (1988). 103. Bloxham DP, Bradshaw D, Cashin CH, Dodge BB, Lewis EJ, Westmacott D, Barber W.E, Machin PJ, Osbond JM, Self CR, Smithen CE, Tong BP. Brit J Rheumatol 1987; 26 (Suppl. 2):2.

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104. Bradshaw D, Dodge BB, Franz PH, Lee SC, Wilson SE. Abstracts, 3rd Interscience World Conference on Inflammation. Monte-Carlo, 1989, 183. 105. Sedgwick AD. Abstracts, 3rd Interscience World Conference on Inflammation. Monte-Carlo; 1989:183. 106. Kraulis PJ, MOLSCRIPT: a program to produce both detailed and schematic plots of protein Structures. J Appl Cryst 1991; 24:946–950. 107. Merrit EM, M RASTER 3D version 2.0: a program for photorealistic molecular graphics. Acta Crystallogr 1994; D50:869–873. 108. Nicholls A, Sharp KA, Honig B. Protein folding and association: insights from the interfacial and thermodynamic properties of hydrocarbons. Proteins 1991; 11:281–296. 109. Russell RB, Barton CJ, Proteins, 1992; 14:309–323. 110. Barton CJ, Protein Engineering, 1989; 6:37–40.

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17 Structure and Functional Studies of Interferon: A Solid Foundation for Rational Drug Design Michael A. Jarpe Cambridge NeuroScience, Inc., Cambridge, Massachusetts Carol H. Pontzer University of Maryland, College Park, Maryland Brian E. Szente* and Howard M. Johnson University of Florida, Gainesville, Florida I. Introduction The interferons (IFNs) were discovered in 1957 by Isaacs and Lindenman when they observed that a substance secreted by virally infected cells could protect other cells from viral infection [1a]. They called this substance interferon and found that it was a protein that caused uninfected cells to produce other proteins that made them resistant. Researchers since then have been finding a growing family of structurally related molecules: the interferons. Through the years, the interferons have been given many different names including immune, fibroblast, leukocyte, Type I, and Type II interferons. The recognized nomenclature includes alpha, beta, omega, tau, and gamma (α, β, ω, τ, and γ) interferons. Alpha, beta, omega, and tau all belong to the similar Type I subclass. Gamma is the sole member of the Type II or immune interferon class. The Type I interferons all share a greater sequence homology to each other than they do to IFN-γ (for a recent general review of the IFNs, see Reference 1b). The IFNs exert their actions on cells via cell surface receptors. Type I IFNs share the IFN Type I receptor (IFN-R1) while IFN-γ has its own unique Type II receptor. The signal transduction pathways of Type I and Type II *

Current affiliation: Brigham and Women's Hospital, Boston, Massachusetts.

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Figure 1 Interferon activity.

receptor activation are similar. These pathways involve ligand and receptor binding followed by the activation of tyrosine kinases and the phosphorylation of various proteins and their subsequent interaction with transcription elements on DNA. The two receptor activation pathways differ at the level of ligand binding. Type I IFNs bind their receptor as a complex of a single ligand, ligand binding element, and an accessory molecule. The Type II binding event occurs as a dimer of IFN-γ binding to two identical receptor molecules leading to receptor dimerization and activation. The IFNs of all subclasses posses antiviral activity. Additionally, they produce cellular responses that are distinct from antiviral activity, including antiproliferative and immunomodulatory activities (Figure 1). These activities have led to an interest in their use as potential therapeutics to combat viral disease, cancer, and autoimmune disease. Currently, the IFNs have a worldwide market in excess of 2 billion dollars annually. There are six FDA-approved indications in the United States with several more in clinical trials (Table 1). In fact, two of the top-ten grossing biotechnology-based drugs on the U.S. market are IFNs. Intron A is an IFN-α used for immune protection and has an annual U.S. market of $570 million. Roferon-A is another IFN-α used for hairy-cell leukemia and Kaposi's sarcoma with an annual market of $170 million. These sales are despite profound negative side effects associated with IFN treatment. High doses are required to achieve positive clinical results and can lead to severe flu-like symptoms including nausea, vomiting, and fever. These side effects can cause patients to drop out of treatment before beneficial effects are seen. Another drawback of IFNs as drugs is that they require parenteral delivery. The IFNs are protein drugs that must be administered by injection and

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Page 437 Table 1 Approval Indications for IFNs FDA Approved IFN-α

Clinical Trials

Chronic hepatitis

HIV infection

Kaposi's sarcoma

Colon tumors

Genital warts (papillomavirus)

Kidney tumors Bladder cancer

Hairy cell leukemia

Malignant melanoma Non-Hodgkin's lymphoma Chronic myelogenous leukemia Throat wartz (papillomavirus)

IFN-β

Relapsing remitting multiple sclerosis

Basal cell carcinoma

IFN-γ

Chronic granulomatous disease

Kidney tumors Leishmaniasis

cannot be given orally. For many of the clinical indications, treatments of many months are needed requiring repeat injections. These drawbacks, coupled with the market value of IFN-related treatments, now and in the future, have created an interest in producing second-generation molecules that can mimic IFN activity. These “mimetics” could potentially have greater specificity with fewer side effects. They may also have the advantages of reduced manufacturing costs and more versatile delivery. The design of mimetics can be achieved through structure-based drug design methodologies that are currently being developed. However, in order to apply structure-based drug design to a protein, a solid understanding of the structure/function relationship is needed. A three-dimensional structure, taken alone, gives little insight into the activity of a protein. Structure/function studies must be done for the full potential of structurebased drug design to be realized.

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Structure/function studies can take a variety of forms and use a number of techniques including the use of molecular biology, synthetic peptides, and antibodies, or combinations of these methods. Molecular biology is a powerful tool for structure/function analysis. Mutagenesis of cDNAs to produce mutant proteins with point mutations, truncations, or deletions can identify functional sites. One drawback to this approach, especially with large proteins, is the proverbial “needle in a haystack” problem. One has difficulty determining where to begin placing mutations. The synthetic peptide approach can be equally as powerful. One can synthesize individual domains or segments of proteins and test them for agonist or antagonist activities thereby identifying functional domains. Synthetic peptides can be used to map the epitope specificity of antibodies that block the activity of a protein. Peptides can also be used to produce monospecific antisera to a defined region of a protein. The antibody approach has also proved quite useful in determining functional sites of

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proteins. One potential disadvantage to using antibodies is the possibility of over interpreting the blocking results because of steric hindrance. A large antibody molecule may inhibit function through binding to a distant site and covering up a functional site. These approaches have all been used with success on a variety of proteins, but are best used in combination. For example, the information obtained from synthetic peptides and antibodies can significantly narrow down the region for site-directed mutagenesis studies. The entire sequence is narrowed to a segment, which reduces the size of the “haystack” in which the needle is hidden. Even though these approaches are powerful methods for determining functional sites on proteins, they are limited if not coupled with some form of structural determination. As Figure 2 illustrates, molecular biology and synthetic peptide/antibody approaches are not only interdependent, they are tied in with structural determination. Structural determination methods can take many forms, from the classic x-ray crystallography and NMR for three-dimensional determination, to two-dimensional methods such as circular dichroism and Fourier Transformed Infrared Spectroscopy, to predictive methods and modeling. A structural analysis is crucial to the interpretation of experimental results obtained from mutational and synthetic peptide/antibody techniques.

Figure 2 Flow diagram of structure/function studies.

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Page 439 Table 2 Three-Dimensional Structure Studies of the IFNs Type of Study

Reference

X-ray studies IFN-β

2

IFN-γ human

3

IFN-γ bovine

C.T. Samudzi, J.R. Rubin, unpublished data

IFN-γ rabbit

4

IFN-γ human + receptor

5

NMR studies IFN-γ human

6

IFN-γ mouse N-terminal peptide (1–39)

7

Models IFN-α2a

8

IFN-α8

9

IFN-τ sheep

10 T. Senda, S.I. Saitoh, Y. Mitsui, J.Li, and R.M. Roberts, unpublished data

Note: This is not meant to be an exhaustive list of all structural studies of the IFNs. It only highlights some of the three-dimensional studies that have been conducted.

While there are no hard-and-fast rules for conducting structure/function studies, the approaches taken for studying the IFNs can be used to illustrate some of the methods that have been successful. Over the years, a large body of work has accumulated on the IFNs, including a number of structural studies. Table 2 summarizes some of the studies exploring the three-dimensional structure of the IFNs. The following sections review some of the structure/function studies that have begun to elucidate important features of IFN activity and form a basis for future rational drug design.

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II. Type IFNs A great deal of structure/function analysis has been done on the Type I IFNs. One Type I IFN in particular, IFN-τ has received attention recently because of its lower cytotoxicity compared to the IFNαs. Structure/function studies have concentrated on comparing IFN-τ with IFN-α. Therefore, IFN-τ provides an excellent example of structure/function studies of the Type I IFNs. First isolated from the conceptuses of sheep, IFN-τ is the major conceptus secretory protein responsible for signaling maternal recognition of pregnancy in ruminants [11]. it is produced in large quantities (200 µg in 30 h from a day 16 conceptus culture). The protein was purified using a combination of anion exchange and molecular sieve chromatography.

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A. IFN-τ Synthetic Peptide Studies A sheep blastocyst library was screened with a probe based on the N-terminal sequence of the IFN-τ protein and the cDNA obtained (Table 3). Surprisingly, it exhibited 45–55% homology with various IFNs from human, mouse, rat, and pig and 70% homology with bovine IFN-ω [12]. It shared both molecular weight (19 kDa) and pI (5.4–5.6) with IFN-αs, while its length, 172 amino acids, was equivalent to the IFN-ωs. In competition studies, IFN-τ was found to compete with IFNs α, β, and ω for binding to the Type I IFN receptor [13]. In contrast, IFN-τ exhibited several unique properties such as its reproductive function, its poor inducibility by virus, and its apparent reduced cytotoxicity. Thus, IFNτ conceptus protein appears to be a novel IFN. Structural studies began with production of overlapping synthetic peptides, each 30–35 amino acids in length, corresponding to the entire

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sequence of the molecule [14]. The peptides were used in competition assays with the native molecule. Peptide inhibition of a particular function would implicate the region of the molecule that it represented in the elicitation of that function. The effect of the IFN-τ peptides on the antiviral activity of ovine IFNτ was examined in a dose/response assay using Madin Darby bovine kidney (MDBK) cells challenged with vesicular stomatitis virus. The carboxy-terminal peptide oIFN-τ(139–172) was found to be the most effective inhibitor of antiviral activity. Three additional peptides, oIFN-τ(1–37), (62–92), and (119–150), also reduced IFN-τ antiviral activity. This suggested that multiple regions of the IFN-τ molecule interact with the Type I IFN receptor and elicit antiviral activity. These regions are underlined in Table 3. The data were consistent with studies of antiviral activity and receptor binding with IFN-α analogs demonstrating that 3 distinct sites, located in the amino-terminal, internal, and carboxy-terminal regions of the molecule, influenced human IFN-α activity [15]. To verify functional results using synthetic peptides, antipeptide antisera were produced [14]. All antipeptide antisera were reactive with the native molecule. Interestingly, antisera titers correlated with the hydropathic index of the peptide, rather than with the predicted surface accessibility of the specific region in the 3-D configuration. Consistent with the peptide studies, antisera against the same four regions of the molecule inhibited IFN-τ activity while antisera to other regions did not. Since IFN-τ and IFN-α bind to the same receptor, the ability of the IFN-τ synthetic peptides to block both bovine and human IFN-α was examined. Interestingly, only three of the four inhibitory peptides were effective competitors of IFN-α. Cross-inhibition of IFN-α by the internal and carboxy-terminal peptides was observed and suggested that these residues may adopt a similar conformation in both molecules and bind to a common site on the receptor. The aminoterminal peptide failed to reduce IFN-α function entirely. Thus, either the IFN-α amino-terminus has a much higher affinity for receptor or the IFN-τ aminoterminus binds a unique site on the receptor complex that may be associated with its unique properties. As expected, none of the peptides blocked the antiviral activity of IFN-τ, which interacts with a different receptor. Next, it was determined whether the same active regions of IFN-τ were involved in additional systems. The Type I IFN receptor on cells has been reported to be somewhat more promiscuous than on other cell types [16]; therefore, vesicular stomatitis challenge of Fc-9 cells was performed. Only the carboxyterminal peptide inhibited IFN-τ activity in this system [17]. This suggested that it was the carboxyterminus that was crucial to receptor interaction. In studies examining IFN-τ-treated feline immunodeficiency virus infected FeT-1 cells and human immunodeficiency virus-infected peripheral

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blood lymphocytes, peptide inhibition of IFN-τ antiretroviral activity implicated both the amino- and carboxy-termini as functionally important [17]. The structural basis of the antiproliferative activity of IFN-τ was also investigated. While multiple regions were again involved in IFN-τ antiproliferative activity, it was the area adjacent to the carboxy terminus, rather than the carboxy-terminus itself, which was the most crucial for antiproliferative activity, inhibiting cell division by blocking entry into the S phase of the cell cycle [18]. Since, for a particular IFN-α subtype, antiviral potency does not necessarily correlate with antiproliferative potency, localization of these functions in different domains of the molecule is not unexpected [19]. Within all known IFN-αs, the 8 amino acids from 139 to 147 are highly conserved. These residues are contained in both carboxy-terminal peptides, but while they may be involved in antiviral activity, they do not appear to be solely responsible for antiproliferative activity since the two peptides are not equivalent inhibitors of IFN-τ antiproliferative activity. This observation is consistent with inhibition of antiviral activity but not antiproliferative activity by a monoclonal antibody in this conserved region in human IFN-α and with the requirement for tyrosine at position 123 for human IFN-α1 antiproliferative activity [20,21]. It has also been reported that mutations around Arg33 affected both antiviral and antiproliferative activity of human IFN-α4 on human cells [22], while the amino-terminus did not appear to be as important in IFN-τ antiproliferative activity on bovine cells. B. IFN-τ Monoclonal Antibodies Another approach to structure/function analysis of IFN-τ involved generation of anti-IFN-τ monoclonal antibodies. Four monoclonal antibodies were produced that reacted with the native IFN-τ protein. They were epitope mapped using the available IFN-τ peptides. Two of the antibodies were directed against the carboxy-terminus of the molecule, one against a region adjacent to the aminoterminus, and the final one appeared to react with a conformational, rather than a linear determinant (C. Pontzer, unpublished data). When these antibodies were used as competitors in binding assays, all four inhibited IFN-τ binding to the Type I IFN receptor on MDBK cells. That anti-IFN-τ carboxy-terminal antibodies would inhibit binding is not unexpected, but the inhibitory activity of the monoclonal antibodies directed against the more amino-terminal region was not anticipated. There is the caveat that warns that results using monoclonal antibodies to delineate function sites must be interpreted with caution since their size may cause significant steric hindrance. To proceed further with structural studies of IFN-τ, access to larger quantities of pure protein was required. The obvious route to this end entailed production of recombinant protein. A synthetic gene for IFN-τ was designed

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that allowed for optimal expression in both bacterial and yeast systems [23]. In addition, restriction sites were incorporated at intervals throughout the length of the sequence to allow for cassette mutagenesis. Using the Pichia pastoris expression system, 50 mg of purified IFN-τ were obtained from a one-liter culture. C. IFN-τ Binding and Signal Transduction Detailed receptor-binding studies were performed comparing recombinant human IFN-α and IFN-τ [24]. The Kd of 125I-IFN-τ and 125I-IFN-αA for receptor on MDBK cells was 3.9 × 10-10 M and 4.45 × 10-11 M, respectively. Consistent with the higher binding affinity, IFN-αA was several fold more effective than IFN-τ as a competitive inhibitor. Functionally, the two IFNs had similar specific antiviral activities, but IFN-τ was 30 fold less toxic to MDBK cells at high concentrations. Phosphorylation of the signal transduction proteins, Tyk2, Stat1a, and Stat2 did not appear to be involved in the cellular toxicity associated with IFN-α relative to IFN-τ. Excess IFN-τ did not block the cytotoxicity of IFN-αA, suggesting that they recognize the receptor differently. While maximal IFN antiviral activity required only fractional receptor occupancy, toxicity was associated with maximal occupancy. Thus, “spare” receptors may exist with respect to certain biological properties, and IFNs may induce a concentrationdependent selective association of receptor subunits. D. Structural Biology of IFN-τ In order to better interpret the information derived from the above studies, an understanding of the 3-D structure of IFN-τ is required. Prior to resolution of the crystal structure, modeling techniques were employed for structural predictions [10]. For IFN-τ, the homology it shares with the other IFNs can be exploited. Since the x-ray coordinates for IFN-β are known [2] (Figure 3), it was used as a template for predicting the topology of IFN-τ. When the sequences of IFN-τ and IFN-β are aligned, the overall homology is approximately 30%. When residues are compared on the basis of conservative substitutions, the similarity rises to about 50%, and if only the location of hydrophobic residues is compared, the similarity is approximately 75%. This is important because hydrophobicity is thought to be a critical factor in driving protein folding. The interferons IFN-β, IFN-α-2, and several other cytokines including IL-2, IL-4, growth hormone, and GM-CSF belong to a family in which all share a four-helix bundle structural motif. Four-helix bundles exhibit a characteristic apolar periodicity in the α helices where every third or fourth residue is apolar, forming a hydrophobic strip down one side of the helix, which facilitates packing. The aligned helical regions of IFN-τ show the same apolar periodicity, suggesting a four-helix bundle motif.

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Figure 3 Stereo view of IFN-β crystal structure [2].

The structure of IFN-τ was also examined by CD [10]. Analysis of the IFN-τ spectra predicts that the secondary structural elements derived from CD spectra indicate approximately 70% α-helix. The remainder of the molecule is either predicted to be random or a combination of β sheet and turn. Since it is known that algorithms that predict secondary structures from CD spectra are most accurate at identifying α helices, we are confident that IFN-τ is mainly α helical. The CD spectra for the synthetic peptides of IFN-τ were also obtained. The peptides IFN-τ(1–37), IFN-τ(62–92), IFN-τ(119–150), and IFN-τ(139–172) all show the presence of α helix, while IFN-τ(34–64) and IFN-τ(90–122) are mainly random. The presence of an α helix in the peptides supports the CD analysis of the intact protein and also roughly indicates the location of helical segments. The secondary structure of IFN-τ, including the location of the α helices and loop region, was then predicted using a neural network-based computer program called PHD that relies on sequence alignments of all proteins related to the target sequence [25,26]. When this prediction is correlated with the CD data, peptides that possess considerable α helicity are predicted to contain entire helical segments, and conversely, peptides with little helicity are predicted to be within loop regions. A model of the 3-D structure of IFN-τ was constructed using a distance geometry-based homology modeling method with mouse IFN-β acting as a template. The distance constraints were generated between residues within IFN-τ that are homologous to residues of IFN-β. Dihedral-angle restraints of α helices were generated from the secondary-structure prediction of IFN-τ. No constraints were applied to the 13-residue carboxy tail of IFN-τ, which is absent in IFN-β, since it is likely to be flexible in a manner similar to other proteins such as IFN-γ. Additional distance constraints were added from putative disul-

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Figure 4 Stereo view of IFN-τ model. Highlighted sequences are from 1–37, 62–92, and 139–172.

fide bridges between residues 1 and 99 and residues 29 and 139. Several structures were generated using distance geometry routines, and the energy was minimized and averaged to yield a final model [16]. A similar model was built by Senda et al. (unpublished results) using a homology modeling method. This model was also built using the x-ray coordinates of IFN-β and shows a similar topology to the IFN-β three-dimensional structure (Figure 4). The most striking feature of both models is that those discontinuous regions, previously determined to be functionally important, are localized to one side of the molecule and found to be spatially contiguous (Figure 4). This observation is consistent with multiple binding sites on IFN-τ interacting simultaneously with the Type I IFN receptor and emphasizes the importance of structural modeling in the understanding and interpretation of functional data. III. Type II IFN A. Functional Sites on the IFN-γ Molecule The production of IFN-γ-neutralizing antibodies specific for an N-terminal peptide of human IFN-γ provided the first evidence that the N-terminus of IFN-γ contained an important functional site [27]. A similar approach was used to produce N-terminus-specific neutralizing antisera against murine IFN-γ [28]. Subsequent studies using IFN-γ synthetic peptides to map the epitope specificity of monoclonal antibodies to murine IFN-γ showed that N-terminal specific monoclonal antibodies neutralize IFN-γ antiviral activity [29]. In receptor-

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competition studies, murine IFN-γ N-terminal peptide consisting of residues 1–39 [IFN-γ (1–39)] blocked both binding to receptor and antiviral activity of IFN-γ [30]. Overlapping peptides of other regions of the IFN-γ molecule failed to block binding and function of IFN-γ [31]. Thus the combination of peptide mapping of epitope specificities and receptor competition using peptides has identified the Nterminus as a structurally and functionally important region of the IFN-γ molecule. This region is highlighted in the sequence of human IFN-γ found in Table 3. Interestingly, site-specific antibodies to the C-terminus of murine IFN-γ, which were induced using the peptide consisting of residues 95–133 [IFN-γ (95–133)], also neutralized IFN-γ activity, however IFN-γ (95–133) failed to block binding of IFN-γ to receptor and IFN-γ activity simultaneously. Antibodies to internal peptides failed to block both antiviral activity and binding of IFN-γ to receptor. In studies with recombinant murine IFN-γ receptor, which consisted of the entire α chain except for the transmembrane domain, the C-terminal peptide did block binding of IFN-γ to receptor [32]. Thus we have the interesting paradox wherein the IFN-γ C-terminal peptide blocked binding of IFN-γ to the recombinant, soluble receptor and yet did not block binding to the cell-surface receptor. One interpretation of these findings has allowed us to formulate the “velcro-key” model of binding to receptor that involves both N- and Cterminal domains of IFN-γ (Figure 5). The N-terminus binds in the “lock and key” manner characterized by specific ligand-receptor binding. The hydrophilic C-terminus binds to a region of the receptor distinct from that for the N-terminus, most likely through its polycationic region, which is conserved across species barriers. Binding of this type would exhibit high affinity and low specificity, similar to a piece of velcro. The C-terminal peptide of IFN-γ would therefore act as a poor competitor for cell-surface binding due to its low specificity alone. This interaction becomes specific in the context of the whole IFN-γ molecule and may increase the affinity of receptor binding. An alternative explanation that may also account for the inability of the C-terminal peptide to compete for cell-surface interactions is that its binding site is located not on the extracellular domain of the receptor, but rather on the intracellular domain. The primary differences between the cell-surface form of the IFN-γ receptor and (2) the accessibility of the recombinant receptor's cytoplasmic domain. A synthetic peptide corresponding to the membrane proximal region of the cytoplasmic domain of the murine IFN-γ receptor was able to bind IFN-γ and specifically compete with the binding of IFN-γ (95–133) to fixed/permeabilized cells [33]. Studies by others have reaffirmed the importance of both the N- and C-terminal regions of IFN-γ in function. Using recombinant DNA techniques, it has been shown that deletion of residues from the Nterminus of the molecule

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Figure 5 Velcro-key model of IFN-γ binding to its receptor. (From Reference 25. Copyright 1992. The American Association of Immunologists.)

results in decreased receptor binding [34]. Deletions or substitutions at the C- terminus have a direct effect on function of the molecule [35–38]. Epitope mapping of neutralizing monoclonal antibodies has also revealed an internal region of the molecule (from residues 84–94) as being functionally important [39]. This sequence bears strong homology to the nuclear localization sequence (NLS) of the SV40 large T antigen and has recently been demonstrated to be fully functional as an NLS for IFN-γ [40]. Thus, internal regions of the IFN-γ molecule are also likely to play an important functional role. B. IFN-γ Receptor α Chain Sites of Interaction with IFN-γ Both the human and the murine IFN-γ receptors consist of a ligand-binding subunit and a speciesspecific cofactor molecule. It is through interaction with this cell-surface receptor complex that IFN-γ exerts its biological effects. The IFN-γ molecule and its N-terminal peptide IFN-γ (1–39) bind specifically to the http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_447.html (1 of 2) [4/9/2004 12:11:36 AM]

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cell-surface receptor and to a recombinant, soluble form of the ligand-binding chain of the receptor [25]. Synthetic peptides corresponding to the sequence of the extracellular domain of the ligand-binding subunit were used to define the region of the receptor to which the N-terminus of IFN-γ binds. Receptor peptide MIR (95–120) competed most strongly with IFN-γ binding to both cell-surface and recombinant, soluble receptor [41]. Additionally, antisera to this peptide and the adjacent overlapping peptide, MIR (118–143), inhibited the binding of IFN-γ to the recombinant, soluble receptor. Therefore, the receptor domain responsible for binding the N-terminus of IFN-γ is defined by the region encompassing residues 95–120 of the ligand-binding subunit of the IFN-γ receptor and may extend further into the neighboring sequence. Antibodies to the C-terminal region of IFN-γ have been shown to be potent neutralizers of IFN-γ activity [29]. However, no cell-surface binding site for the C-terminus of IFN-γ could be localized using either antisera or synthetic peptides. Furthermore, as indicated above, the C-terminal IFN-γ peptide, IFN-γ (95–133), competed specifically with the intact IFN-γ molecule for binding to a recombinant, soluble form of the receptor, which consists of both the extracellular and the intracellular domains [42]. It was hypothesized that since the intracellular portion of the soluble receptor was accessible, in contrast to that of the cell-surface receptor, the C-terminus of IFN-γ might indeed be binding to this region. In studies using synthetic peptides corresponding to the cytoplasmic domain of the murine IFN-γ receptor, only peptide MIR (253–287) specifically bound both murine IFN-γ and its C-terminal peptide, MuIFN-γ (95–133) [33]. This peptide corresponds to the membrane proximal region of the receptor's cytoplasmic region. Antibodies to this receptor peptide inhibited the binding of the C-terminus of murine IFN-γ to the receptor in cells which had been fixed and permeabilized. Analogous binding studies with human IFN-γ and its C- terminal peptide, HuIFN-γ (95–134), yielded a similar result [43]. Surprisingly, the binding of the IFN-γ C-terminal peptides to their cytoplasmic binding sites is not species restricted, which is in contrast to the binding of the whole molecule at the cell surface. Both human and murine IFN-γ and their C-terminal peptides bound equally well to receptor peptides of either human or murine origin [43]. Thus, a receptor binding site for the C-terminus of the IFN-γ molecule has been localized to the membrane proximal region of the ligand-binding subunit's cytoplasmic domain (Figure 6). Previously, there have been several reports of human IFN-γ having activity on murine cells when administered cytoplasmically [44–46]. With the identification of a cytoplasmic binding site for IFN-γ, which is not species restricted, the question arose as to whether this might be the basis for these earlier observations. Thus, C-terminal IFN-γ peptides of both human and murine origin were used to stimulate murine macrophage lines P388D1 WEHI-3. Macrophages were chosen particularly for their capacity to nonspecifically endocytose material,

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Figure 6 Proposed receptor activation pathway for IFN-γ. (From Reference 53. Copyright 1995. The American Association of Immunologists.)

and we took advantage of this as a means of introducing the IFN-γ peptides into these cells. The IFN-γ Cterminal peptides induced a potent antiviral state in the murine macrophages and upregulated expression of MHC class II molecules, both in a dose-dependent fashion [43]. These effects were demonstrated to be sequence specific, as a scrambled version of the murine C-terminal peptide lacked activity. Furthermore, a truncated form of the murine C-terminal peptide, lacking the sequence of basic amino acids (RKRKR), was also without activity. The absence of activity of this truncated peptide was linked directly to a loss of its ability to bind to the receptor [43]. Therefore, interaction of IFN-γ, via its Cterminus, with its cytoplasmic binding site is important for function and requires the presence of a region of basic amino acids near the C-terminus of the molecule. C. Structural Biology of IFN-γ and the IFN-γ Receptor Structure-function studies of IFN-γ carried out using the synthetic peptide approach and site-specific antibodies indicated that both the N- and C-terminal regions of the protein were not only functionally important, but also accessible at the surface of the molecule. The x-ray crystal structure of human IFN-γ has been determined and reveals that in the IFN-γ homodimer both the N- and the

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Figure 7 Stereo view of IFN-γ [40]. Regions 1–39 of chain A and 95–119 of chain B of the dimer are highlighted.

C-terminus were indeed accessible [3]. Figure 7 illustrates the close proximity of the N- and C-termini of IFN-γ. The subunits of the homodimer are oriented head to tail, such that the N-terminal helix-loophelix (corresponding to residues 1–39) of one IFN-γ molecule interacts with the C-terminus of the second IFN-γ molecule. As mentioned above, synthetic peptides were also instrumental in identifying the region of the receptor to which the N-terminus of IFN-γ binds. Recently, the crystal structure of a complex between murine IFN-γ and the murine IFN-γ Rα subunit has been determined [5]. The synthetic peptides and corresponding antisera had predicted an interaction of murine IFN-γ residues (1–39) with receptor region (95–143). The crystal structure confirmed these observations, indicating an interaction of IFN-γ residues (1–42) with receptor residues (108–132). However, the crystal structure did not define an extracellular binding site for the C-terminus of IFN-γ. There has been some speculation that the basic amino acid residues of the C-terminus may interact with an acidic patch on the receptor's extracellular domain, which would support the previously mentioned “velcro-key” model, but that the crystallization conditions precluded this interaction [5]. It is quite possible that such an interaction may occur as a transitional state prior to the internalization of the C-terminal portion of IFN-γ and interaction with the cytoplasmic region of the receptor. An alternative explanation for the apparent lack of a binding site for the IFN-γ C-terminus on the receptor's extracellular face is that its primary site of interaction is

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with the cytoplasmic portion of the receptor as described above. Thus, the orientation of the C-terminal portion of the IFN-γ molecules in the receptor complex should be such that they are situated near to the cell membrane. When one examines the structure of the receptor-ligand complex, it is easy to see that this is indeed the case. Studies are currently under way to determine the crystal structure of the complex between the cytoplasmic domain of the IFN-γ receptor and the IFN-γ molecule, in particular the Cterminus. With regards to the definition of sites of interaction between receptors and ligands, the synthetic peptide approach has repeatedly proven to be an accurate indicator of structurally important regions of the IFN-γ/IFN-γ R system. D. Signal Transduction by IFN-γ Within the past several years some of the immediate-early signal transduction events initiated in response to IFN-γ stimulation have been elucidated. Treatment of cells with IFN-γ leads to the rapid activation of two protein tyrosine kinases, JAK1 and JAK2 [47]. The JAK kinases are a newly emerging family of protein kinases important in signaling via cytokines and growth factors. These proteins are unrelated to the src family of tyrosine kinases and are characterized as being larger, having two putative phosphotransferase domains and containing no characteristic SH2 or SH3 domains [48–51]. Members of the Janus kinase family are found associated with the cytoplasmic domains of cytokine and growth factor receptors at or near to the membrane proximal region [51]. In the resting cell, JAK1 and JAK2 are found associated with the α and β/AF-1 chains of the IFN-γ receptor, respectively [52]. These kinases as well as the ligand-binding chain of the IFN-γ receptor are tyrosine phosphorylated in response to IFN-γ treatment [47,53,54]. This leads in turn to the tyrosine phosphorylation of a latent cytoplasmic transcription factor, known variously as p91, Stat 91, or Stat 1 α on tyrosine residue 701 [55]. It is interesting to note that the IFN-α signal-transduction pathway partially overlaps with that of IFN-γ. Stimulation of cells by IFN-α leads to the activation of JAK1 and another Janus family kinase, Tyk2 [56,57]. In turn, this cascade leads to phosphorylation of two latent cytoplasmic transcription factors, p84 (Stat 1β) and p113 (Stat 2β) in addition to the p91 (Stat 1α) activated by IFN-γ. The identification of tyrosine kinases that directly associate with the subunits of the IFN-γ receptor lead to the question of how the binding of IFN-γ might affect these proteins. Recently, the synthetic peptide method was used to identify two regions of the murine IFN-γ receptor's α chain as being important for interaction with the kinase JAK2 [58]. One of these regions lies in the distal portion of the cytoplasmic tail (residues 404–432), while the other (residues 283–309) is nearer to the membrane proximal region to which the C-terminal part of IFN-γ binds (residues 253–287). The fact that there are adjacent binding sites for JAK2 and IFN-γ implied a potential for interaction between the IFN-γ

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ligand and the machinery of signal transduction, namely JAK2. It was found that both intact murine IFNγ and its C-terminal peptide (95–133) are capable of specifically mediating an increase in the degree of association between the recombinant, soluble IFN-γ receptor and JAK2. These findings were further supported as IFN-γ and IFN-γ (95–133) caused an increase in the amount of JAK2 coprecipitating with the receptor from intact murine macrophages [58]. This has been the first such demonstration of an extracellular cytokine ligand participating directly in interaction with cytoplasmic signaling elements. E. IFN-γ as a Candidate for Rational Drug Design The IFN-γ molecule is a potentially attractive model for the rational application of drug-design strategies. Reagents exist that are capable of either positively or negatively modulating the in vivo effects of IFN-γ. An initial target is quite simply at the level of receptor-ligand interaction. Synthetic peptide analogs of the N-terminal region have been successfully applied in vitro to inhibit interaction of intact IFN-γ with cell-surface receptors [30]. Interestingly, it has been observed that the N-terminal region of mouse IFN-γ has the ability to interact with the human receptor [59]. It was shown that the mouse peptide IFN-γ (1–39) had a 10-fold greater ability to block the binding of human IFN-γ to cellsurface receptors. This was shown to be correlated with a more stable structure in solution for the murine peptide and illustrates the importance of stable structure to receptor binding, which may be exploited when designing peptide mimetics. The solution structure of this peptide has also been determined and could provide the beginning steps for determining the structural requirements of an antagonist [7]. Future studies could focus on cocrystallization of the peptides with receptor or NMR studies of peptide domains of the receptor and IFN-γ. Recombinant, soluble forms of the extracellular domain of the ligand-binding subunit of the receptor have also been used in analogous fashion both in vitro to inhibit cell-surface binding and in vivo to interfere with disease progression [60]. Therefore prevention of potentially deleterious effects of IFN-γ may be achieved by preventing initial interactions with receptor molecules at the cell surface. A second candidate region lies within amino acid residues 84–94 of human IFN-γ and the corresponding region of its murine homologue. This portion of the molecule functions as a nuclear localization signal and, therefore, is also an attractive target for drug design. In cells treated with IFN-γ, the IFN-γ molecule traffics rapidly to the nucleus of the cell, usually within one to two minutes. When the IFN-γ molecule is crosslinked to its receptor, the resultant receptor-ligand complex migrates to the nucleus [40]. The implication is that this sequence may therefore be of use in artificially targeting proteins from the cytoplasm directly to the nucleus. This is potentially a very attractive method

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for directed subcellular targeting of synthetic transcriptional activators or repressors that might not otherwise be directed to the nucleus. Finally, the C-terminal region of IFN-γ offers another possibility. With respect to the induction of an antiviral state, or the upregulation of MHC class II molecule expression, synthetic peptides corresponding to the C-terminal 39 amino acids of either human or murine IFN-γ function as potent agonists. This activity is due to the interaction of these peptides with the cytoplasmic domain of the IFNγ receptor and the associated protein tyrosine kinases. Furthermore, in contrast to the stringent species specificity of intact IFN-γ, the action of the C-terminal peptides agonists is not limited by species constraints. This property therefore renders the C-terminal portion of the IFN-γ molecule an attractive model for the development of IFN-γ agonists and antagonists. The identification of the C-terminus of IFN-γ as that part of the molecule that contacts the cytoplasmic portion of the receptor implies that in developing IFN-γ agonists, the primary focus should be on this region of the protein. Corresponding antagonists may be developed based upon the portion of the receptor to which the IFN-γ C-terminus binds. IV. Conclusion The interest in IFNs as therapeutics has existed from their initial discovery in 1957. Since then scientists have been trying to understand the mechanism of their action and apply that knowledge to the treatment of many different diseases, meeting with some success. The effort now is to understand how IFNs work at the molecular level, with the goal being to design better, more specific therapeutics. Through structure/function studies, we now know where the functional sites lie on many of the IFNs. We also know the sites of interaction with their receptors and second messenger systems. From these studies, initial candidates for structure-based drug design have been identified. Although more work is needed to further characterize the IFNs and their receptor systems, the challenge now is to apply our existing knowledge and create second generation molecules that can modulate the many activities of these fascinating proteins. References 1a. Isaacs A and Lindenmann J. Virus Interference. I. The interferon. Proc R Soc London Ser B 1957; 147:258. 1b. Johnson HM, Bazer FW, Szente BE, Jarpe MA. How interferons fight disease. Scientific American 1994; 270:40–47. 2. Senda T, Shimazu T, Matsuda S, Kawano G, Shimizu H, Nakamura KT, Mitsui Y. Three-dimensional crystal structure of recombinant murine interferon-β. EMBO J 1992; 11:3193–3201.

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3. Ealick SE, Cook WJ, Vijay-Kumar S, Carson M, Nagabhushan TL, Trotta PP, Bugg CE. Threedimensional structure of recombinant human interferon-g. Science 1991; 252:698–702. 4. Samudzi CT, Burton LE, Rubin JR. Crystal structure of recombinant rabbit interferon-gamma at 2.7 A resolution. J Biol Chem 1991; 266:21791–21797. 5. Walter MR, Windsor WT, Nagabhushan TL, Lundell DJ, Lunn CA, Zavodny PJ, Narula SK. Crystal structure of a complex between interferon-g and its soluble high-affinity receptor. Nature 1995; 376:230–235. 6. Grzesiek S, Dobeli H, Gentz R, Garotta G, Labhardt AM, Bax A. 1H, 13C, and 15N NMR backbone assignments and secondary structure of human interferon-gamma. Biochemistry 1992; 31 (35):8180–90. 7. Sakai TT, Jablonski MJ, DeMuth PA, Krishna NR, Jarpe MA, Johnson HM. Proton NMR sequence specific assignments and secondary structure of a receptor binding domain of mouse γ-interferon. Biochemistry 1993; 32:5650. 8. Murgolo NJ, Windsor WT, Hruza A, Reichert P, Tsarbopoulos A, Baldwin S, Huang E, Pramanik B, Ealick S, Trotta PP. A homology model of human interferon alpha-2. Proteins 1993; 17(1):62–74. 9. Seto MH, Harkins RN, Adler M, Whitlow M, Church WB, Croze E. Homology model of human interferon-alpha-8 and its receptor complex. Protein Science 1995; 4:655–70. 10. Jarpe MA, Johnson HM, Bazer FW, Ott TL, Curto EV, Rama Krishna N, Pontzer CH. Predicted structural motif of IFN-τ. Protein Engineering 1994; 7:863–867. 11. Bazer FW. Mediators of maternal recognition of pregnancy in mammals. Proc Soc Exp Biol Med 1992; 199:373–384. 12. Imakawa K, Anthony RV, Kazemi M, Marotti KR, Polites HG, Roberts RM. Interferon-like sequence of ovine trophoblast protein secreted by embryonic trophectoderm. Nature 1987; 330:377–379.

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13. Stewart HJ, McCann SHE, Barker PJ, Lee KE, Lamming GE, Flint APF. Interferon sequence homology and receptor binding activity of ovine trophoblast antileuteolytic protein. J Endocrinol 1987; 115:R13–R15. 14. Pontzer CH, Ott TL, Bazer FW, Johnson HM. Structure/function studies with interferon tau: Evidence for multiple active sites. J Interferon Res 1994; 14:133–141. 15. Fish EN, Banerjee K, Stebbing N. The role of three domains in the biological activity of human interferon-α. J Interferon Res 1989; 9:97–114. 16. Novick D, Cohen B, Rubinstein M. The human interferon α/β receptor: characterization and molecular cloning. Cell 1994; 77:391–400. 17. Pontzer CH, Yamamoto JK, Bazer FW, Ott TL, Johnson HM. Potent anti-feline immunodificiency virus and anti-human immunodeficiency virus effect of interferon tau. J Immunol 1995; (in press). 18. Pontzer CP, Bazer FW, Johnson HM. Antiproliferative activity of a pregnancy recognition hormone, ovine trophoblast protein-1. Cancer Res 1991; 51:5304–5307. 19. Pestka S, Langer JA, Zoon KC, Samuel CE. Interferons and their actions. Annu Rev Biochem 1987; 56:727–777. 20. Barasoain I, Portolès A, Aramburu JF, Rojo JM. Antibodies against a peptide representative of a conserved region of human IFN-α. Differential effects on the antiviral and antiproliferative effects of IFN. J Immunol 1989; 143:507–512.

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21. McInnes B, Chambers PJ, Cheetham BF, Beilharz MW, Tymms MJ. Structure-function studies of interferons-α: Amino acid substitutions at the conserved residue tyrosine 123 in human interferon-α1. J Interferon Res 1989; 9:305–314. 22. Waine GJ, Tymms MJ, Brandt ER, Cheetham BF, Linnane AW. Structure-function study of the region encompassing residues 26–40 of human interferon-α4: Identification of residues important for antiviral and antiproliferative activities. J Interferon Res 1992; 13:42–48. 23. Ott TL, Van Heeke G, Johnson HM, Bazer FW. Cloning and expression in Saccharomyces cerevisiae of a synthetic gene for the type-1 trophoblast interferon ovine trophoblast protein-1: Purification and antiviral activity. J Interferon Res 1990; 11:357–364. 24. Subramaniam PS, Khan SA, Pontzer CH, Johnson HM. Differential recognition of the type In IFN receptor by IFN-τ and IFN-α is responsible for their differential cytotoxicities. 1995; (submitted). 25. Sander C, Schneider R. Database of homology-derived structure and the structural meaning of sequence alignment. Proteins 1991; 9:56–68. 26. Rost B, Sander J. Prediction of protein structure at better than 70% accuracy. J Mol Biol 1993; 232:544–599. 27. Johnson HM, Langford MP, Lakchaura B, Chan TS, Stanton GJ. Neutralization of native human gamma interferon by antibodies to a synthetic peptide encodded by the 5' end of human gamma interferon cDNA. J Immunol 1982; 129:2357–2359. 28. Langford MP, Gray PW, Stanton GJ, Lakchaura B, Chan T-S, Johnson HM. Antibodies to a synthetic peptide corresponding to the N-terminal end of mouse gamma interferon (IFN-α). Biochem Biophys Res Comm 1983; 117:866–871. 29. Russell JK, Hayes MP, Carter MJ, Torres BA, Dunn BM, Russell SW, Johnson HM. Epitope and functional specificity of monoclonal antibodies to mouse interferon gamma: the synthetic peptide approach. J Immunol 1986; 136:3324–3328. 30. Magazine HI, Carter JM, Russell JK, Torres BA, Dunn BM, Johnson HM. Use of synthetic peptides to identify an N-terminal epitope on mouse gamma interferon that may be involved in function. Proc Natl Acad Sci USA 1988; 185:1237–1241. 31. Jarpe MA, Johnson HM. Topology of receptor binding domains of mouse IFN-α. J Immunol 1990; 145:3304–3309. 32. Griggs ND, Jarpe MA, Pace JL, Russell SW, Johnson HM. The N-terminus and C- terminus of interferon gamma are binding domains for cloned soluble interferon gamma receptor. J Immunol 1992; 149:517–520. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_455.html (1 of 2) [4/9/2004 12:12:22 AM]

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33. Szente BE, Johnson HM. Binding of IFN-α and its C-terminal peptide to a cyto-plasmic domain of its receptor that is essential for function. Biochem Biophys Res Comm 1994; 201:215–221. 34. Zavodny PJ, Petro ME, Chiang TR, Narula SK, Leibowitz PJ. Alterations of the amino terminus of murine interferon gamma: expression and biological activity. J Interferon Res 1988; 8:483–494. 35. Arakawa T, Hsu YR, Parker CG, Lai PH. Role of polycationic C-terminal portion in the structure and activity of recombinant human interferon gamma. J Biol Chem 1986; 261:8534–8539. 36. Leinikki PO, Calderon J, Luquette MH, Schreiber RD. Reduced receptor binding by a human interferon gamma fragment lacking 11 carboxyl-terminal amino acids. J Immunol 1987; 139:3360–3366. 37. Wetzel R, Perry LJ, Veilleux C, Chang G. Mutational analysis of the C-terminus of human interferon gamma. Prot Eng 1990; 3:611–623.

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38. Lundell D, Lunn C, Dalgarno D, Fossetta J, Greenberg R, Reim R, Grace M, Narula S. The carboxylterminal region of human interferon-gamma is important for biological activity: mutagenic and NMR analysis. Prot Eng 1991; 4(3):335–341. 39. Zu X, Jay FT, The E1 functional epitope of the human interferon-α is a nuclear targeting signal-like element. J Biol Chem 1991; 266:6023–6026. 40. Bader T, Wietzerbin J. Nuclear accumulation of interferon-gamma. Proc Natl Acad Sci USA 1994;91:11831–11835. 41. Van Volkenburg MA, Griggs ND, Jarpe MA, Pace JL, Russel SW, Johnson HM. Binding site on the murine interferon-gamma receptor for interferon-gamma has been identified using the synthetic peptide approach. J Immunol 1993; 151:6206– 6213. 42. Fernando LP, LeClaire RD, Obici S, Zavodny PJ, Russell SW, Pace JL. Stable expression of a secreted form of the mouse IFN-α receptor by rate cells. J Immunol 1991; 147:541–547. 43. Szente BE, Soos JM, Johnson HM. The C-terminus of IFN-α is sufficient for intracellular function. Biochem Biophys Res Comm 1994; 203:1645–1654. 44. Fidler IJ, Fogler WE, Kleinerman ES, Saiki I. Abrogation of species specificity for activation of tumoricidal properties in macrophages by a recombinant mouse or human interferon gamma encapsulated in liposomes. J Immunol 1985; 135:4289–4296. 45. Sancéau J, Sondermeyers P, Béranger F, Falcoff R, Vaquero C. Intracellular human interferon triggers an antiviral state in transformed murine L cells. Proc Natl Acad Sci USA 1987;84:2906–2910. 46. Smith MR, Muegge K, Keller JR, Kung HF, Young HA, Durum SK. Direct evidence for an intracellular role for interferon-gamma. J Immunol 1990; 144:1777–1782.

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47. Igarashi K, Garotta G, Ozmen L, Ziemiecki A, Wilks AF, Harpur AG, Larner AC, Finbloom DS. Interferon gamma induces tyrosine phosphorylation of interferon gamma receptor and regulated association of protein tyrosine kinases, Jak1 and Jak2, with its receptor. J Biol Chem 1994; 269:14333–14336. 48. Firmbach-Kraft I, Byers M, Shows T, Dalla-Favera R, Krolewski JJ. Tyk2, prototype of a novel class of non-receptor tyrosine kinase genes. Oncogene 1990; 5:1329–1336. 49. Bernards A. Predicted tyk2 protein contains two tandem protein kinase domains. Oncogene 1991; 6:1185–1187. 50. Wilks AF, Harpur AG, Kurban RR, Ralph SJ, Zurcher G, Ziemiecki A. Two novel protein tyrosine kinases, each with a second phosphotransferase-related catalytic domain, define a new class of protein kinase. Mol Cell Biol 1991; 11:2057–2065. 51. Ihle JN, Witthuhn BA, Quelle FW, Yamamoto K, Silvennoinen O. Signaling through the hematopoietic cytokine receptors. Annu Rev Immunol 1995; 13:369–398. 52. Sakatsume M, Igarashi K-I, Winestock KD, Garotta G, Larner AC, Finbloom DS. The Jak Kinases differentially associate with the a and b (accessory factor) chains of the interferon-g receptor to form a functional receptor unit capable of activating STAT transcription factors. J Biol Chem 1995; 270:17528–17534. 53. Khurana Hershey GK, McCourt DW, Schreiber RD. Ligand-induced phosphorylation of the human interferong receptor. J Biol Chem 1990; 265:17868–17875.

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54. Greenlund AC, Farrar MA, Viviano BL, Schreiber RD. Ligand induced interferon-gamma receptor tyrosine phosphorylation couples the receptor to its signal transduction system (p912). EMBO J 1994; 13:1591–1600. 55. Shuai K, Start GR, Kerr IM, Darnell JE. A single phosphotyrosine residue of Stat91 required for gene activation by interferon gamma. Science 1993; 261:1744–1746. 56. Muller M, Briscoe J, Laxton C, Guschin D, Ziemiecki A, Silvennoinen O, Harpur AG, Barbieri G, Witthuhn BA, Schindler C, Pellegrini S, Wilks AF, Ihle JN, Stark GR, Kerr IM. The protein tyrosine kinase JAK1 complements defects in interferon alpha/beta and gamma signal transduction. Nature 1993; 366:129–135. 57. Barbieri G, Velazquez L, Scrobogna M, Fellous M, Pellegrini S. Activation of the protein kinase tyk2 by interferon α/β. Eur J Biochem 1994;223:427–435. 58. Szente BE, Subramaniam PS, Johnson HM. Identification of IFN-γ receptor binding sites for JAK2 and enhancement of binding by IFN-γ and its' C-terminal peptide IFN-γ(95–133). J Immunol 1995; 155:95–133. 59. Jarpe MA, Johnson HM. Stable conformation of IFN-γ receptor binding peptide in aqueous solution is required for IFN-γ antagonist activity. J Interferon Res 1993; 13:99. 60. Ozmen L, Roman D, Fountoualakis M, Schmid G, Ryffel B, Garotta G. Soluble interferon-gamma receptor: a therapeutically useful drug for systemic lupus erythematosus. J Interferon Res 1994; 14(5):283–284.

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18 The Design of Anti-Influenza Virus Drugs from the X-ray Molecular Structure of Influenza Virus Neuraminidase Joseph N. Varghese Biomolecular Research Institute, Melbourne, Victoria, Australia I. Introduction Influenza has plagued humankind since the dawn of history and continues to affect a significant proportion of the population irrespective of age or previous infection history. These periodic epidemics that reinfect otherwise healthy people have devastated communities world wide. Some pandemics like the 1917–1919 “Spanish flu” were responsible for the death of tens of millions of people throughout the world. The origins, spread, and severity of influenza epidemics have been a puzzle that has only in the last two decades been adequately addressed. In early times it was thought that the disease was the evil influence (sic) of the stars, and other extraterrestial objects. At present it is generally accepted that the disease is of viral origin, spread by aerosols produced by infected animals, and the continual production of new strains of the virus results in reinfection of the disease (reviewed in Reference 1). There are three types of influenza virus classified on their serological cross-reactivity with viral matrix proteins and soluble nucleoprotein (A, B, and C). Only type A and B are known to cause severe human disease. Type B is only found in humans, while type A has a natural reservoir in birds and some mammals like pigs and horses [2]. Influenza, an orthomyxovirus, is a 100 nm lipid-enveloped virus (Figure 1) containing an eight-segment negative single-stranded genome [3]. Two of the segments code for the surfaces glycoproteins, hemagglutinin (which binds to terminal sialic acid), and neuraminidase (which

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Figure 1 A schematic diagram of an influenza virus particle that illustrates its constituent components and morphology. The surface antigens hemagglutinin and neuraminidase are attached to the lipid and matrix protein shell that encapsulates the eight negative-stranded RNA genes of the virus and associated nucleoprotein and polymerase.

cleaves terminal sialic acid) and which appear as spikes protruding out of the viral envelope. The viral target in humans is the upper respiratory tract epithelial cells. Replication (see Figure 2) begins with penetration of the virion through the mucin layer covering the epithelial surface, followed by attachment to the viral receptor by the hemagglutinin. Penetration of the cell is achieved by endocytosis and the virion core is released after the fusion of the virion and vesicle membrane mediated by the hemagglutinin. Fusion is enabled by a conformational change in the hemagglutinin made possible by lowering the pH of the endosome by the M2 ion channel protein. Following replication, the progeny virions are released by budding off the cell membrane [4,5].

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Figure 2 A simplified schematic of the replication cycle of an influenza virion in the host respiratory epithelia. Details of viral transport through mucin and pathways of viral spread on budding from the epithelial cell membrane during replication is not understood. Neuraminidase activity is important for release of budding progeny virions, desialyation of viral glycoproteins, and probably facilitates transport through sialic-acidrich mucin.

Release of virions occur 8 hours post infection and the onset of infection is sudden, resulting in pyroxia, muscular and joint pain, and a dry cough [6]. Virus shedding continues for up to a week, when a rise in virus-specific antibody clears the virus from the host. The vulnerability of the host succumbing to viremea during this week of rising viral titer is mediated by interferon induction [7] 48 hours post infection, which attenuates viral replication until the cell-mediated immune response begins to clear the virus. The severity of the illness is thought to depend on the level of cross protection arising from antibodies raised from previous influenza infections [19]. The course of the illness can be debilitating, and no effective treatment is available at present to halt the progression of the disease. Death can result for susceptible populations (neonate and elderly) primarily as a result of secondary infections [8]. This chapter shall examine a structural basis for the continual emergence of new influenza strains, and the reasons current vaccines against influenza fail to protect against all

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strains of influenza. The discovery of the active site of influenza neuraminidase and the exploitation of its structural conservation shall be discussed in terms of the design of potent neuraminidase inhibitors. The potential therapeutic use of these inhibitors as antiviral drugs against influenza virus infections shall be examined. A. Antigenic Variation The plethora of different strains of virus that are responsible for the continued reinfection of virus in humans is primarily related to mutations in the viral genes of two surface glycoproteins, hemagglutinin and neuraminidase [9]. The current paradigm for this genetic variation [10,11] is that these mutations arise primarily from incremental changes in the amino acid sequences of these glycoproteins by selection pressure of the immune system of the infected host. This mechanism termed “Antigenic Drift” accounts for most of the strain variation within a particular subtype of influenza. However, infrequently a mutation arises by genetic reassortment of viruses from different animal hosts (“Antigenic Shift”) whereby an entirely new gene for one of the surface glycoproteins is generated that is significantly different (~50%) in amino acid sequence from the parent virus. This is the mechanism by which new subtypes of influenza arise and are primarily responsible for the major pandemics that occur. Strains of influenza virus are classified by type (A, B, or C), geographic location, date of original isolation, and the subtype of the hemagglutinin and neuraminidase antigens. There exist 9 known subtypes (N1 to N9) of neuraminidase and 13 known subtypes (H1 to H13) of hemagglutinin for influenza A in all animal populations. Two neuraminidase (N1 and N2) and three hemagglutinin (H1, H2, and H3) subtypes of influenza A have occurred in strains that have infected humans since 1933 when isolates were first characterized [12]. Prior to 1933 there is indirect evidence of antigenic shift occurring in human populations [13]. The N1 subtype was associated with virus isolated between 1933 and 1957, after which time the N2 subtype appeared in the Asian influenza. No major change in the structure of neuraminidase has occurred since, although the hemagglutinin subtype has changed from H2 to H3 in 1968 in the Hong Kong pandemic, and H1N1 reappeared in 1978 as the Russian pandemic. Influenza B, which infects only human hosts, has only one subtype, but like type A undergoes continual antigenic drift. B. Current Therapeutics Amantidine and Rimantidine are the only class of drugs that have been approved for therapy. At high concentration (>50 mg/mL) Amantidine is thought to buffer the pH of the endosome and prevent the conformational

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change of the hemagglutinin necessary for fusion. Drug-resistant mutants arise where the hemagglutinin trimers are thought to be less stable than the wild type [14]. At low concentrations (<1 mg/mL) Amantadine blocks the activity of the viral M2 ion channel protein, which plays a role in virus uncoating and glycoprotein maturation [15]. Amatadine prevents fusion by altering the ability of the virus to change the ion balance [16] within the endosome and the trans-Golgi network. However Amatadine rapidly gives rise to drug-resistant strains by mutations in the hemagglutinin and the M2 protein that circumvent or block the activity of the drug. Rimantadine has been shown to lead to drug resistance in humans 2 days post treatment [17]. C. Influenza Vaccines Influenza virus vaccines [18,6] prepared from killed (formalin inactivated) virus are used currently worldwide as the only prophylactic treatment available against the disease. Killed vaccines contain whole virus or fractionated subunits. These vaccines attempt to incorporate antigens from influenza strains that will circulate in the community during an expected outbreak, conferring immunity to viral strains that are closely related to the strains the vaccine is made from. However antigenic drift reduces the susceptibility of the virus to neutralization by antibodies raised by immunization. For example, commercial vaccines containing inactivated A/Beijing/353/89 H3N2 strain circulating among humans from 1990 to 1993 provided significantly less protection [20] against the antigenic drift [21] variant A/Georgia/03/93. A more radical vaccination treatment, involving the direct injection into muscle of plasmid, DNAencoding hemagglutinin and M1 influenza viral proteins has been attempted successfully on animal models [22]. This is thought to involve incorporating the plasmid into muscle cells and eliciting a cellmediated immune (CMI) response that offers cross protection overcoming antigenic drift in the virus. The success of the method arises from a CMI response to segments of conserved amino acids in these proteins. Problems arising from possible autoimmune response to DNA and problems in ensuring the exogenous DNA is not integrated into the cell genome or sensitive cell lines elsewhere in the host have precluded testing in humans at present. Furthermore, mutations in the nuclear proteins of influenza (which have been relatively stable to date) arising from CMI selection pressure could undermine the efficacy of this technique. II. Viral Neuraminidase Activity Enzyme activity on the surface of influenza virus was first detected by Hirst [23], who observed that red blood cells once agglutinated by influenza virus

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could not again be agglutinated by either the eluted virus or fresh virus preparations. This activity is now attributed to neuraminidase, which is one of the two integral membrane glycoproteins of influenza virus (for reviews see References 24 and 25). A. Function Neuraminidase is an exoglycosidase that destroys the hemagglutinin receptor by cleaving the αketosidic linkage of terminal sialic acid [(N-actylneuraminic acid (Neu5Ac))] to an adjacent sugar [26,27]. Viral hemagglutinin binds specifically to Neu5Ac-containing receptors on the surface of susceptible cells [28]. Neuraminidase, which also removes terminal sialic acid from a range of glycoconjugates, plays an important, but not completely understood, role in the viral replication cycle. Without neuraminidase activity viruses [29] were thought to be immobilized by mucosal secretions in the upper respiratory tract. By removing terminal sialic acid from the sialic-acid-rich mucous layer [27,30] protecting target cells, neuraminidase could facilitate penetration of the virus to the cell surface. It has been shown that neuraminidase-deficient virus [31] can still replicate in vivo, albiet at a much reduced rate [32]. This shows that neuraminidase does not play an essential role in viral entry, replication, assembly or budding in mice, but has an important role in the spread of the infection by preventing aggregation at the cell surface and possible immobilization in the mucin by hemagglutinin. Once replication is initiated in the infected cell, the freshly synthesised viral glycoproteins have to be desialylated to prevent self-aggregation at the infected host cell surface by hemagglutinin binding to terminal sialic acid on these glycoproteins. Finally on elution of progeny virions from infected cells, neuraminidase activity is required to facilitate viral escape from the cell surface. Inactivation or inhibition of neuraminidase during budding has been observed to result in aggregation of virions on the cell surface [33–35]. Inhibition of this glycohydrolase could provide a means of controlling this disease by slowing the rate of viral attachment and subsequent release of progeny virions allowing the host immune system to eliminate the virus while the number of infected cells is low. B. Morphology There are between 50 to 100 neuraminidase spikes per virion [36] which is approximately 10% of the visible spikes projecting out of the surface of the virion [37]. These spikes can be removed from the virus by treatment with detergent [38]. Electron microscopic images of the neuraminidase spikes [39] reveal a mushroom-shaped molecule made up of a boxlike head of about 80 ×

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80 × 40 Å with a narrow centrally attached stalk (15 Å wide and 100 Å long), which terminates into a hydrophobic knob anchored in the viral envelope. The detergent-released spikes can be digested by pronase to release the neuraminidase “heads,” which retain full antigenic and enzyme activity [40,41]. The molecule was found to be a tetramer of molecular weight 240,000, reducing to 200,000 when treated with pronase [42]. A low resolution x-ray image of crystallized neuraminidase heads [43] established that the enzyme had circular 4-fold symmetry. III. Molecular Structure of Neuraminidase Crystals of pronase-released heads of the N2 human strains of A/Tokyo/3/67 [44] and A/RI/5+/57 were used for an x-ray structure determination. The x-ray 3-dimensional molecular structure of neuraminidase heads was determined [45] for these two N2 subtypes by a novel technique of molecular electron density averaging from two different crystal systems, using a combination of multiple isomorphous replacement and noncrystallographic symmetry averaging. The structure of A/Tokyo/3/67 N2 has been refined [46] to 2.2 Å as has the structures of two avian N9 subtypes [47–49]. Three influenza type B structures [50] have also been determined and found to have an identical fold with 60 residues (including 16 conserved cysteine residues) being invariant. Bacterial sialidases from salmonella [51] and cholera [52] have homologous structures to influenza neuraminidase, but few of the residues are structurally invariant. A. Structural Topology The protein fold consists of a symmetric arrangement of six four-stranded antiparallel β sheets arrange as blades of a propeller (Figure 3), the propeller axis being approximately parallel to but titled away from the circular 4-fold axis of the tetramer. This tilt angle varies between the known subtypes. This topology has now also been found in the seven β sheet propeller structure of bacterial methylamine dehydrogenase [53] and galactose oxidase [54], and the eight β sheet propeller structure of methanol dehydrogenase [55]. Each sheet of neuraminidase has a “W” topology (+1,+1,+1) [56] with four strands connected by reverse turns (Figure 3). The first strand of each sheet enters from the top, approximately parallel to and near the propeller axis; the fourth strand exits from the bottom, approximately perpendicular to the propeller axis. Top and bottom surfaces of the head refer to the faces of the tetramer away and towards the viral membrane respectively. Each sheet thus has a characteristic 90° right-hand twist between the inner- and outermost strand. The six sheets and their connections to each other are topologically identical.

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Figure 3 (a) A MOLSCRIPT [107] stereo diagram of the neuraminidase tetramer viewed from above down the symmetry axis. The different shaded arrows represent β strands comprising the six β sheets that form the “propeller” framework of each subunit. (b) A stereo diagram of a neuraminidase monomer viewed down the 4-fold axis, and α-sialic acid is shown bound in the active site of the enzyme, which lies in a pocket formed by the six β sheets near the pseudo six-fold axis of each subunit.

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The first strand of each sheet is connected across the top of the submit to the fourth strand of the preceding sheet. The N-terminus lies across the bottom of the subunit, and builds the fourth strand of the sixth sheet before entering the first sheet. The C-terminus strand builds the third strand of the sixth sheet, enters the subunit interface from the bottom, and runs parallel with the outermost (fourth) strand of the sixth β sheet.

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B. Protein Structure The 6 β sheets of the subunit, although topologically identical, vary in size and detailed structure. However, the six sheets can be considered as maintaining an approximate 6-fold symmetry relationship to each other about an axis parallel to the mean direction of the first strand of each sheet and through the centroid of these directions. Four disulfide bridges are formed between adjacent sheets as well as two between the sheets, which distort the sheet structure. Similar pairing has now been found in one of the domains of CD4 [57]. Also short β bulges occur on the inner and outer strands of most of the sheets. The packing of the β sheets is stabilized by hydrophobic interactions at the sheet interfaces and hydrogen bonding at the periphery of the sheets. In addition the intersheet disulfide bonds and ion pairs stabilize the structure. In general the loops connectioning the β sheets are shorter and tighter on the bottom of the subunit compared to the outer loops, which tends to be more elaborate. The intersheet loop connecting the fourth and fifth loop is the most extensive in the structure and is stabilized primarily by a disulfide bridge between Cys318 (sequence numbering of A/Tokyo/3/67 will be used throughout) and Cys337, a conserved ion pair Asp330—Arg364, and a putative Ca2+ ion binding site. This calcium binding site has approximate octahedral coordination with the main-chain oxygens of residues 293, 297, 345, and 348, a carboxylate oxygen of Asp324, and a water molecule in N2 and type B neuraminidases. In N9, the mainchain oxygen at 343 is replaced by a water molecule. Calcium has been shown to be necessary for neuraminidase activity [58] and this site is connected to the active site via conserved residues; however, the functional role of calcium in the structure is unknown, although it has been shown that calcium is essential for the thermostability of the molecule [59]. C. Carbohydrate Structure Carbohydrate at four N-linked glycosylation sites were observed in N2 neuraminidase at residues 86, 146, 200, and 234 in the x-ray structure. Two N-acetylglucosamines were resolved at Asp86 and Asp234, both at the bottom surface of the monomer. The carbohydrate at Asp200 consists of eight sugar residues with linkages consistent with known mannose-rich simple N-linked carbohydrates [60]. This oligosaccharide emerges from the side of the monomer and covers a neighboring subunit (see Figure 4). The oligosaccharide site at Asn146 is the most conserved of all neuraminidase glucosylation sites, except that of the neurovirulent virus A/WSN/33 [61]. The absence of this glycosylation site in A/WSN/33 has been shown to confer neurovirulence in mice [62]. It is a complex sugar containing Nacetylgalactosamine [63] that is not found in any other of the known oligosaccharides of influenza virus glycoproteins and is the only glycopeptide antigenically related to chick embryo “host antigen”

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Figure 4 A CPK model of two tetramers of A/Tokyo/3/67 neuraminidase heads as seen in the crystal structure. The four carbohydrate spikes emanating from the top of the molecule (at Asn146) interdigitate and form an open “barrel” structure. They form an unusual crystal contact. The dark spheres represent carbohydrate and the light spheres represent protein. The carbohydrate at Asn200 starts from one subunit and covers a neighboring subunit on the same tetramer. The other carbohydrates lie on the underside of the tetramer.

[63,64]. The oligosaccharide appears as a spike emanating from the top of the monomer, forming a crystal contact with a neighboring tetramer in crystals of A/Tokyo/3/67 neuraminidase. The four carbohydrate spikes of a tetramer form an open “barrel” structure of eight carbohydrate chains with the neighboring tetramer, with no apparent intercarbohydrate contacts. This oligosaccharide may play an important but as yet unidentified role in neuraminidase structure or activity. D. Antigenic Variation in Neuraminidase Structures Comparison of all known sequences of approximately 390 residues of the neuraminidase globular head [24], indicates that only 54 (excluding 16 conserved cysteine residues) are invariant (Figure 5a). Apart from 21 residues involved

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Figure 5 (a) Stereo image of a CPK atomic model of an influenza virus neuraminidase tetrameric head viewed distal to the viral membrane. The darker-shaded atoms represent totally conserved residues that for the most part form the enzyme active-site pocket. The lighter shaded atoms represent strain-variable residues and carbohydrate. (b) A stereo image of the enzyme active-site pocket of a subunit of neuraminidase with the same shading scheme.

in preserving the structural integrity of the molecule [46], the main clustering of these invariant residues is within the enzyme active site (Figure 5b), where 17 are in the active site and 16 are neighboring the active site. This is a cavity on the upper surface of the molecule into which sialic acid has been observed to bind [65,66,50]. Excluding the active-site pocket, strain variation occurs over the entire surface of the neuraminidase heads. The active site was found to be in

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Figure 6 Stereo image of a ball-and-stick model of sialic acid bound to the active site of Tern/N9 neuraminidase. The hydrogen-bond interactions with conserved residues are shown as dotted lines. Nitrogen atoms are shaded black, oxygen atoms are shaded dark gray, and carbon atom are shaded light gray.

a pocket of totally conserved (over all animal subtypes) residues [65]. In this way the enzyme active-site pocket is surrounded by highly variable surface residues that prevent immune recognition of the active site by antibody molecules [25]. The x-ray diffraction studies of neuraminidase—antibody complexes have shown that the footprint of an antibody in the complex is larger than the exposed surface of the conserved region of the active site [67–69]. These structural results indicate that antibodies are unable to exert mutational pressure on the conserved active site because they cannot bind there without engaging strain-variable residues as part of the binding surface. However, as antibodies bind to strain-variable elements of the structure, the virus can overcome host immune pressure by point mutations of the residues that do not have a catalytic or structural role [70,48] but are able to disrupt the antigen—antibody binding interface. The rapid emergence of these escape mutants explains the failure in producing a universal vaccine for influenza. E. The Enzyme Active Site The structures of N-acetyl neuraminic acid (sialic acid Neu5Ac) and the 2-deoxy-2,3-dehydro-N-acetyl neuraminic acid (Neu5Ac2en) inhibitor complexed with N2 neuraminidase [66] revealed the nature of the interactions of the

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molecules in the active-site pocket (Figure 6). Sialic acid binds in the active site in the α-anomer and, in a distorted half-chair conformation, through the same face as used in its interaction with hemagglutinin [71]. The carboxylate group of the sugar interacts with three guanidinium groups of argine residues 118, 292, and 371 and has an equatorial conformation with respect to the sugar ring (this group, axial toward the floor in the undistorted structure, is probably held equatorial by interactions with these arginine residues). The NH group of the 5-N-acetyl side chain interacts with the floor of the active-site cavity via a bound water molecule. The oxygen of the 5-N-acetyl side chain is hydrogen bonded to the Nε of Arg 152, while the methyl group lies in a hydrophobic pocket near Ile222 and Trp178. The last two hydroxyl groups of the 6-glycerol side chain are hydrogen bonded to carboxylate oxygens of Glu276 and the 4hydroxyl is directed to a carboxylate oxygen of Glu119. The glycosidic oxygen O2 interacts with a carboxylate oxygen of Asp151. Similar binding of sialic acid in the active site was observed in type B virus [50]. Comparison of active sites of N2, N9, and type B neuraminidase [72] show there are no significant differences between active-site orientations, except for some minor displacements of Arg224 and Glu276, where the major interactions with the 6-glycerol group of sialic acid occur. However there are differences in the water structure in the active sites of the different subtypes. The Gly405 residue (in N2 and N9) is replaced by a tryptophan in type B, which displaces four water molecules that lie in a solvent pocket bounded by arginines at residues 371 and 118. The Val240 residue (in N2 and N9) is replaced by a methionine in type B, which displaces two water molecules that form a channel under Arg 224, decreasing the flexibility of the active site of type B in this region. These waters are not displaced in the sialic acid/neuraminidase complex in N9 and N2 and would alter the hydrogen-bonding pattern of the complexes when compared to type B. Comparison of the active sites of influenza neuraminidases and bacterial sialidases [51,52] indicates that there is considerable conservation of the catalytic site at the carboxylate-binding end. The residues Asp151, Arg118, Glu277, Arg292, Val or Ile349, Arg371, Tyr406, and Glu425 are conserved over all known viral and bacterial strains. The arginyl residues 118, 292, and 371 position the 2-carboxylate group and the Val(or Ile)349, Glu425, and Glu277 are important in positioning the triarginyl cluster. The residues Asp151 and Tyr406 are presumably important in bond cleavage, but the precise mechanism is still unclear. These eight residues (Figure 7) are thus most likely to be conserved in all neuraminidases. Differences between viral, bacterial, and mammalian neuraminidase structures may correspond with the different role these enzymes have in vivo. These differences are likely to be in the interactions of the 6glycerol, 5-N-acetyl, and 4-OH groups of silaic acid. In the influenza virus, the turnover rate must be balanced against the requirement to maintain

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Figure 7 Stereo image of the active site of Tern/N9 showing the active-site residues surrounding 4-guanidino-Neu5Ac2en (black). Those residues that are conserved in both influenza and bacterial neuraminidase are shaded gray, and those that are conserved only in influenza virus neuraminidase are not shaded.

sufficient sialic acid at the cell surface to enable attachment via the hemagglutinin. This balance may require some configuration of residues in the active site not directly responsible for catalysis but only involved in the binding and release of sialic acid. IV. Neuraminidase Inhibitor Design Earlier screening programs [73] failed to identify potent inhibitors of viral neuraminidase. The first inhibitor synthesized [74] with a Ki value in the micromolar range was Neu5Ac2en. This was based on the proposed transition state of the reaction, where the anomeric carbon (C2) bound to the ketosidic oxygen has a trigonal state. Several analogues of Neu5Ac2en were synthesized soon after, and the most potent of these, a trifluoracetyl derivative, had a Ki of only 0.8 µM [75]. While this compound showed that in cell culture it retarded virus shedding [34,76], it failed as an effective antiviral agent in animals [77]. The x-ray structure of Neu5Ac2en/neuraminidase complexes have been determined for N2 [66], type B [78], and N9 [79]. The Neu5Ac2en molecule binds in the active site of neuraminidase with the carboxylate oxygen atoms placed in the same location as the carboxylate of sialic acid. The 5-N-acetyl, 4-

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hydroxy, and 6 Neu5Ac2en -glycerol are positioned isosterically in the two molecules. An alternative approach to develop sialidase inhibitors has been made using synthetic thioglycoside analogs of gangliosides such as Neu5Aca(2-S-6)Glcb(1-1)Ceramide [80] were shown to inhibit different subtypes of human and animal influenza virus with Ki values of up to 2.8 µM. These metabolically stable ganglioside analogs contain a thioglycosidic linkage to the terminal neuraminic acid that resists cleavage by the enzyme. Several flavonoid neuraminidase inhibitors have been isolated from plant extracts [81], one of which 5,7,4'-trihydroxy-8-methoxyflavone, was a more potent inhibitor than Neu5Ac2en. Recently, in vivo anti-influenza virus activity of a Kampo (Japanese herbal medicine) preparation has shown promising results in inhibiting influenza virus replication in mice [82], but the mode of action of these compounds is unclear. A. Enzyme Mechanism The similar positioning of the carboxylate oxygens and ring of Neu5Ac2en and sialic acid suggests that Neu5Ac2en is probably a transition state analog. As Neu5Ac2en has a higher affinity for neuraminidase than sialic acid, a mechanism of catalysis was proposed [83] that involves the distortion of the substrate by the formation of a oxycarbonium ion intermediate, which has a similar structure to Neu5Ac2en. However the structural results from sialic acid/neuraminidase complexes suggest that the tighter binding of Neu5Ac2en more likely comes from the relaxation of the conformational strain arising from the transition from chair to boat of the pyranose ring of sialic acid in the active site [66]. Evidence for a sialyl cation transition state by isotopic effects [84] support the existence of a oxycarbonium ion intermediate. However the structural basis for neuraminidase activity is still unclear. It has been suggested [66] that the tyrosyl oxygen of Tyr406, assisted by the sialic acid carboxylate itself, could stabilize the developing charge on the oxycarbonium ion intermediate. The reaction would be completed by the activation of a water molecule by a deprotonated Asp151 and its attack on the carbonium, resulting in the formation of the α-anomer of sialic acid [85]. However the pH-activity profile of neuraminidase [86,87,78] suggests a bell-shaped profile, which indicates normal activity from a pH range of about 4.5 to 9. This would indicate that the role of Asp151 as the acid group in the catalysis is unclear. A nonspecific proton donor has been proposed [78], probably a water molecule as the acid group, with a deprotonated Tyr406 stabilizing the oxycarbonium ion, and a proton transferred from water to the departing aglycon group. It has also been postulated that a proton is eliminated at C3 leading to the transformation

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of the oxycarbonium ion into Neu5Ac2en, which is produced irreversibly at low levels from sialic acid by the enzyme [78]. An SN1-type mechanism has been suggested [88] that is facilitated by an activated water molecule, which can be expelled upon inhibitor binding. The catalytic mechanism could possibly proceed without an acid group: the electrostatic potential of the enzyme could lower the barrier preventing the breaking of the ketosidic bond and the solvent could protonate the aglycon after release [89]. Clearly details of the enzyme mechanism have yet to be elucidated definitively. Structural considerations indicate that only Tyr406 (and possibly Glu277) and the triarginyl cluster are essential in the enzyme mechanism and that Asp151 is implicated. B. Inhibitor Design Principles All the nearest-neighbor interactions between sialic acid or Neu5Ac2en and the protein are with totally conserved amino acids. Thus an inhibitor designed to bind only to the conserved active-site residues of neuraminidase would inhibit neuraminidase activity across all strains of influenza. This would enable the development of an antiviral drug that would affect the spread of viral replication potentially in three ways, i.e., transport through the protective mucosal layer, desialyation of freshly synthesized viral glycoproteins, and elution of progeny virions from infected cells. The development of potent inhibitors was based on the structural information of the N2 neuraminidase conserved active site and its complex with sialic acid and Neu5Ac2en [66]. There are no reports of de novo molecules designed to fit into the cavity, and the most useful approach was to consider molecules that were structurally related to Neu5Ac2en. This involved the design of molecules that would bind isosterically to Neu5Ac2en but which were modified to increase the number of favorable interaction with the protein. The method of Goodford [90] enabled the calculation of favorable binding sites for a variety of chemical probes. The validity of this method was indicated by its ability to identify the positions of the carboxylate binding site of sialic acid as an energy minima for a carboxylate probe (Figure 8a), and the successful prediction of known bound-water sites in the active sites by a water probe. Utilizing this methodology, predictions of energetically favorable substitutions to Neu5Ac2en were examined [91]. A replacement of the hydroxyl at the 4-position of the pyranose ring of Neu5Ac2en by an amino group was identified by this procedure as an energetically favorable substitution. A protonated primary amine probe identified a favorable binding site of -16 kcal mol-1 at this location and in a pocket in the active site near two conserved glutamate residues Glu119 and Glu227 (Figure 8b). This suggested that the substitution of the 4-hydroxyl group by an amino group would increase the overall binding interactions by forming a salt link with Glu119. Furthermore the substitution of the 4hydroxyl group with a much bulkier guanidinyl group would lead to even tighter binding as a result

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Figure 8 Stereo image of the residues in the active site of Tern/N9 complexed with 4-guanidino—Neu5Ac2en overlayed with GRID maps [90] (caged mesh contours) for (a) a carboxylate oxygen probe, contoured at -12 kcal/mol; (b) an amino nitrogen probe, contoured at -12 kcal/mol.

of lateral interactions of the terminal nitrogens of the guanidinyl group and the carboxylate groups of Glu119 and Glu227.

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This led to the design and syntheses of 4-amino-Neu5Ac2en and 4-guanidino-Neu4Ac2en [91] which bound to A/Tokyo/3/67 with a Ki of 50 nM and 0.2

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nM, respectively. These compounds were later shown by x-ray studies of 4-guanidino and 4-aminoNeu4Ac2en complexed with A/Tokyo/3/67 neuraminidase to bind close to that predicted by the design studies. However details of the interactions of the guanidinyl group of the 4-guanidino-Neu4Ac2en with the glutamic acid groups (Glu119 and Glu277) in the floor of the active site were slightly different. This was confirmed on a higher resolution x-ray study (Figure 9) of a 4-guanidino-Neu4Ac2en complexed with Tern N9 neuraminidase [72]. One of the primary guanidinyl nitrogens of 4-guanidino-Neu4Ac2en is hydrogen bonded to the main-chain oxygen at residue 178, a carboxylate oxygen of Glu227, and a water molecule. The other primary guanidinyl nitrogen interacts with the main-chain oxygen of residues 178 and 151. The secondary guanidinyl nitrogen interacts with the carboxylate of Glu119 and Asp151. The interactions with Glu119 are electrostatic and van der Waals in character and lack hydrogenbonding geometry (postulated in the design study) as the carboxylate group of Glu119 stacks parallel to the guanidinyl group. Furthermore, theoretical energy-minimized structures of the complex using AMBER [92] converged to the x-ray structure only if the protein nonhydrogen atoms were kept rigid in the x-ray structure [72]. Otherwise this resulted in active site residues showing large distortions in their conformation. This is an example of the difficulty in correctly modeling even modest changes in the interactions of an inhibitor/active site complex. The 4-guanidino analog shows potent inhibition of neuraminidase activity in all known wild strains of influenza. Furthermore it is very specific to in-

Figure 9 Stereo image of the Tern/N9 4-guanidino-Neu5Ac2en complex showing the hydrogen-bond interactions (dotted lines) of the inhibitor with conserved residues in the active site of the enzyme. Nitrogen, oxygen, and carbon atoms are shaded black, dark gray, and light gray, respectively.

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fluenza, as it shows weak inhibition to bacterial, para influenza, and mammalian neuraminidases [91,93]. This is possibly due to the specific interactions of the 4-guanidino group within the subpocket of the active site of influenza neuraminidase that is not conserved in other neuraminidases. In bacterial neuraminidases [51] this pocket is much smaller, and would prevent the binding of the 4-guanidino group in this region of the active site. This is consistent with the proposition that the interactions of sialic acid with the active site of neuraminidases are function specific at the C4, C5, and C6 position of sialic acid and that modification at these positions confer specificity to the target enzyme [72]. Other approaches [94] to structure-based design of inhibitors have to date produced only millimolar inhibition of neuraminidase activity. V. Antiviral Activity In vitro inhibition of viral replication in tissue culture was demonstrated earlier [34] for the trifluro derivative of Neu5Ac2en, but its antiviral activity in vivo was not demonstrated [77]. As a consequence of this, efforts were directed towards hemagglutinin, which was then considered a better target for antiinfluenza drugs. The interest in neuraminidase inhibitors as anti-influenza drugs has only been revived with the success of 4-guanidino-Neu5Ac2en and its analogs in attenuating viral titer in mice when administered directly into the lungs [91,95]. A. Inhibition In Vitro Von Itzstein and co-workers [91] have shown that the 4-amino-and 4-guanidino-Neu5Ac2en inhibit influenza strains A/Singapore/1/57 and B/Victoria/102/95 in MDCK cells with IC50 values (the concentration required to inhibit plaque formation in MDCK cells by 50%) of 1.5 mM and 0.065 mM (4amino) and 0.014 mM and 0.005 mM (4-guanidino) respectively. These IC50 values, in particular for the 4-guanidino compound, are well below those found for amantadine, ribovarin, and Neu5Ac2en. Furthermore in comparison to Neu5Ac2en, the 4-guanidino-Neu5Ac2en inhibitor was 100-fold less active against human lysosomal sialidase and over 1000-fold more active against a wide range of clinical isolates of influenza A and B, including amantidine and rimantadine resistant variants [93]. The inhibition of virus replication in MDCK cells has been confirmed [96] and this knowledge prompted the extension of the inhibition studies to human respiratory epithelium cells in vitro [97], which indicated high antiviral activity for strains of A(HINI) and A(N3N2) isolates. They also found that delayed administration of the drug—after viral replication was well estab-

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lished—was associated with inhibition of virus replication. However viral titer was higher (1.3 log10 compared to 4.0 log10 at 10 mg/mL concentration of the drug) for the delayed administration compared with viral titer when the drug was present throughout the period of viral exposure. The clinical significance of this in the treatment of established infections has yet to be explored in detail, although preliminary clinical trails [98] have indicated some positive results. B. Administration and Inhibition In Vivo The earlier work of Palase and Schulman [77] indicated that the failure in inhibiting viral replication in mice after intranasal and subcutaneous treatment with Neu5aC2en would also occur for Neu5Ac2en analogs. This failure of Neu5Ac2en as an antiviral treatment in animal models can now be ascribed to the rapid excretion of the compound [99] thereby not delivering sufficient concentration of the inhibitor to the infected tissue. It had been shown [91] that the antiviral activity in mice is considerably more, when the 4-guanidino compound is administered intranasally than when the drug is injected intraperitoneally. This can be attributed to the localization of sufficiently high concentration of inhibitor in the lining of the nasal and respiratory epithelia where influenza virus replication is believed to occur. Animal trials with ferrets challenged with influenza virus have shown the 4-guanidino Neu5Ac2en compound is effective in studies [91] involving prophylactic administration of the drug. When the drug is administrated intranasally, 50 µg/kg, twice daily, one day before infection with the virus and the succeeding six days, it substantially reduces virus titer in nasal washing and abolishes fever that usually appears 3 days after infection. The drug is currently undergoing clinical trials, and the initial results from a double blind, randomized, placebo-controlled trial using this compound have been positive both for early treatment and prophylaxis of experimental inoculation of human volunteers [98] with influenza A/Texas/91. C. Drug Resistance Although the active site of influenza virus has been conserved in all known field strains of the virus, the possibility of drug resistance needs to be addressed. Experience with influenza and other viruses, in particular HIV, have shown [100] that drug-resistant mutants arise very rapidly, resulting in the effectiveness of antiviral drugs being short lived. One attempted solution to the problem is the use of several drugs during therapy [101], making it more difficult for the virus the develop resistance. In the case of influenza virus, there have to date been no reports of drug resistance from field strains. However it has recently been reported [102,103]

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that 4-guanidino-Neu5Ac2en-resistant mutants can arise from multiple serial passages of virus in MDCK cells in the presence of the inhibitor. It has been demonstrated [104] that almost all of the mutations arise in the hemagglutinin receptor binding site and not on the neuraminidase. These altered HA variants, which have weaker binding to HA receptors, appear to arise as a result of increased inhibition of the neuraminidase by the drug. This is consistent with the earlier proposition that the rate of desialylation of receptor is critically related to the rate of attachment to receptor, for the virus infection and elution. The decreased activity of the neuraminidase by the drug selects HA mutants with decreased binding to receptors. However a drug-resistant neuraminidase mutation has been isolated [103,105] that results in a single active-site residue mutation—with apparently unaltered activity—of glutamic acid 119 to glycine. The crystal structure of this mutant and its complex (Figure 10) with 4-guanidino-Neu5Ac2en has been determined [105] and the structure suggests that the decrease in inhibitor binding arises from the loss of stabilizing interaction with the 4-guanidino group of the drug [72] and alterations in the solvent structure of the active site. This alteration arises from a water molecule that binds near the location of one of the carboxylate oxygens of the glutamic acid in the wild type molecule. The location of the 4-guanidinoNeu5Ac2en drug in the complex with the mutant enzyme is isosteric compared to the drug/wildtype complex [72]. The only differences are the interactions with residue 119.

Figure 10 Stereo image of the Glu119Gly Tern/N9 mutant complexed with 4-guanidino-Neu5Ac2en. The inhibitor binds in a similar conformation as with the wild type enzyme. The atom labeled “Wat” represents the water molecule found in the mutant enzyme that assumes the role of the glutamic acid in the wild type enzyme. Nitrogen, oxygen, and carbon atoms are shaded black, dark gray and light gray, respectively.

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VI. Conclusion It has now been over a decade since the structure of influenza virus neuraminidase was determined [45,65]. The development of the 4-substituted Neu5ac2en analog inhibitors dates from 1987 when the structure of sialic acid and Neu5Ac2en complexed with neuraminidase were determined to sufficient accuracy to permit modeling of potential inhibitors [66]. The development was a multidisciplinary collaboration of biochemists, crystallographers, molecular modelers, and synthetic chemists and culminated in the synthesis [106] and biological testing of the compounds [91]. Preliminary data on the efficacy of these drugs on humans indicate its effectiveness in both prophylaxis and treatment of the disease [98]. The 4-guanidino compound is in Phase 2 trials (October, 1995). While drug resistance of the virus has been observed in vitro, it will be interesting to see if variants arise in animal studies as they do for Amantidine. This is one of the first rationally designed antiviral drugs to be synthesized and portends well for this methodology to be used as an additional weapon in controling the many pathogens that have plagued humanity for so long. Acknowledgments The author would like to acknowledge Peter Colman, my collaborator in neuraminidase crystallography, Mike Lawrence for the GRID maps shown here and reading this manuscript, Brian Smith for discussions on enzyme mechanisms, Jenny McKimm-Breschkin for discussions on drug resistance, Bert van Donkelaar for technical support, and Paul Davis for computing support. References 1. Kilbourne ED. Influenza. New York: Plenum, 1987. 2. Webster RG, Schafer JR, Suss J, Bean Jr WJ, Kawaoko Y. Evolution and ecology of influenza viruses. In: Hannoun C, et al. eds. Options for the Control of Influenza Virus II. Amsterdam: Excerpta Medica, 1993:177–185. 3. Lamb RA. Genes and proteins of the influenza virus. In: Krug RM, ed. The Influenza Viruses, ed. New York: Plenum, 1989: 1–87. 4. Murphy JS, Bang FB. Observations with the electron microscope on cells of chick chorio-allantoic membrane infected with influenza virus. J Exp Med 1952; 95:259. 5. Compans RW, Dimmock NJ. An electron microscopic study of single-cycle infection of chick embryo fibroblasts by influenza virus. Virology 1969; 39:499. 6. Murphy BR, Webster RG. Orthomyxovirus. In: Fields BN, Knipe DM, eds. Virology. New York: Raven Press, 1990:1091–1152.

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7. Ennis FA, Meager A. Immune interferon produced to high levels antigenic stimulation of human lymphocytes with influenza virus. J Exp Med 1981; 154:1279–1289. 8. Sprenger MJW, Beyer WEP, Kempen BM, Mulder PGH, Masurel N. Risk factors for influenza mortality. In: Hannoun C, et al. eds. Options for the Control of Influenza Virus II. Amsterdam: Excerpta Medica, 1993:15–23. 9. Smith FI, Palase P. Variation in influenza virus genes. Epidemiological, pathogenic, and evolutionary consequences. In: Krug RM, ed. The Influenza Viruses. New York: Plenum, 1989:319–359. 10. Skehel JJ. The origin of pandemic influenza virus. Symp Soc Gen Microbiol 1974; 24:321–342. 11. Webster RG, Laver WG. Antigenic variation in influenza viruses. In: Kilbourne ED, ed. Influenza Virus and Influenza. New York: Academic, 1975:269–314. 12. Smith W, Andrewes CH, Laidlaw PP. A virus obtained form influenza patients. Lancet 1933; 2:66–68. 13. Beveridge WI. Influenza: the last great plague. London: Heinemann, 1977. 14. Daniels RS, Downie JC, Hay AJ, Knossow M, Skehel JJ, Wang ML, Wiley DC. Fusion mutants of the influenza virus haemagglutinin glycoprotein. Cell 1985; 40:431–439. 15. Hay AJ, Thompson CA, Geraghty A, Hayhurst S, Grambas S, Bennett MS. The role of the M2 protein in influenza virus infection. In: Hannoun C, et al. eds. Options for the Control of Influenza Virus II. Amsterdam: Excerpta Medica, 1993:281–288. 16. Pinto LH, Holsinger LJ, Lamb RA, Influenza virus M2 protein has ion channel activity. Cell 1992; 69:517–528. 17. Hayden FG. Update on antiviral agents and viral drig resistance. In: Mandell GL, Douglas RG, Bennett JE, eds. Principles and Practice of Infectious Disease. Vol. 2. 2d ed. New York: Churchill Livingston. 1993:3–15. 18. Tyrrell DAJ. Influenza vaccines. Phil Trans R Soc Lond 1980; B288:449–460. 19. Fox JP, Cooney MK, Hall CE, Foy HM. Influenza virus infections in Seattle families, 1975–1979. II. Pattern of infection of invaded households and relation of age and prior antibody to occurrence of infection by time and age. Am J Epidemiol 1982; 116:228–242. 20. Chakraverty et al. Influenza Activity—United States and worldwide, and composition of the 1993–94 influenza vaccine. JAMA 1993; 269:1778–1779.

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21. Both GW, Sleigh MJ, Cox NJ, Kendal AP. Antigenic drift in influenza virus H3 haemagglutinin from 1968 to 1980: multiple evolutionary pathways and sequential amino acid changes at key antigenic sites. J Virol 1983; 48:52–60. 22. Donnelly JJ, Friedman A, Martinez D, Montgomery DL, Shiver JW, Motzel SL, Ulmer JB, Liu MA. Preclinical efficacy of a prototype DNA vaccine: enhanced protection against antigenic drift in influenza virus. Nature Med 1995; 1:583–586. 23. Hirst GK. Adsorption of influenza hemagglutinins and virus by red blood cells. J Exp Med 1942; 76:1740–1743. 24. Colman PM, Influenza virus neuraminidase: Krug RM, ed. Enzyme and antigen. In: The Influenza Viruses. New York: Plenum, 1989:175–218. 25. Colman PM. Influenza virus neuraminidase: structure, antibodies, and inhibitors. Protein Science 1994; 3:1687–1696.

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26. Klenk E, Faillard H, Lempfrid H. Uber die enzymatishe Wirkung von Ifluenza. Z Physiol Chem 1955; 301:235–246. 27. Gottschalk A. Neuraminidase: the specific enzyme of influenza virus and vibrio cholerae. Biochem Biophys Acta 1957; 23:645–646. 28. Rogers GN, Paulson JC. Receptor determinants of human and animal influenza virus isolates: difference in receptor specificity of the H3 haemagglutinin based on species of origin. Viroloy 1983; 127:361–373. 29. Burnett FM. Mucins and mucoids in relation to influenza virus action. IV. Inhibition by purified mucoid of infection and haemagglutinin with the virus strain WSE. Aust J Exp Biol Med Sci 1947; 26:381–387. 30. Gottschalk A. Neuraminidase: its substrate and mode of action. Adv Enzymol 1958; 20:135–145. 31. Liu C, Air GM. Selection and characterization of a neuraminidase-minus mutant of influenza virus and its rescue by cloned neuraminidase genes. Virol 1993; 194:403–407. 32. Liu C, Eichelberger MC, Compans RW, Air GM. Influenza type A virus neuraminidase does not play a role in viral entry, replication, assembly, or budding. J Virol 1995; 69:1099–1106. 33. Palase P, Tobita K, Ueda M, Compans RW. Characterization of temperature sensitive influenza virus mutants defective in neuraminidase. Virology 1974; 61:397–410. 34. Palase P, Compans RW. Inhibition of influenza virus replication in tissue culture 2-deoxy-2,3dehydro-N-trifluroacetylneuraminic acid (FANA): mechanism of action. J Gen Virol 1976; 33:159–163. 35. Griffin JA, Compans RW. Effect of cytochalsin B on the maturation of enveloped viruses. J Exp Med 1979; 150:379–391. 36. Bucher DJ, Palase P. The biologically active proteins of influenza virus neuraminidase. In: Kilbourne ED, ed. Influenza virus and influenza. New York: Academic, 1975:83–123. 37. White DO. Influenza viral proteins: identification and synthesis. Curr Top Microbiol Immunol 1974; 63:1–48. 38. Laver WG. The structure of influenza virus 3. Disruption of the virus particle and separation of neuraminidase activity. Virology 1963; 20:251–262. 39. Laver WG, Valentine RC. Morphology of the isolated hemagglutinin and neuraminidase subunits of influenza virus. Virology 1969; 38:105–119.

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40. Drzenick R, Frank H, Rott R. Electron microscopy of purified influenza virus neuraminidase. Virology 1968; 36:703–707. 41. Laver WG. Crystallization and peptide maps of neuraminidase “heads” from H2N2 and H3N2 influenza virus strains. Virology 1978; 86:78–87. 42. Blok J, Air GM, Laver WG, Ward CW, Lilley GG, Woods EF, Roxburgh CM, Inglis AS. Studies on the size, chemical composition, and partial sequence of the neuraminidase (NA) from type A influenza virus show that the Nterminal region of the NA is not processed and serves to anchor the NA in the viral membrane. Virology 1982; 119:109–121. 43. Colman PM, Laver WG. The structure of influenza virus neuraminidase at 5A resolution. In: Structural aspects of recognition and assembly in biological macromolecules. I.S.S. Rehovot, 1981:869–872.

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44. Wright CE, Laver WG. Preliminary crystallographic data for influenza virus neuraminidase “heads”. J Mol Biol 1978; 120:133–136.

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45. Varghese JN, Laver WG, Colman PM. Structure of the influenza virus glycoprotein antigen neuraminidase at 2.9Å resolution. Nature 1983; 303:35–40. 46. Varghese JN, Colman PM. Three-dimensional structure of the neuraminidase of influenza virus A/Tokyo/3/67 at 2.2Å resolution. J Mol Biol 1991; 221:473–486. 47. Baker AT, Varghese JN, Laver WG, Air GM, Colman PM. The three-dimensional structure of neuraminidase of subtype N9 from an avian influenza virus. Proteins 1987; 1:111–117. 48. Tulip WG, Varghese JN, Baker AT, van Donkelaar A, Laver WG, Webster RG, Colman PM. Refined atomic structures of N9 subtype influenza virus neuraminidase and escape mutants. J Mol Biol 1992; 221:487–497. 49. Tulip WG, Varghese JN, Laver WG, Webster RG, Colman PM. Refined crystal structure of the influenza virus N9 neuraminidase-NC41 Fab complex. J Mol Biol 1992; 227:122–148. 50. Burmeister WP, Ruigrok RWH, Cusack S. The 2.2Å resolution crystal structure of influenza B neuraminidase and its complex to sialic acid. EMBO J 1991; 11:49–56. 51. Crennell SJ, Garman EF, Laver WG, Vimr ER, Taylor GL. Crystal structure of a bacterial sialidase (from Salmonella typhimurium LT2) shows the same fold as an influenza virus neuraminidase. Proc Natl Acad Sci USA 1993; 90:9852–9856. 52. Crennell SJ, Garman E, Laver G, Vimr E, Taylor GL. Crystal structure of Vibro Cholerae neuraminidase reveals dual lectin-like domains in addition to the catalytic domain. Structure 1994; 2:535–544. 53. Vellieux FMD, Huitema F, Groendijk HN, Kalk KH, Frank J, Jonjejan JA, Duine JA, Petratos K, Drenth J, Hol WGJ. Structure of quinoprotein methylamine dehydrogenase at 2.25A resolution. EMBO J 1989; 8:2171–2178. 54. Ito N, Phillips SEV, Stevens C, Ogel ZB, McPherson MJ, Keen JN, Yadav KDS, Knowles PF. Novel thioether bond revealed by a 1.7A crystal structure of galactose oxidase. Nature 1991; 350:87–90. 55. Xia Z-X, Dia W-W, Xion J-P, Hao Z-P, Davidson VL, White S, Mathews FS. The three dimensional structure of methonol dehydrogenase from two methylotrophic bacteria at 2.6A resolution. J Biol Chem 1992; 267:22289. 56. Richardson JS. The anatomy and taxonomy of protein structure. Advan Protein Chem 1981; 34:167–339. 57. Wang J, Yan Y, Garrett TPJ, Liu J, Rogers DW, Garlick, Tarr GE, Husain Y, Reinherz EL, Harrison SC. Atomic structure of a fragment of human CD4 containing two immunogloblin-like domains. Nature 1990; 348:411–418. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_483.html (1 of 2) [4/9/2004 12:28:58 AM]

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58. Baker NJ, Gandhi SS. Effect of Ca++ on the stability of influenza virus neuraminidase. Arch Virol 1976; 52:7–18. 59. Burmeister WP, Cusack S, Ruigrok RWH. Calcium is needed for the thermostability of influenza B virus neuraminidase. J Gen Virol 1994; 75:381–388.87. 60. Wagh PV, Bahl OP. Sugar residues on proteins. Crit Rev Biochem 1981; 307–377. 61. Hitte AL, Nayak DP. Complete nucleotide sequences of the neuraminidase gene of human influenza virus A/WSN/33. J Virol 1982; 41:730–734. 62. Li S, Schulman J, Itamura S, Palase P. Glycosylation of neuraminidase determines the neurovirulence of influenza A/WSN/33 virus. J Virol 1993; 67:6667–6673. 63. Ward CW, Murry JM, Roxburgh CM, Jackson DC. Chemical and antigenic characterisation of the carbohydrate side chains of an Asian (N2) influenza virus neuraminidase. Virology 1983; 126:370–375.

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64. Ward CW, Elleman TC, Azad AA. Amino acid sequence of the pronase-released heads of neuraminidase subtype N2 from the Asian strain A/Tokyo/3/67 of influenza virus. Biochem J 1982; 207:91–95. 65. Colman PM, Varghese JN, Laver WG. Structure of the catalytic and antigenic sites in influenza virus neuraminidase. Nature 1983; 303:41–44. 66. Varghese JN, McKimm-Breschkin JL, Caldwell JB, Kortt AA, Colman PM. The structure of the complex between influenza virus neuraminidase and sialic acid, the viral receptor. Proteins 1992; 14:327–332. 67. Colman PM, Laver WG, Varghese JN, Baker AT, Tullock PA, Air GM, Webster RG. Threedimensional structure of a complex of antibody with influenza virus neuraminidase. Nature 1987; 326:358–363. 68. Colman PM, Tulip WR, Varghese JN, Tulloch PA, Baker AT, Laver WG, Air GM. 3-D structures of influenza virus neuraminidase-antibody complexes. Proc Roy Soc Lond 1989; B,323:511–518. 69. Malby RL, Tulip WR, Harley VR, McKimm-Breschkin JL, Laver WG, Webster RG, Colman PM. The structure of a complex between the NC10 antibody and influenza virus neuraminidase and comparison with the overlapping binding site of the NC41 antibody. Structure 1994; 2:733–746. 70. Varghese JN, Webster RG, Laver WG, Colman PM. The three-dimentional structure of an escape mutant of the neuraminidase of influenza virus A/Tokyo/3/67. J Mol Biol 1988; 200:201–203. 71. Weis W, Brown JH, Cusack S, Paulson JC, Skehel JJ, Wiley DC. Structure of the influenza virus haemagglutinin complexed with its receptor, sialic acid. Nature 1988; 333:426–431. 72. Varghese JN, Epa VC, Colman PM. Three-dimensional structure of 4-guanidino-Neu5Ac2en and influenza virus neuraminidase. Protein Science 1995; 4:1081–1087. 73. Edmond JD, Johnston RG, Kidd D, Rylance HJ, Sommerville RG. The inhibition of neuraminidase and antiviral action. Br J Pharmacol Chemother 1966; 27:415–426. 74. Meindl P, Tuppy H. 2-Deoxy-2,3-dehydrosialic acids. I. Synthesis and properties of 2-deoxy-2,3dehydro-N-acetylneuraminic acids and their methy esters. Monatsh Chem 1969; 100:1295–1306. 75. Meindl P, Bodo G, Palase P, Schulman J, Tuppy H. Inhibition of neuraminidase activity by derivatives of 2-deoxy-2,3-dehydro-N-acetylneuraminic. Virology 1974; 58:457–463. 76. Palase P, Schulman JL, Bodo G, Meidl P. Inhibition of influenza and parainfluenza virus replication in tissue culture by 2-deoxy-2,3-dehydro-N-trifluroacetylneuraminic acid (FANA). Virology 1974; 59:490–498.

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77. Palase P, Schulman JL. Inhibitors of viral neuraminidase as potential antiviral drugs. In: Chemoprophylaxis and virus infections of the upper respiratory tract. vol. 1. Boca Raton, Florida: CRC Press, 1977:189–205. 78. Burmeister WP, Henrissat B, Bosso C, Cusack S, Ruigrok RWH. Influenza B virus neuraminidase can synthesize its own inhibitor. Structure 1993; 1:19–26. 79. Bossart-Whitaker P, Carson M, Babu YS, Smith CD, Laver WG, Air GM. Three dimensional structure of influenza A N9 neuraminidase and its complex with the inhibitor 2-deoxy 2,3-dehydro-Nacetyl neuraminic acid. J Mol Biol 1993; 232:1069–1083.

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80. Suzuki Y, Sato K, Kiso M, Hasegawa A. New ganglioside analogs that inhibit influenza virus sialidase. Glycoconjugate J 1990; 7:349–356. 81. Nagai T, Miyaichi Y, Tomimori T, Suzuki Y, Yamada H. Inhibition of influenza virus sialidase and anti-influenza virus activity by plant flavonoids. Chem Pharm Bull 1990; 38(5):1329–1332. 82. Nagai T, Yamada H. In vivo anti-influenza virus activity of Kampo (Japanese herbal) medicine “Shoseiryu-to” and its mode of action. Int J Immunopharmacol 1994; 16(8):605–613. 83. Miller CA, Wang P, Flashner M. Mechanism of Arthro-bacter sialophilus neuraminidase: the binding of substrates and transition state anologs. Biochem Biophys Res Commun 1978; 83:1479–1487. 84. Chong AKJ, Pegg MS, Taylor NR, von Itzstein M. Evidence for a sialyl cation transition state complex in the reaction of sialidase from influenza. Eur J Biochem 1992; 207:335–343. 85. Friebolin H, Brossmer R, Keilich G, Ziegler D, Supp M. 1H-NMR spektroskopischer nachweis der N-acetyl-a-D-neuraminsaure als primares spaltprodukt der neuraminidasen. Hoppe Seyler's Z Physiol Chem 1980; 361:697–702. 86. Lentz MR, Webster RG, Air GM. Site-directed mutation of the active site of influenza neuraminidase and implications for the catalytic mechanism. Biochemistry 1987; 26:5351–5358. 87. Chong AKJ, Pegg MS, von Itzstein M. Characterization of an ionisable group involved in the binding and catalysis by sialidase from influenza virus. Biochem Int 1991; 24:165–171. 88. Taylor NR, von Itzstein M. Molecular modeling studies on ligand binding to sialidase from influenza virus and the mechanism of catalysis. J Med Chem 1994; 37:616–624. 89. Janakiraman MN, White CL, Laver WG, Air MA, Luo M. Structure of influenza virus neuraminidase B/Lee/40 complexed with sialic acid and a dehydro anolog at 1.8A-resolution: implications for the catalytic mechanism. Biochem 1994; 33:8172–8179. 90. Goodford PJ. A computational procedure for determining energetically favorable binding sites on biologically important macromolecules. J Med Chem 1985; 28:849–857. 91. Von Itzstein M, Wu W-Y, Kok GB, Pegg MS, Dyson JC, Jin B, VanPhan T, Symthe ML, White HF, Oliver SW, Colman PM, Varghese JN, Ryan DM, Woods JM, Bethell R C, Hotham VJ, Cameron JM, Penn CR. Rational design of potent sialidase-based inhibitors of influenza virus replication. Nature 1993; 363:418–423. 92. Pearlman DA, Case DA, Caldwell J, Seibel G, Singh UC, Weiner PK, Kollman PA. AMBER 4.0. San Francisco, California: University of California, 1990.

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93. Woods JM, Bethell RC, Coates JAV, Healy N, Hiscox SA, Pearson BA, Ryan DM, Ticehurst J, Tilling J, Walcott SM, Penn CR. 4-Guanidino-2-4-Dideoxy-2,3-Dehydro-N-Acetylneuraminic Acid is a highly effective inhibitor both of the sialidase (Neuraminidase) and growth of a wide range of influenza A and B viruses In Vitro. Antimicro Agents and Chemotherapy 1993; 37:1473–1479. 94. Luo M, Jedrzejas MJ, Singh S, White CL, Brouillette WJ, Air GM, Laver WG. Benzoic acid inhibitors of influenza virus neuraminidase. Acta Cryst 1995; D51:504–510.

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95. Ryan DM, Ticehurst J, Dempsey MH, Penn CR. Inhibition of influenza virus replication in mice by GG167 (40guanidino-2,4-dideoxy-2,3-dehydro-N-acetylneuraminic acid) is consistent with extracellular activity of viral neuraminidase. Antimicrob Chem Chemother 1994; 38(10),2270–2275. 96. Thomas GP, Forsyth M, Penn CR, McCauley JW. Inhibition of growth of influenza virus in vitro by 4-guanidino-2,4-dideoxy-2,3-dehydro-N-acetylneuraminic acid. Antiviral Res 1994; 24:351–356. 97. Hayden FG, Rollins BS, Madren LK. Anti-influenza activity of the neuraminidase inhibitor 4guanidino-Neu5Ac2en in cell culture and in human respiratory epithelium. Antiviral Res 1994; 25:123–131. 98. Hayden FG, Treanor JJ, Betts RF, Lobo M, Esinhart JD, Hussey EK. Safety and efficacy of the neuraminidase inhibitor GG167 in experimental human influenza, JAMA 1996; 275:295–299. 99. Nohle U, Beau J-M, Schauer R. Eur J Biochem 1982; 126:543–548. 100. Kimberlin DW, Crumpacker CS, Straus SE, Biron KK, Drew WL, Hayden FG, McKinlay M, Richman DD, Whitley RJ. Antiviral resistance in clinical practice. Antiviral Res 1995; 26:423–438. 101. Madren LK, Shipman C, Hayden FG. In vitro inhibitory effects of combinations of anti-influenza agents. Antiviral Chem Chemotherapy 1995; 6(2):109–113. 102. McKimm-Breschkin JL, Marshall D, Penn CR. Phenotypic changes observed in influenza viruses passaged in 4-amino or 4-guanidino-Neu5Ac2en in vitro. Abstracts, 9th Internat. Conf. on Negative Strand Viruses. Estoril, Portugal, 1994:260. 103. Gubareva LV, Bethell R, Hart GJ, Penn CR, Webster RG. Characterization of mutants of influenza virus selected with 4-guanidino-Neu5Ac2en. Abstracts, 14th Annual Meeting, Amer. Soc. for Virol. Austin, Texas, 1995:W44-1. 104. McKimm-Breschkin JL, Blick TJ, Sarasrabudhe AV, Tiong T, Marshall D, Hart GJ, Bethell RC, Penn CR. Generation and characterisation of variants of the NWS/G70C influenza virus after in vitro passage in 4-amino-Neu5Ac2en and 4-guanidino-Neu5Ac2en. Antimicrob Agents Chemotherapy 1995; in press. 105. Blick TJ, Tiong T, Sahasrabudhe A, Varghese JN, Colman PM, Hart GJ, Bethell RC, McKimmBreschkin JL. Generation and characterization of an influenza virus variant with decreased sensitivity to the neuraminidase specific inhibitor 4-guanidino-Neu5Ac2en. Virology 1995; 214:475–484. 106. von Itzstein M, Wu W-Y, Jin B. The synthesis 2,3-didehydro-2,4-dideoxy-4-guanidiny-Nacetylneuraminic acid: a potent influenza virus inhibitor. Carbohydr Res 1994; 259:301–305. 107. Kraulis PJ. MOLSCRIPT: a program to produce both detailed and schematic plots of protein structures. J Appl Cryst 1991, D50:869–873. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_486.html (1 of 2) [4/9/2004 12:30:20 AM]

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19 Rhinoviral Capsid-Binding Inhibitors: Structural Basis for Understanding Rhinoviral Biology and for Drug Design Vincent L. Giranda Abbott Laboratories, Abbott Park, Illinois Guy D. Diana ViroPharma, Inc., Malvern, Pennsylvania I. Introduction A cure for the common cold has been sought after so long that it has become a clicheé: “We can _____(do something difficult) but we can't cure the common cold”. There are, unfortunately, a number of factors that conspire to make the cure for the common cold ephemeral. In spite of these difficulties, dramatic progress has been made in producing chemotherapies for human rhinoviruses (HRVs), which are the major cause of the common cold in humans [1]. Although most colds are generally both mild and self-limiting, they are responsible for both a large proportion of visits to physicians and lost work time [2]. Billions of dollars are spent in the United States alone on symptomatic relief from this disease. The ubiquitousness of this disease has led many people to seek a cure for many years (the discovery of the class of HRV inhibitors described here is over 20 years old). This chapter will describe the influence of structure-based approaches in the design of a class of antipicornaviral agents called capsid-binding inhibitors. Any effort to inhibit HRV replication, and thus cure many common colds, is made more difficult by three factors. First, there are at least 102 described serotypes of HRVs, and it seems likely that there are many more serotypes not yet described [3]. Rhinoviruses are responsible for about 40–60% of the colds in humans [1,4]. Therefore, a chemotherapeutic agent would need to be effective

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against most of the HRV serotypes to be useful in approximately 50% of common colds. This percentage may be somewhat higher because some of the other causes of cold-like illnesses, particularly the enteroviruses (e.g., Coxsackie and echoviruses), are also inhibited by capsid-binding compounds [5–7]. In order to be efficacious, drugs must necessarily have a broad spectrum of activity. The second difficulty in producing HRV chemotherapy stems from the relatively innocuous nature of HRV infection. Compounds must be able to be very safely administered, with a minimum of drug-drug interactions, if therapy is to be acceptable. An analogy may be drawn between common headache remedies and common cold chemotherapy. Common headache cures such as nonsteroidal antiinflammatory agents and acetaminophen clearly can cause serious side effects (gastrointestinal bleeding or catastrophic liver failure) particularly if misused [8]. In spite of this possibility, serious complications from these agents are quite rare. One would suspect that an antirhinoviral agent would need to be at least as safe as headache remedies. The third major difficulty in developing cold cures arises from the fact that the HRVs are RNA viruses. When presented with any selective pressure, including chemotherapeutic or antibody challenge, RNA viruses mutate rapidly [9]. This ability to mutate is most clearly illustrated in influenza viruses (RNA viruses), where new strains continuously arise to circumvent immunity in a population. Influenza A viruses have been shown to mutate around the anti-influenza drug Amantadine, after a single passage through a susceptible human host. The mutated viruses shed from a host treated with Amantadine are now resistant to Amantadine. These mutated viruses appear to be as virulent as the parent strain of virus [10]. Any effort at antirhinoviral therapy must attend to these three issues: (1) serotypic diversity; (2) exceptional safety; and (3) viral resistance. Therefore, any structure-based approach cannot concentrate on potency alone, but must also attend to these three issues as well. The requirement for inhibiting multiple targets has been addressed in tangible ways using structure-based design and will be discussed here. Safety issues have been addressed in limited published data from clinical and preclinical studies, but structure-based design has not played any significant role in addressing these problems. One might argue that structure-based approaches have aided in the design of clinical backups. These backups are then brought forward after an initial drug fails for safety reasons. Drug-resistant mutations created in the laboratory have also been examined structurally and will be discussed. The importance of resistance developing in the clinical setting has not yet been answered. This chapter will first introduce the target, the HRVs, and describe their anatomy and life cycle. Emphasis will be placed on the viral capsid and its disassembly or uncoating. How structural information has helped in our under-

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standing viral physiology will be highlighted. This will be followed by a description of capsid-binding antirhinoviral compounds. We will leave discussions of other HRV targets, e.g., proteases, to other authors more knowledgeable in these subjects. Drug structure—activity relationships will be discussed, followed by a discussion of drug resistance. Finally clinical trials and future prospects for capsidbinding inhibitors in any antirhinoviral armamentarium will be discussed. II. The Human Rhinovirus Anatomy The human rhinoviruses are picornaviruses (pico = small; rna = RNA), a family of small (300 Å in diameter), positive sense, single-stranded RNA viruses. Other generas in this family include the enteroviruses (e.g., polioviruses); the aphthoviruses (e.g., foot-and-mouth disease virus); the cardioviruses (e.g., mengovirus, EMC virus); and the heparnaviruses (e.g., hepatitis A virus). With the exception of the heparnaviruses, a crystallographic structure is known for at least one of the viruses in each genera [5,11–17]. These structures show remarkable similarities, which will be described. In spite of these similarities, caution should be exercised when trying to generalize data gathered in one genus to other members of the picornavirus family. The picornaviruses share an icosahedral structure (Figure 1). The icosahedral protein coat that encapsidates the viral RNA is made up of 60 symmetrically arranged protomers. Each of these protomers is comprised of four viral polypeptides, termed VP1 through VP4. This was the extent of our structural knowledge of the picornaviruses until the 1985 structure of HRV14 was published by Michael Rossmann and coworkers [16]. This was followed rapidly by structural determination of a variety of other picornaviruses. These structures provided the framework on which to base current paradigms for the picornaviral life cycles, particularly with regard to their assembly, attachment, and uncoating. The VP1, VP2, and VP3 all contain a core eight-stranded antiparallel β barrel (Figure 2). These polypeptides have surfaces on both the exterior and interior of the virion particle, facing both solvent and viral RNA. The fourth polypeptide, VP4, is considerably smaller than the others and does not contain the eight-stranded barrel motif. The VP4 resides entirely on the interior surface of the virion, in close association with the viral RNA. The N-terminus of VP4 is known to be myristoylated in both rhinoand polioviruses [18,19]. Three regions of the picornavirus structure deserve special attention because they appear to play crucial roles in the viral life cycle as well as bear on the function of the capsid-binding compounds. These regions are the canyon, the VP1 hydrophobic pocket, and the β cylinder.

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Figure 1 Schematic illustration of the icosahedral rhinovirus 14. (a) Shown is the icosahedron comprised of 60 copies each of VP1 (light gray), VP2 (black), and VP3 (gray). The shaded circles around each five-fold axis indicate the canyon positions. Also indicated is the approximate position of the VP1 hydrophobic pocket that lies underneath the surface of the virion. (b) An icosahedral pentamer is expanded with one viral protomer shown as a protein ribbon diagram. (c) This pentamer is seen in a cutaway view. Here VP1 is white, VP2 and VP4 black, and VP3 gray. A capsid-binding compound is depicted as black spheres inside the VP1 ribbon diagram. The cross hatched regions on the (c) schematic (right) indicate areas that disorder when HRV14 crystals are exposed to acid.

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A. The Canyon The HRVs have been divided into the major and minor receptor groups based on two identified cellular receptors [3]. The major group, which is comprised of approximately 90 serotypes, binds to the intercellular adhesion molecule 1 (ICAM-1) [20]. The minor group, about 10 serotypes, binds to the low density lipoprotein receptor family [21]. The canyons are depressions approximately 15 to 20 Å deep that encircle each icosahedral five-fold axis (Figure 1). When first seen in HRV 14, these canyons were postulated to be the site at which a cellular receptor would bind. Subsequent electron-microscopic data revealed that ICAM-1 does indeed bind in the canyon as predicted, although in a somewhat different orientation than early models [22,23]. These canyons allow the receptor binding sites to escape immunological surveillance because the canyons are too narrow to allow an immunoglobulin to contact the canyon floor. Directly underneath the floor of the canyon lies a second important structure, the VP1 hydrophobic pocket. B. The VP1 Hydrophobic Pocket The VP1 hydrophobic pocket is the site where the capsid-binding compounds reside. This hydrophobic pocket in VP1 was not initially apparent in the HRV14 structure because it exists in a closed conformation in the native HRV14. The addition of a capsid-binding antiviral agent induces the pocket to open. This was first seen by Smith and coworkers when the first crystal structure of a capsid-binding drug, WIN 51711, was solved bound in HRV14 [24]. (WIN is the designation for a Sterling Winthrop compound.) This was a seminal event in the structure-based design of these capsid-binding compounds; before this structure was solved the exact site at which the compounds bound was unknown.

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Figure 2 Panels (a), (b), and (c) depict VP1, VP2, and VP3 β barrels, respectively. In all cases the view is such that the virion exterior would be on the right side of the page. A capsid-binding compound is depicted as gray spheres as it is found inside VP1. http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_492.html (1 of 2) [4/9/2004 12:34:08 AM]

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Subsequent studies have shown this pocket is present in every known HRV and enterovirus structure. This pocket is inside the β barrel of VP1, directly underneath the canyon floor where ICAM-1 binds (Figure 1c). The proximity of the hydrophobic drug-binding site to the receptor-binding site explains the effects these compounds have on viral attachment in specific HRV serotypes (see below). C. The β Cylinder The β cylinder is a complicated structure that lies on the inside of the viral coat (Figure 1c). There is one β cylinder at each icosahedral five-fold axis. The β cylinder is formed by winding the five-fold related VP3 N-termini around the symmetry axis. In close association with this cylinder is the N-terminal regions of VP1 and VP4. The myristoyl moiety attached to VP4 is observed in the poliovirus structures. This moiety is in close association with the cylinder formed by VP3 [19]. In HRVs, density consistent with the myristoyl moiety is also seen near the β cylinder [25]. In the HRVs, structural disorder of the first 25–28 Nterminal residues of VP4 obscures the connection between VP4 and the myristoyl group. The addition of myristic acid to protein is seen in many viral and cellular

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proteins. It is typically a signal moiety that directs the protein to which it is attached towards cellular membranes [26]. Above the β cylinder is an ion, thought to be Ca2+, which lies right on the five-fold axis and coordinates to the five adjacent VP1 polypeptides [25,27]. The proximity of the β cylinder to structures shown later to become external during uncoating (portions of VP1, VP4) suggests that it may be important in uncoating. III. The Human Rhinovirus Life Cycle The HRVs must undergo a number of transitions to replicate (Figure 3). First, they must attach to the cell surface at a cellular receptor. They are then internalized into the cell via the endosomal compartment. Following internalization, they must eject their RNA out of the viral capsid, through a lipid bilayer, and into the cytosol in a manner that preserves the integrity of the RNA. Replication

Figure 3 A schematic diagram depicting some of the required steps in viral replication.

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of the RNA and production of a polyprotein ensues. The polyprotein is then processed by viral proteases to form the viral polypeptides. The coat must then assemble, package RNA, and leave the cell. The capsid-binding compounds' effect on propagation have been shown to occur at the attachment, uncoating, and assembly steps (25,27–30). The relative effect on each of these steps appears to be variable and has not yet been extensively characterized. The most universal effect appears to be on the uncoating step. During uncoating several different rhinovirus subparticles are observed. These are thought to be intermediates in the uncoating process [32–33]. The fully infectious 149S particle becomes an 125S Aparticle, which has lost VP4 but still maintains viral RNA. This A-particle is no longer infectious. The Aparticle then releases RNA to become an 80S empty shell. The formation of these types of particles can be induced by association of the virion with the cell receptor or acidification [32,34]. In poliovirus, attachment to cells has been shown to lead to a particle that has lost VP4 and has externalized the N-terminal region of VP1, which normally resides on the virion interior. There has been considerable debate about how the HRVs accomplish the transfer of RNA from inside the virion into the cell cytosol. This step is crucial for productive uncoating. An important question concerns the requirement for acidification of the endosome for HRVs to release their RNA. Evidence that appeared to conflict was found in a number of studies using either entero- or rhinoviruses (35–39). This question was later addressed by experiments that specifically separated entero- and rhinovirus behavior [40]. These experiments showed that HRVs, unlike poliovirus, require a pH-lowering step for productive infection. This pH lowering is likely to occur in the endosomal compartment. It should be noted that HRV and enteroviruses have been classified historically based on their resistance to acid: HRVs are acid-labile, while enteroviruses are stable in acid. Consequently, differences in behavior between the rhino- and enteroviruses in an acidic environment within the cell are not surprising. To replicate the changes in HRV that might be induced by acidification in the endosome, HRV14 was acidified in the crystalline state and examined via x-ray diffraction. When compared to the native HRV14, the acidified HRV14 capsid becomes disordered in three regions: the Ca2+ ion on the five-fold axis; a region of the β cylinder and the adjacent portion of VP4; and the GH loop [41]. The GH loop is the region of structure that connects β-strands G and H and lies directly between the hydrophobic pocket and the receptor binding site (Figure 4). It forms the roof of the VP1 hydrophobic pocket and the floor of the canyon at the receptor binding site. Mutants that are resistant to acid in vitro were isolated [41,42]. These mutants cluster about the GH loop (Table 1, Figure 4), and typically would be thought of as mutants that stabilize protein structure (e.g., larger or more

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Page 496 Table 1 VP1 Acid- and Drug-Resistant Mutations Phenotype

Mutations in VP1

Acid resistant

H1078L, H1078Y, N1100K, N1100T, D1101E, N1145S, W1163R, V1188A, V1191A, T1216I, M1221L, M1224L, A1225V

Drug resistant (compensation)

N1100S, N1105S, V11531, N1219S, S1223G

Drug Resistant (exclusion)

V1188L, V1188M, C1199F, C1199Y, C1199W, C1199R

branched amino acids) [43–46]. The proximity of these mutations to the GH loop coupled with the loop's movement on acidification suggest that the GH-loop movement plays an important role in uncoating. It has also been suggested that binding of ICAM-1 to the virions may also play a role in uncoating. In support of this hypothesis, it has also been observed

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Figure 4 A ribbon diagram of HRV14 VP1 bound to WIN 61605 (small gray spheres with black bonds). The VP1 region that disorders under acid conditions is depicted by black color on the ribbon diagram. Residues that can be mutated to be acid stable, drug resistant (compensation type) or to either phenotype are shown as black, white, and gray ball-and-stick models, respectively. The majority of mutations are near the site of drug binding as well as the site of acid-induced disorder in VP1.

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that in some HRVs (particularly HRV14 and to a lesser extent in HRV3, but not HRV16) the addition of ICAM-1 itself, in the absence of acid, can induce structural changes that mimic uncoating in the capsid [34]. However, these changes are unlikely to lead to productive uncoating because they occur in the extracellular space. Therefore, the binding of ICAM alone, at the cell surface, is unlikely to be sufficient to cause productive uncoating of HRVs. It seems that the hydrophobic pocket in HRVs is maintained to allow an induced transition around the GH loop that is required for productive uncoating. This transition could be induced by acidification in the endosome, receptor binding, or a combination of the two. Filling this pocket in VP1 with a drug or naturally occurring factor would inhibit this transition, thus inhibit uncoating. As expected, binding of compounds in the VP1 pocket has also been shown to inhibit intracellular uncoating as well as either acid-or heat-induced uncoating of the virus [28,29,47]. An attractive hypothesis suggests that the GH loop transition precedes or allows the externalization of VP4, which would be required for the formation of the uncoating intermediate particles. Remember, VP4 contains the myristoyl moiety, which can signal VP4 to associate with a membrane. The VP4 would drag the N-terminal region of VP1 (to which it is closely associated) to the exterior of the virion. This would result in a particle with the N-terminus of VP1 exposed, as has been observed in poliovirus [48–50]. The sequence of this exposed region of VP1 suggests that it can form an amphipathic helix. The VP1 helices could then insert into the membrane and form a pore, which could allow the passage of RNA through the lipid bilayer into the cytosol. This is reminiscent of the pore formation by colicin [51]. The observation that both the Ca2+ ion plus the β cylinder and VP4 become disordered under acidic conditions are consistent with this hypothesis. These are regions that would need to disorder to allow the externalization of VP4 and the N-terminus of VP1. IV. Capsid-Binding Compounds Capsid-binding compounds were discovered long before the emergence of the HRV crystal structure [52]. They were initially discovered on screening of compounds that had been produced by an insect pheromone project at Sterling Winthrop. Examples of compounds known or presumed to bind in this pocket are shown in Figure 5 [52–69]. The prototypical WIN drug contains an oxazoline ring attached to a phenoxy group, which is in turn linked by an aliphatic chain to an isoxazole ring. The three rings will be termed A, B, and C here (Figure 6). Compounds of this type were the first to be shown to inhibit the viral uncoating and also to stabilize the virion to heat-induced denaturation [29].

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Figure 5 Some compounds which have been shown to inhibit picornavirus replication. These are thought to bind in the VP1 hydrophobic pocket. References are indicated on the figure.

The structure of the first compound to be solved in complex with HRV 14 was found to be bound in an extended conformation within the VP1 hydrophobic pocket [24]. The compound is almost entirely buried within the capsid of the virion. Since in the native HRV14 the pocket exists in a closed configuration, binding required large motions in VP1 to accommodate the drug. These

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Figure 6 Some WIN compounds depicting different rings and linkers. Note WIN 52452 has no C ring.

motions, of up to approximately 4.5 Å, are most pronounced in the region of the GH loop. In contrast, HRV1A and HRV16, a minor and major receptor group virus respectively, have their pockets in open conformations even in the absence of drug [13,15]. For these serotypes, drug binding typically shifts positions of capsid atoms a maximum of 1 to 2 Å, smaller than those shifts seen in HRV14. Drugs bind in these pockets in a fashion similar to that seen in HRV14, extended and almost entirely buried by VP1. The structures of five HRVs have been solved to date: HRVs 1A, 3, 14, 16, and 50. In all of these HRVs, as well as the polio- and coxsackie viruses, VP1 hydrophobic pockets have been observed [5,12,13,15,17,24,70,71] (HRV3, Zhao, R. et al., personal correspondence; HRV50, Giranda V. L. et al., unpubhttp://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_499.html (1 of 2) [4/9/2004 12:36:54 AM]

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Figure 7 HRV14 VP1 hydrophobic pocket schematic with WIN 61605 illustrating some of the terminology commonly used to describe this pocket. Notice the GH loop separates the pocket from the canyon floor.

lished data). These pockets all share similar features and have been described as foot shaped, with a hydrophobic toe region, a heel region capable of hydrogen bonding, and a pore region near the ankle of the foot (Figure 7). Many of the picornavirus structures have been shown to have electron density in their VP1 pockets even in the absence of any added drug. These densities have been modeled as fatty acids or similar compounds [12,15,56,70–72]. The occurrence of these pocket factors have led some to hypothesize that these factors perform a similar function as do capsid-binding inhibitors, that is, to stabilize the virions [15,24,41,73,74]. Teleologically one could argue that the virion would pick up a fatty acid in its VP1 pocket before its egress from the cell, which would then stabilize the virion in transit to new hosts. The HRVs are known to be stable for long periods of time on surfaces and the dominant mode of transmission is thought to be hand-to-hand contact [75,76]. When a new host cell is reached, the stabilization factor might exit the pocket. This would allow the necessary conformational transition (probably at the GH loop) to occur and allow productive uncoating. A. Multiple Targets The use of these structures in a traditional structure-based drug design approach has been limited by the large number of unknown target serotype structures. There have been useful studies that have grouped picornaviruses based on their susceptibility to various antiviral agents (see below) [7,77]. However, in order to design a truly broad-spectrum single drug, the structural elements common among many serotypes must be considered.

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Because of the large number of serotypes, a successful inhibition strategy needs to consider whether it is better to have a drug that is exceedingly potent against a small subset of HRV serotypes (e.g. 30%) or a drug that is somewhat less potent but effective against most serotypes (e.g. 90%). Because of these considerations, different parameters are required to describe viral inhibition. Two important values that have been used extensively to describe potency for antipicornaviral compounds include the mean inhibitory concentration (MIC) and the MIC80. The MIC is the concentration that inhibits the viral progeny production by 50% in a cell-based plaque assay. The mean MIC is the average MIC over the number of serotypes against which the compounds have been tested. The MIC80 is the MIC at which at least 80% of the serotypes tested will be inhibited by at least 50%. Another way to think about the MIC80 is that it is the MIC value for the serotype that is at the 80th percentile rank for viruses inhibited. For example, if ten viruses were tested, the MIC80 would be the MIC concentration of the drug that inhibits the 8th most sensitive virus [15]. B. Potency and Binding Energetics It would be reasonable to assume that the activity of a drug (MIC) against a specific serotype would be related to its binding energy. This is an important consideration because algorithms that are used to predict potency rely on estimations of binding energy. The only experiment completed to directly study this correlation suggests a rough correlation in a small number of samples. This study however was limited to a small number of compounds in a single chemically similar series [78]. It is unclear whether this relationship will hold over diverse chemical entities. One could imagine a series of compounds that binds, but is ineffective at stabilizing the virion to any extent, thus ineffective in inhibiting viral replication. This appears to be what was observed when fragments of WIN compounds that only contained the A and B rings were examined. These fragments were less able to stabilize the virus to heat-induced denaturation than intact compounds. Although the binding constant for these compounds has not been determined, the diminished thermostabilization occurred at concentrations of compound sufficient to allow the compounds to be seen bound in the VP1 pocket via x-ray crystallography [79]. This observation suggests that the drug binding and inhibition have been decoupled to some degree. Another question concerning correlating binding affinity to viral inhibition is the number of sites per virion required to be occupied before the virion can no longer uncoat: all 60, or a few? Further, is there any positive or negative cooperativity in the interaction? These questions have not yet been answered.

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C. Structure-Activity Relationships Structural features, as stated above, have been found that are common to the known HRV structures. These features place constraints on compounds that have been demonstrated in a large number of structure—activity relationships. Shape considerations, hydrophobicity requirements, hydrogen-bonding requirements and compound flexibility will be discussed. Pocket-Shape Considerations Compound Length. The results of x-ray studies on several HRV serotypes have shown that the hydrophobic pockets vary in size, but all pockets impose structural constraints on compounds. The pockets, which almost completely enclose the compounds, require that both drug length and width must be limited. The results of structure—activity relationships for one series of WIN compounds have shown that the optimum length of the aliphatic linking region between the phenoxy (B ring) and isoxazole (C ring) ring is between 3 and 6 carbons (Table 2). This optimum length will vary in other series based on the extent of substitution on the A and C rings (see below) [54,80–82]. This effect on linker length is obvious both for individual serotypes like HRV14 and for the MIC80, which measures many serotypes. This effect of pocket length can also be seen when substituents are added to either the A or C ring which would lengthen the compounds. Additions of alkyl chains of four or more carbons tend to decrease potency, whereas shorter chains may increase potency (Table 3) [54,83]. In studies on compound length, HRV14 appears to be more sensitive to longer compounds than are many other rhinoviruses. This is consistent with the observation that the hydrophobic pocket of HRV14 is longer than that of HRVs

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1A, 16, and 50. Cluster analysis examining the sensitivity of 100 different picornaviruses against 15 different inhibitors showed that the sensitivities of different viruses depends on the shape of the pocket, with shorter pockets preferring shorter compounds [7]. This analysis is consistent with the structural observations, i.e., the viruses with long pockets (e.g., HRV14 and polioviruses) cluster together. A cluster distinct from that which contains HRV14, and prefers shorter compounds, is the group that contains the shorter pocket viruses: HRVs 1A, 16, and 50 (Figure 8). It is interesting that these drug sensitivity groups do not correspond to the receptor binding groups of the viruses. For example, both HRV14 and 50 are major receptor group viruses, but the pocket of HRV50 is clearly shaped more like the pocket of the minor receptor group virus 1A. The argument that the two different drug sensitivity groups may constitute distinct classes of HRVs has been bolstered by genetic examination, which has shown that clusters with similar amino acid sequences correlate well with the drug sensitivity [84]. As with the pocket shapes, these genetic clusters do not correlate with the two receptor groups into which the HRVs are placed. Attempts have been made to break through the length barrier imposed by the pocket structure. One could conceive of a drug that could pass through the pore at the heel of the pocket and connect the VP1 hydrophobic pocket to the canyon floor. Such a drug may have activity that could inhibit uncoating via stabilization of VP1 as well as inhibit attachment by directly blocking the receptor site. Attempts have been made to synthesize just such compounds, with extended “tails” attached to the C-ring. While these compounds bind, their tail does not extend through the pocket pore, but rather coils up within the pocket [85].

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Figure 8 Solvent accessible surfaces (dot surfaces) of (a) HRV1A, (b) 14, (c) 16, (d) 50 filled with WINs 56291, 61605, 56291, 61209 respectively (ball and stick). In all cases the cutaway view of the pocket has the viewer looking from inside the virion. The pocket of HRV14 is narrower and longer than the others and this is reflected in the drug structure—activity relationships (Tables 1–3).

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Compound Width. Bulk considerations have also been examined for the phenoxy or B ring [80]. In HRV14, which has a long narrow pocket, compounds with no substituents on the phenoxy ring were the most potent. However most other serotypes were more sensitive to compounds with substituents on the phenoxy ring, either a dimethyl or a dichloro. The serotypes that preferred disubstituted compounds included HRV1A and 50, both known to have wider, shorter pockets (Table 4). This is also in agreement with the drug-clustering analysis discussed above [7]. Induced VP1 Pocket Changes. The shape constrains discussed above are by nature somewhat qualitative. The precision with which one could predict the binding energy of a compound based on its shape is limited by the ability of the pocket to conform to its occupant. Conformational changes within the pocket are most pronounced between native and drug-bound HRV14, but present to a lesser extent in both HRV1A and 16. The extent of conformational changes produced by different drugs on the same virus may differ. This observation is illustrated in a study of a compound SCH 38057 in HRV14. This compound has a substantially different structure than the WIN compounds, and when bound to HRV14, induces changes in HRV14 that are also quite different from changes resulting from the binding of a WIN compound [62]. The ability of compounds to effect conformational changes in the capsid when bound and the variability of these changes both between drug classes and viral serotypes have made a priori predictions of potency based on shape considerations very difficult. Comparative Molecular Field Analysis (CoMFA) studies has been useful, when examining similar compounds in a single serotype (HRV14), in demonstrating spatial constraints [85], but generalizing these constraints over many serotypes and drug classes is much more difficult.

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Hydrophobicity Requirements Drug binding is enhanced by hydrophobicity in that portion of the drug that binds to the pocket toe. Quantitative structure-activity relationship (QSAR) analysis of these compounds have consistently shown that the most predictive parameter of antiviral activity is a measure of hydrophobicity, the octanol:water partition coefficient (logP) [80,82,85]. These studies have also consistently shown that there is no apparent correlation between electrostatic potential or dipole moment and potency. This evidence suggests that the drug—pocket interactions in the toe of the pocket are low-intensity hydrophobic interactions. Supporting this hypothesis is the finding that many structurally diverse molecules can be accommodated in this region, and even structurally similar molecules can bind in distinctly different conformations. Closely related structures have been shown to shift along their long axis by as much as 1.6 Å with respect to their phenoxy substituents [55]. There is even more variability with respect to the isoxazole placement (Table 5, Figure 9). No single hydrogen-bonding or electrostatic interaction appears to predominate within the pocket toe.

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Figure 9 A solvent-accessible surface of HRV14 (from the 61605 bound structure, dot surface) overlaid with WIN 61605 (ball and stick), 51711, 56826, and 56291 (a). The atoms closest to the toe pocket wall (top) for each compound are in approximately the same position, while the isoxazole ring (C ring) positions vary significantly. The atom closest to the toe pocket wall is quite different in (b), which compares WIN 61605 (ball and stick) to R61837 (tubes) and SCH 38057 (lines).

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The correlation of potency with logP might suggest that A rings with the lowest hydrogen-bonding potential would be the most potent. This has not been shown to be true. In fact, rings with heterocyclic nitrogen atoms are preferred to furan and thiophene rings, which would be expected to have less hydrogen-bonding potential [86,87]. This observation even extends to tetrazole rings, which have often been used in pharmaceutical design to replace hydrophilic groups such as carboxylates or esters. Explanations for the variability in potency due to A-ring changes have not been satisfactory. These heterocycles lack any consistent pattern of hydrogen bonds with the pocket residues (Table 6) [88]. Like QSAR analysis, structural analysis has not been able to show any relationship between hydrogenbonding groups in the A ring, or dipole moment, and potency [80,85]. Extensive structural characterization of many different A-ring heterocycles has not yet been done. Difficulties predicting relative potency of these compounds a priori stem from the lack of understanding of solvation/desolvation effects as well as difficulties in characterizing the low-intensity hydrophobic interactions. Consequently, it seems likely that new structure—activity relationships about the A-ring heterocycle will continue to be determined based on empirical findings. Hydrogen-Bonding Requirements Capsid-binding compounds with a hydrogen-bond accepting atom in the region of the drug that binds to the VP1 pocket heel appear to demonstrate greater potency than do their non-hydrogen-bonding counterparts (Table 7) [55,89]. Studies using HRV14 where Asn1219, a hydrogen-bond donor in the pore region, has been mutated to Ala have shown that the compounds bind as well in the mutated virus as in the native virus [27]. This might suggest that the hydrogen bonding to Asn1219 at the pocket pore is unimportant. Recent structures of compounds in HRV14 have shown that other hydrogen-bond donors can function in place of Asn1219, even if Asn1219 is still present. These groups in HRV14 are the hydroxyl of Ser1107 and the backbone nitrogen of Leu1106. Other hydrogen-bonding groups are present in other rhinoviruses that coordinate to tightly bound waters, which can also act as hydrogen-bond donors [55,56]. This provides a flexible hydrogen-bonding network that can then

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accommodate a variety of different hydrogen-bonding groups in the C ring of the drug (Figure 10). This hydrogen-bonding network has also been observed in all classes of capsid-binding compounds that have had their structures determined in HRVs [58,62]. Reports of compounds that bind to the capsid and inhibit picornavirus uncoating, but would seem to have little hydrogen-bonding potential (e.g., dichloroflavan) have not been structurally examined [60]. The effect of adding a hydrogen-bonding group to these compounds near the pocket pore is unknown, but one might expect that the potency of the compound would improve. Remember that WIN compounds without a hydrogen-bonding group at the pore region still have antiviral activity, albeit this activity is much weaker than that of an analogous drug with the hydrogen-bonding group present.

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Figure 10 Possible hydrogen bonds in the pore region of HRV14 bound to WIN 61605 (ball and stick). The waters (spheres W1 and W2) could potentially form hydrogen bonds to WIN 61605 as well as the side chains of Asn1219 and Ser1107 as well as the backbone of Leu1106 (residues highlighted as tubes). The viewer is looking from the virion exterior.

The fluidity of the hydrogen-bond donation network at the pore could allow a large number of structurally distinct molecules to bind in this location. If the function of the pocket is to bind fatty acids or other structurally diverse pocket factors and thus stabilize the virion. The flexibility of the hydrogenbonding network at the pore seems ideally suited for such a purpose. Drug Flexibility The WIN compounds all contain a flexible linking region that allows them to conform to differently shaped interiors of VP1 hydrophobic pockets. This flexibility is seen in many compounds that share the WIN-drug mechanism of action. However, some compounds do not contain a region as obviously flexible as an aliphatic linker (e.g., those with unsaturated rings as linkers, Figure 5). Flexibility in the aliphatic linking region of the WIN compounds has been explicitly studied and the results suggest that such a property is important for http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_512.html (1 of 2) [4/9/2004 12:44:42 AM]

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broad spectrum activity (Table 8) [56,90]. While certain less-flexible compounds may have increased potency for a particular serotype of HRV, these compounds have lower potency versus other serotypes. It is not known if the effect of flexibility is an equilibrium or kinetic effect. The flexibility might allow the compounds to expand or contract to fill available space in the VP1 hydrophobic pocket. Alternatively, the flexibility may allow the compounds to achieve a conformation required to enter or leave the pocket, but this conformation would not be seen in the crystallographic experiment. If this is true, modeling of the equilibrium structure of compounds in the pocket will not be accurate predictors of compound potency. Structure—Activity Relationship Summary The combination of classical structure—activity relationships combine with the structures of several compounds in a number of HRV serotypes have led to a paradigm for designing potent compounds. The effects of pocket shape, requirements for hydrophobicity in the toe of the pocket, as well as the potential for hydrogen bonding in the heel region appear to be strong determinants of antiviral potency. The requirement for a broad spectrum has so far limited a more detailed paradigm, such as one that might exist for designing inhibitors to a specific serine protease. In spite of this vagueness of antirhinoviral structure—activity relationships, they have been useful in guiding new compound synthe-

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sis. Large numbers of compounds, which might have been synthesized in the absence of any structural knowledge, have been eliminated from the list of inhibitors to be created, because these compounds would not have fit well into the VP1 pocket due to its finite size and hydrophobic nature. The goal of being able to predict with greater accuracy the potency and spectrum of compounds before they are synthesized awaits three developments: the structures of more HRVs and compounds, the ability to more accurately model hydrophobic interactions, and probably the most difficult, the ability to predict changes in the HRVs that occur due to drug binding. V. Viral Resistance As would be expected for any virus, particularly an RNA virus, resistance to capsid-binding compounds has been observed in the laboratory. The effect such mutations have on the life cycle of the virus is important. Mutations that occur in regions where changes are poorly tolerated, because that region serves a vital function in the viral life cycle, are likely to lead to less virulent viruses. The almost universal inhibition of rhino- and enteroviruses by capsid-binding compounds, coupled with the observation that all of the viruses with determined structures have a VP1 hydrophobic pocket, suggests that these pockets serve an important and similar function. It seems unlikely that so many viruses would maintain such a pocket if it were not a selective advantage. Drug-resistance mutants have been classified into two groups, exclusion mutants and compensation mutants. A. Exclusion Mutants The exclusion mutants' behavior has been readily and adequately determined by biochemical and crystallographic means [91,92]. The mutations occur within the hydrophobic pocket of VP1 and thermostabilization studies have shown that these mutations preclude the binding of drug in the VP1 pocket. One mutation site in HRV14 has been located at position 1188, which is on the side of the pocket closest to the viral interior, away from the canyon. Mutations at this site that convey resistance are Val rarrow.gif Leu or Val rarrow.gif Met. In both cases the mutation is to a larger side chain, which would be expected to fill the pocket. The crystal structure of the Val rarrow.gif Leu mutation confirms this hypothesis and demonstrates that the Leu side chain occupies space that would normally be occupied by an antiviral drug. The second site found in HRV14 is at Cys1199. Mutations to Phe, Tyr, Trp, and Arg all confer resistance at this site. Again, all of these mutations are to larger side chains. The hypothesis that these mutations function by excluding

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drug from the pocket is confirmed by crystallographic analysis showing that the Phe side chain in the Cys rarrow.gif Phe mutation occupies the site which would be occupied by a drug if it were bound. B. Compensation Mutants More intriguing and more difficult to understand are the compensation mutants. These viruses bind compounds within their hydrophobic pockets, but are still able to replicate and are therefore able to compensate for the bound drug (Table 1, Figure 4). These mutations, like the mutations that convey acidresistance phenotypes, cluster about the hydrophobic pocket of VP1 [27,28,92]. Two hypothesis have been presented in order to explain the behavior of the compensation mutants. The first suggests that there is a link between the binding of receptor by virion and its ability to bind compounds in the VP1 pocket. The second hypothesis suggests that the effect of a mutation on protein stability leads to the drug-resistant phenotype. These two hypotheses are not mutually exclusive. Linked-Binding Hypothesis The compensation mutants can be subdivided into those that occur on the canyon floor and those that are inside the VP1 hydrophobic pocket. The linked-binding hypothesis suggests that the subset of compensation mutations that occur on the canyon floor increase the binding affinity of receptor. An increase in binding for one of these mutants, Val1153 rarrow.gif Ile, has been observed [28]. This increased binding would then in turn cause a decrease in the affinity of the capsid for the drug. This would result in the drug being less effective in inhibiting uncoating. The second subset of compensation mutants are those that occur inside the hydrophobic pocket but do not interact with the receptor binding site. These mutations might directly decrease drug-binding affinity in the VP1 pocket due to decreased hydrophobic interactions caused by the smaller amino acid side chains [27]. In both subsets of mutations, the binding affinity of a drug is reduced either directly or through the effects of receptor binding. This decreased affinity for drug would be displayed as a drugresistant phenotype. Stabilization—Destabilization Hypothesis An alternative explanation for the behavior of the drug compensation mutants suggests that these mutants allow for increased conformational flexibility in the capsid. This increased conformational flexibility would compensate for the decrease in flexibility, which is manifested by decreased acid or thermal lability, induced by the binding of drug. The increase in flexibility would allow a

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conformational transition to occur, presumably near the GH loop, that would be required for viral uncoating. The conformational transition would lead to productive uncoating, even in the presence of bound drug. This hypothesis suggests that drug-resistant mutations would be of the type that destabilize protein structures. Such mutations are typically from larger to smaller side chains or from highly branched to less branched side chains [43–45]. Four of five observed compensation mutants are of this type (Table 1). The remaining mutant, Val1153 rarrow.gif Ile, while a priori might not be predicted to destabilize the capsid, has been shown experimentally to decrease viral thermostability when compared with native HRV14 [28]. The advantage of the stabilization-destabilization hypothesis is that it allows for a single mechanism of resistance for both compensation mutants occurring in the hydrophobic pocket as well as those that are outside the pocket near the receptor binding site. Both of these sites are close to the GH loop of VP1 that becomes disordered under acidic conditions. This stabilization—destabilization hypothesis also explains the behavior of acid-resistant mutants, which are predominately of the type which should stabilize a protein to conformational changes. Wild-type viruses have a preferred stability profile, which allows uncoating under the proper conditions. Mutations or addition of drug can perturb this profile, resulting in decreased replication. A second perturbation might then compensate for the first, restoring virulence (Figure 11). This behavior has been observed in HRV1A, in which mutants have been isolated that require presence of drug to replicate. These mutations are more acid-labile

Figure 11 The effects of various rhinovirus manipulations. The solid line depicts the native rhinovirus uncoating profile. Mutations and drugs can effect this profile to make the virus more or less stable to pH- or temperature-induced changes.

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than the wild-type virus. At the optimal drug concentration for growth, the pH-stability of these mutations is equivalent to that of native HRV1A in the absence of drug. Increasing drug concentration beyond that which is optimal for growth further increases the acid-stability of these mutant viruses so that they are more acid-stable than native virus without drug (Daniel Pevear, personal communication). C. Clinical Resistance and Virulence The clinical importance of these mutations has not been clearly demonstrated. In at least one case a drugresistant mutant was able to grow in vitro in single-cycle growth experiments as well as wild-type HRV14 [28]. Frequently, however, drug-resistant or acid-stable mutants will not grow as well in cell culture as native virus [93]. The final analysis of the clinical importance of resistant mutants awaits the results of clinical trails. In the one study published to date, HRV2 mutants of the compensation type (resistant to the capsid-binding compound chalcone) were tested in healthy human volunteers [65]. The drug-resistant mutants caused significantly fewer colds than the normal virus. Mutants that require the presence of drug to grow did not cause any apparent disease. In this single case it appears that the mutant viruses were not as virulent as the parent strain. VI. Animal and Clinical Studies The idea that an inhibitor that binds to a nonenzymatic, nonreceptor site of a virion could inhibit viral replication in vivo would a priori be considered to be an unlikely scenario by most in the field of structure-based design. If this idea were proposed knowing only the structure of the native virus (especially HRV14, which has a closed-pocket conformation) skepticism would abound. This type of project arose not from a structure-based approach, but from the tried-and-true screening approach. The compounds were first shown to be effective in inhibiting viral replication in vitro [52]. This was followed by an experiment showing that these compounds could inhibit at least one enterovirus in a mouse model [94]. In this study, suckling mice were inoculated with an echovirus and if untreated they developed a fatal paralytic illness. When treated either prophylactically or within a few days of the viral inoculation, virtually all of the mice were protected. This is dramatic proof of the concept that these compounds can inhibit picornaviral infection in vivo. There are important differences between mice and men and between enteroviral paralytic illness and rhinoviral upper respiratory tract infections.

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The differences between mice and men should be obvious to the casual reader. Enterovirus-induced paralytic illnesses are systemic infections, and the virus must at some point move through the body and be subject to circulating drug. Upper respiratory infections resulting from rhino-or enteroviruses tend to be localized to the pharynx, which would require that drug titers remain high in this region of the body. In spite of these difficulties, there have been some dramatic successes in clinical trials of antirhinovirals, although clearly there are still obstacles to overcome. Two different classes of antipicornavirus compounds have been shown to be successful at inhibiting at least some types of infection when administered prophylactically. The WIN compound 54954, given orally, has been shown to inhibit Coxsackie virus A21 in humans [6]. The Jaansen compound R77975 has been shown to inhibit HRV9 when administered intranasally 6 times daily [95,96], but R77975 has not been shown to inhibit infection when used therapeutically (after symptoms occur). Further clinical trials with WIN 54954 were suspended due to the developments of side effects. More recently VP 63843 has been shown to have dramatically improved effect when compared with WIN 54954 in the prophylactic treatment of Coxsackie virus A21 infection in humans and it will be tested therapeutically. The inability of R77975 to show a therapeutic effect is more likely due to poor pharmacodynamics rather than a fundamental inability for this compound to inhibit an established infection. The pharmacodynamic problem was witnessed in the R77975 study, which showed that administration intranasally 3 times a day did not yield prophylactic protection. Yet if administered 6 times daily the prophylaxis occurs. Therefore if more potent and metabolically stable compounds are found, more positive clinical results would be expected. VII. Future Directions The structures of the picornaviruses (native, with receptor bound, in the presence of acid, with a myriad of compounds bound, and of acid- and drug-resistant mutants) have yielded valuable information about possible molecular mechanisms for their uncoating. These same studies have suggested the mechanism by which these uncoating inhibitors work. A by-product of this research is the hypothesis that these compounds may mimic naturally occurring factors that occupy the VP1 pocket. The hunt for these natural compounds and their significance is underway. This understanding of the mechanism of action, as well as the structure-activity studies, have also yielded valuable information for future development of antipicornaviral drugs. The determination of compound's size, shape, and other physical requirements for activity will also be of assistance.

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From the animal and human experiments it is clear that the cure for some common colds is within reach. These therapies are also quite likely to be efficacious in enteroviral diseases, for example in cardiogenic coxsackie virus infection [97]. Acknowledgments We would like to acknowledge that there have probably been hundreds of people who have contributed to work reviewed here. We would like especially to thank the many people at ViroPharma, Sterling Winthrop, Purdue University, and the University of Wisconsin at Madison with whom we have worked over the years. We would also like to thank Dirksen Bussiere and Yvonne Martin for reading and editing this manuscript prior to submission. References 1. Rueckert RR. Picornaviridae and their replication. In: Fields BN, Knipe DM, eds. Virology. New York: Raven Press, 1990:507–548. 2. Sperber SJ, Hayden FG. Chemotherapy of rhinovirus colds. Antimicrob Agents Chemother 1988; 32:409–419. 3. Uncapher CR, DeWitt CM, Colonno RJ. The major and minor group receptor families contain all but one human rhinovirus serotype. Virology 1991; 180:814–817. 4. Hamparian VV, Colonno RJ, Cooney MK, et al. A collaboration report: rhinoviruses—extension of the numbering system from 89 to 100. Virology 1987; 159:191–192. 5. Muckelbauer JK, Kremer MK, Minor I, et al. The structure of coxsackievirus B3 at 3.5 Å resolution. Structure 1995; 3:653–667. 6. Schiff GM, Sherwood JR, Young EC, Mason JL, Gamble JN. Prophylactic efficacy of WIN 54954 in prevention of experimental human coxsackie A21 infection and illness. Antivir Res 1992; 17 (Suppl 1):92. 7. Andries K, Dewindt B, Snoeks J, et al. Two groups of rhinoviruses revealed by a panel of antiviral compounds present sequence divergence and differential pathogenicity. J Virol 1990; 64(3):1117–1123. 8. Flower RJ, Moncada S, Vane Jr. The pharmacologic basis of therapeutics. In: Gilman AG, Goodman LS, Gilman A, eds. Analgesic-antipyretics and anti-inflammatory agents; drugs employed in the treatment of gout. New York: Macmillan, 1080:682–728. 9. Drake JW. Rates of spontaneous mutation among RNA viruses. Proc Natl Acad Sci USA 1993; 90(9):4171–4175.

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10. Mast EE, Harmon MW, Gravenstein S, et al. Emergence and possible transmission of amantadineresistant viruses during nursing home outbreaks of influenza A (H3N2). Am J Epidem 1991; 134(9):988–997.

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25. Arnold E, Rossmann MG. Analysis of the structure of the common cold virus, human rhinovirus 14, refined at a resolution of 3.0 Å. J Mol Biol 1990; 211:763–801. 26. Towler DA, Gordon JI, Adams SP, Glaser L. The biology and enzymology of eukaryotic protein acylation. Annu Rev Biochem 1988; 57:69–99. 27. Hadfield AT, Oliveira Ma, Kim KH, et al. Structural studies on human rhinovirus 14 drug-resistant compensation mutants. J Mol Biol 1995; 253:61–73. 28. Shepard DA, Heinz BA, Rueckert RR. WIN 52035-2 inhibits both attachment and eclipse of human rhinovirus 14. J Virol 1993; 67(4):2245–2254. 29. Fox MP, Otto MJ, McKinlay MA. Prevention of rhinovirus and poliovirus uncoating by WIN 51711, a new antiviral drug. Antimicrob Agents Chemother 1986; 30:110–116.

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30. Pevear DC, Fancher MJ, Felock PJ, et al. Conformational change in the floor of the human rhinovirus canyon blocks adsorption to HeLa cell receptors. J Virol 1989; 63:2002–2007. 31. Lonberg-Holm K, Korant BD. Early interaction of rhinoviruses with host cells. J Virol 1972; 9:29–40. 32. Korant BD, Lonberg-Holm K, Yin FH, Noble-Harvey J. Fractionation of biologically active and inactive populations of human rhinovirus type 2. Virology 1975; 63:384–394. 33. Everaert L, Vrusen R, Boeye A. Eclipse products of poliovirus after cold-synchronized infection of HeLa cells. Virology 1989; 171:76–82. 34. Hooverlitty H, Greve JM. Formation of rhinovirus-soluble ICAM-1 complexes and conformational changes in the virion. J Virol 1993; 67(1):390–397. 35. Neubauer C, Frasel L, Kuechler E, Blaas D. Mechanism of entry of human rhinovirus 2 into HeLa cells. Virology 1987; 158:255–258. 36. Zeichhardt H, Wetz K, Willingmann P, Habermehl KO. Entry of poliovirus type 1 and mouse elberfeld (ME) virus into HEp-2 cells: receptor mediated endocytosis and endosomal or lysosomal uncoating. J Gen Virol 1985; 66:483–492. 37. Gromier M, Wetz K. Kinetics of poliovirus uncoating in HeLa cells in a nonacidic environment. J Virol 1990; 68(8):3590–3597. 38. Madshus IH, Olsnes S, Sandvig. Mechanism of entry into the cytosol of poliovirus type 1: requirement for low pH. J Cell Biol 1984; 98:1194–1200. 39. Madshus IH, Olsnes S, Sandvig K. Different pH requirements for entry of the two picornaviruses, human rhinovirus 2 and murine encephalomyocarditis virus. Virology 1984; 139:346–357. 40. Perez L, Carrasco L. Entry of poliovirus into cells does not require a low-pH step. J Virol 1993; 67(8):4543–4548. 41. Giranda VL, Heinz BA, Oliveira MA, et al. Acid-induced structural changes in human rhinovirus14—possible role in uncoating. Proc Natl Acad Sci USA 1992; 89(21):10213–10217. 42. Skern T, Torgersen H, Auer H, Kuechler E, Blaas D. Human rhinovirus mutants resistant to low pH. Virology 1991; 183:757–763. 43. Alber T, Dao-pin S, Nye JA, Muchmore DC, Matthews BA. Temperature-sensitive mutations of bacteriophage T4 lysozyme occur at sites with low mobility and low solvent accessibility in the folded protein. Biochemistry 1987; 26:3754–3758.

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44. Argos P, Rossmann MG, Grau UM, Zuber H, Frank G, Tratschin JD. Thermal stability and protein structure. Biochemistry 1979; 18:5698–5703. 45. Matthews BA. Genetic and structural analysis of the protein stability Problem. Biochemistry 1987; 26:6885–6888. 46. Perutz MF, Raidt H. Stereochemical basis of heat stability in bacterial ferredoxins and in haemoglobin A2. Nature 1975; 255:256–259. 47. Gruenberger M, Pevear D, Diana GD, Kuechler E, Blaas D. Stabilization of human rhinovirus serotype 2 against pH-induced conformational change by antiviral compounds. J Gen Virol 1991; 72:431–33. 48. Fricks CE, Hogle JM. Cell-induced conformational change in poliovirus: Externalization of the amino terminus of VP1 is responsible for liposome binding. J Virol 1990; 64:1934–1945.

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49. Roivainen M, Piirainen L, Rysa T, Narvanen A, Hovi T. An immunodominant N-terminal region of VP1 protein of poliovirion that is buried in the crystal structure can be exposed in solution. Virology 1993; 195:762–765. 50. Yafal AG, Kaplan G, Racaniello VR, Hogle JM. Characterization of poliovirus conformational alteration mediated by soluble cell receptors. Virology 1993; 197:501–505. 51. Stroud R. Ion channel forming colicins. Curr Opin Struc Biol 1995; 5:514–520. 52. McSharry JJ, Caliguiri LA, Eggers HJ. Inhibition of uncoating of poliovirus by arildone, a new antiviral drug. Virology 1979; 97:307–315. 53. Abel MD, Cameron AD, Ha CM, et al. Novel azoylalkyloxy compounds with antipicornaviral activity. Antivir Chem Chemother 1995; 6(4):245–254. 54. Diana GD, Cutcliffe D, Volkots DL, et al. Antipicornavirus activity of tetrazole analogues related to disoxaril. J Med Chem 1993; 36(22):3240–3250. 55. Giranda VL, Russo GR, Felock PJ, et al. The structures of four methyltetrazole-containing compounds in human rhinovirus serotype 14. Acta Cryst D 1995; D51:496–503. 56. Kim KH, Willingman P, Gong ZX, et al. A comparison of the anti-rhinoviral drug binding pocket in HRV14 and HRV1A. J Mol Biol 1993; 230:206–227. 57. Andries K, DeWindt B, Snoeks J, Willebrords R, VanEemeren K. Anthirhinovirus spectrum and mechanism of action of R77975. Antivir Res 1991; 1 Suppl:98. 58. Chapman MS, Minor I, Rossmann MG, Diana GD, Andries K. Human rhinovirus 14 complexed with antiviral compound R 61837. J Mol Biol 1991; 217:455–463. 59. Moeremans M, Raermaeker M, Daneels G, et al. Study of the binding of R 61837 to human rhinovirus 9 and immunobiochemical evidence of capsid-stabilizing activity of the compound. Antimicrob Agents Chemother 1992; 36(2):417–424. 60. Tisdale M, Selway JWT. Effect of dichloroflavan (BW683C) on the stability and uncoating of rhinovirus type 1B. Antimicrob Chemother 1984; 14(Suppl A):97–105. 61. Ash RJ, Parker RA, Hagan AC, Mayer GD. RMI 15,731 (1-[5-tetradecyloxy-2-furanyl]ethanone), a new antirhinovirus compound. Antimicrob Agents Chemother 1979; 16:301–305. 62. Zhang AQ, Nanni RG, Li T, et al. Structure determination of antiviral compound SCH-38057 complexed with human rhinovirus-14. J Mol Biol 1993; 230(3):857–867.

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63. Rozhon E, Cox S, Buontempo P, et al. SCH-38057—a picornavirus capsid-binding molecule with antiviral activity after the initial stage of viral uncoating. Antivir Res 1993; 21(1):15–35. 64. Ishitsuka H, Ninomiya YT, Ohsawa C, Fujiu M, Suhara Y. Direct and specific inactivation of rhinovirus by chalcone Ro 09-0410. Antimicrob Agents Chemother 1982; 22(4):617–621. 65. Yasin SR, al-Nakib W, Tyrrell AJ. Pathogenicity for humans of human rhinovirus type 2 mutants resistant to or dependent on chalcone Ro 09–0410. Antimicrob Agents Chemother 1990; 34:963–966. 66. Ninomiya Y, Shimma N, Ishitsuka H. Comparative studies on the antirhinovirus activity and mode of action of the rhinovirus capsid binding agents, chalcone amides. Antivir Res 1990; 13:61–74. 67. Kenny MT, Dulworth JK, Bargar TM, Torney HL, Graham MC, Manelli AM. In vitro antipicornavirus activity of the 6-substituted 2-(3',4'-dichlorophenoxy)-2H-

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86. Abraham MH, Duce PP, Prior DV, Barratt DG, Morris JJ, Taylor PJ. Hydrogen bonding. Part 9. Solute proton donor and proton acceptor scales for use in drug design. J Chem Soc Perkin Trans II 1989; 1355–1375. 87. Abraham M, Grellier P, Prior D, Morris J, Taylor P. Hydrogen bonding. Part 10. A scale of solute hydrogen-bond basicity using log K values for complexation in tetrachloromethane. J Chem Soc Perkin Trans II 1990; 521–529. 88. Bailey TR, Diana GD, Kowalczyk PJ, et al. Antirhinoviral activity of heterocyclic analogs of WIN54954. J Med Chem 1992; 35(24):4628–4633. 89. Bailey TR, Diana GD, Mallamo JP, et al. An evaluation of the antirhinoviral activity of acylfuran replacements for 3-methylisoxazoles. Are 2-acetylfurans bioisosteres for 3-methylisoxazoles. J Med Chem 1994; 37:4177–4184. 90. Mallamo JP, Diana GD, Pevear DC, et al. Conformationally restricted analogues of disoxaril—A comparison of the activity against human rhinovirus type-14 and type-1A. J Med Chem 1992; 35(25):4690–4695. 91. Badger J, Krishnaswamy S, Kremer MJ, et al. Three-dimensional structures of drug-resistant mutants of human rhinovirus 14. J Mol Biol 1989; 207:163–174. 92. Heinz BA, Rueckert RR, Shepard DA, et al. Genetic and molecular analyses of spontaneous mutants of human rhinovirus 14 that are resistant to an antiviral compound. J Virol 1989; 63(6):2476–2485. 93. Colonno RJ, Condra JH, Mizutani S, Callahan PL, Davies ME, Murcko MA. Evidence for direct involvement of the rhinovirus canyon in receptor binding. Proc Natl Acad Sci USA 1988; 85:5449–5453. 94. McKinlay MA, Frank JA, Steinberg BA. Use of WIN 51711 to prevent echovirus type 9-induced paralysis in suckling mice. J Infec Dis 1986; 154:676–681. 95. Al-Nakib W, Higgins PG, Barrow Gl, et al. Suppression of colds in human volunteers challenged with rhinovirus by a new synthetic drug (R61837). Antimicrob Agents Chemother 1989; 33:522–525. 96. Hayden FG, Andries K, Janssen PAJ. Safety and efficacy of intranasal Pirodavir (R77975) in experimental rhinovirus infection. Antimicrob Agents Chemother 1992; 36(4):727–732. 97. Ilback N-G, Wesslen L, Pauksen K, Stalhandske T, Friman G, Fohlman J. Effects of the antiviral WIN 54954 and the immune modulator LS 2616 on cachectin/TNF and gamma-interferon responses during viral heart disease. Scand J Infect Disease Suppl 1993; 88:117–123.

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20 The Integration of Structure-Based Design and Directed Combinatorial Chemistry for New Pharmaceutical Discovery Roger Bone and F. Raymond Salemme 3-Dimensional Pharmaceuticals, Inc., Exton, Pennsylvania I. New Challenges For Drug Discovery Rapid advances in cell and molecular biology, together with comprehensive genome sequencing efforts, are providing detailed correlations between specific pathological conditions and discrete molecular targets. The same tools of recombinant DNA technology that identify key gene targets also provide the means for target biosynthesis in quantities sufficient for both the high-throughput screening of compound libraries for leads and the structure-based refinement of leads using x-ray crystallography and NMR spectroscopy. The rapid expansion in genomics data makes it inevitable that targets will be identified whose functions are so poorly understood that the most rapid and efficient way to establish their involvement in disease will be through the development of prototype drugs. New approaches to drug discovery that are able to integrate many different types of information are needed to seize this opportunity and drive an optimally efficient discovery process. In what follows, we describe a practical integration of structure-based design and combinatorial chemistry aimed at enhancing the effectiveness of both approaches. Three-dimensional structures provide the information required to most efficiently direct the design and optimization of new lead compounds. Combinatorial chemistry technologies, which are based on high-throughput automated methods of chemical synthesis, produce new classes of lead compounds and permit the rapid generation of structure—activity relationships

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Figure 1 An integrated technology for drug discovery combines the precision of structure-based design with the parallelism of combinatorial synthesis. Chemi-informatics systems track and integrate all data emerging from the discovery cycle.

(SAR). Chemi-informatics plays a key role in this integration by assuring that properties important in drug development are both factored into compound design and cumulatively tracked throughout the discovery process (Figure 1). The product of this approach is a permanently useful set of drug-design parameters. The integration of these technologies promises to produce an increase in the efficiency of drug discovery and may ultimately offer a means for reducing the aggregate failure rate of compounds selected for development. One paradigm for achieving the integration of these technologies is described below. II. Structure-Based Design Structure-based drug design has become a highly developed technology that is in active use in most major pharmaceutical companies. Structure-based design is an iterative process in which lead compounds identified by screening, de novo design, or mechanism-based features are systematically elaborated to improve potency and specificity [1,2]. The process involves successive rounds of structure determination of lead—target complexes, design of lead modifications using molecular modeling tools, synthesis of new drug leads, and measurement of the chemical and biological properties of the modified leads using screens for target

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function. Iterative refinement and optimization of drug leads is an effective strategy for generating potent preclinical candidates. Structure-based design can also be used to design new chemical classes of compounds that present similar substituents to the target using a template or scaffold which is chemically distinct from previously characterized leads [3,4]. Structure determination typically relies on x-ray crystallography or high-field nuclear magnetic resonance to directly visualize the 3-dimensional structure of a molecular target and the structures of complexes of the target with drug leads. Alternately, many targets fall into identifiable classes that frequently enable the development of homology models of the 3-dimensional target structure or a mechanism-based strategy for drug-lead generation. Ongoing genome sequencing efforts have led to the identification of hundreds of potential therapeutic targets, many of which represent possible sources of crossover pharmacology. Homology modeling is a key feature of an integrated drug discovery effort because it allows this genomics information to be utilized early in the development of target ligands or in the engineering of ligand specificity. Although structure-based design is an effective technology, current limitations center on the inability to quantitatively predict how specific modifications of the lead will actually affect ligand binding affinity [5,6]. This reflects the complexity of the drug-binding process and our inability to accurately predict the conformational response of macromolecular structures to ligand binding. In addition, we have only limited ability to accurately calculate molecular energy parameters or to accurately estimate the effects of factors such as polarizability, solvation, and entropy that may have an important influence on drugbinding energetics. Although computational methods will continue to improve, most design work (and algorithms) still relies heavily on heuristic rules (Figure 2) that have been developed through experience and that guide the structural and medicinal chemists in the systematic modification of lead compounds [7]. As a practical consequence, many cycles of serial lead modification are required in order to produce molecules of suitable potency and specificity to be considered preclinical drug candidates. Structural information can increase the efficiency with which pharmacokinetic or toxicological liabilities in lead compounds are eliminated by suggesting where compounds can be modified so as to alter drug properties without affecting target potency. Structural data can also be used to direct de novo design of alternate and distinct chemical classes of lead compounds, each of which might be expected to have a different pharmacological profile [3,4]. New chemical compound classes can also be designed from existing lead compounds by recombining substituents and core regions (scaffolds) from existing lead compounds. Chemically distinct lead series can then be optimized in parallel so that when a preclinical candidate is found to have inadequate drug properties, a backup is immediately available for preclinical evaluation. As

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Figure 2 Some heuristic rules frequently used in structure-based drug design. The positions of “bound” water molecules are key indicators for lead modification sites.

outlined below, both the scaffold modification and structural recombination strategies are key components of an integrated drug discovery technology that combines structure-based design and combinatorial chemistry. III. Combinatorial Chemistry Combinatorial chemical technology enables the parallel synthesis of organic compounds through the systematic addition of defined chemical components using highly reliable chemical reactions and robotic instrumentation [8–11]. Large libraries of compounds result from the combination of all possible reactions that can be done at one site with all the possible reactions that can be done at a second, third, or greater number of sites. Combinatorial chemical methods can potentially generate tens to hundreds of millions of new chemical compounds as mixtures, attached to a solid support, or as individual compounds. The first combinatorial libraries with millions of members were oligopeptide and oligonucleotide libraries for which reliable and versatile chemical reactions already existed [10,12,13]. However, these libraries were not generally useful as a source of leads for small-molecule pharmaceuticals due to the relatively high molecular weight of the resulting leads and other limitations of oligonucleotides and oligopeptides as drugs. More recently, “drug-like” libraries have been produced that offer much greater utility as a source of drug leads (Figure 3). However, practical limitations of the versatility and reliability of the chemical reactions used to make these libraries have resulted in somewhat smaller sets of compounds [14–18].

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Figure 3 Generation of combinatorial libraries. Traditional combinatorial libraries based on polypeptides or polynucleotides were built up by oligomer condensation and achieved a high level of diversity through conformational flexibility. Small-molecule libraries of drug-like molecules can be built up using a variety of strategies that focus on maximizing chemical diversity while (generally) minimizing conformational flexibility.

From the drug discovery perspective, large combinatorial libraries have the same utility as conventional pharmaceutical or natural-product compound libraries, i.e., as a source of leads for new drugs. The design of large combinatorial libraries is driven by the requirement that individual reactions be highly reliable and versatile, while producing libraries with the highest possible degree of chemical diversity. Individual steps must be optimized, the compatibility of building blocks must be examined thoroughly and the synthesis must be automated. As a consequence, a significant investment in time and resources must be made before a library can actually be produced. Designed and used in this way, large combinatorial libraries provide a new source of screening leads. However, libraries with maximum diversity, which may be used for “blind” lead discovery, do not address the principle rate-limiting step in drug discovery, the elaboration of a suitable SAR around the lead compound after an active lead has been discovered.

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While initial combinatorial chemical strategies have focused on the exhaustive synthesis of all members of a given library, it is inevitable that advances in automated chemistry and equipment will soon make it possible to synthesize a vastly greater number of compounds than it will be practical to

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screen. For example, for a simple library created using amine and acid condensations onto a given amino acid scaffold, over a million compounds can be produced from commercially available reagents. If this simple library is expanded to incorporate the hundreds of commercially available or easily synthesized amino acids scaffolds, the number of compounds that can possibly be made increases to greater than 108. Although biological, chemical, and physical assays can be automated so that hundreds of thousands to millions of compounds can be screened annually, the process is expensive and the reliability of the screening process decreases when measurements are made on mixtures of multiple compounds. Various estimates of the number of drug-like molecules with molecular weight (MW)<1000 range from 1050 to 10130, making it clear that there is no way to exhaustively screen all of the compounds that can be made. What is needed instead are strategies for focusing the process of combinatorial chemistry to permit the rapid refinement of drug properties. IV. Integrated Structure-Based Design And Combinatorial Chemistry The limitations and strengths of structure-based design and combinatorial chemistry approaches are complementary. On a basic level, structure-based drug design suffers from the inability to predict the energetic effects of even the most conservative modifications of a lead compound. The result is that many compounds must be individually synthesized to achieve discovery program objectives. Combinatorial chemistry addresses this limitation by providing the methodology to synthesize many compounds in parallel so that an extensive array of structure-activity relationships can be developed. Predictions of the effects of modifying a particular substituent can then be based on empirical SAR data rather than ab initio or semi-empirical computations. Combinatorial chemical approaches can potentially produce millions of compounds, more than are desired or useful, which tax target-screening efforts and compound tracking. Furthermore, the approaches that allow such large numbers of compounds to be screened, using mixture [12,17], solid phase screening [19], or single point assays [16,18], generally result in a reduction in data quality, which makes the development of structure-activity relationships very difficult. Structure-based design methods address this problem by making the search for new leads and development of SAR more directed and by scaling back the numbers of compounds that are synthesized. Instead of producing all possible members of every library, efforts focus on the design of templates that can bias libraries towards a particular target class and the selection of the library members with the greatest probability of interacting with the target. Thus the integrated application of structure-based design and combinatorial chemical technologies can produce synergistic improvements in the effi-

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Figure 4 The information flow in a drug discovery process that connects elements of structure-based design and combinatorial chemistry for drug discovery.

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ciency of drug discovery (Figure 4). The process begins with the knowledge-based design of a library template or scaffold and involves the synthesis of small subsets of library members. As with structurebased design approaches, the process is an iterative one in which SAR data and structural data guide not only the iterative selection of library members for testing, but also the design of later generation library scaffolds. The integrated process is described in detail in the next few paragraphs.

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Figure 5 Generation of a virtual combinatorial library by finding substituents of a custom scaffold that can be accommodated in the binding site of a molecular target or meet other 3D structural criteria. Once the virtual library is synthesized in the computer, individual members can be selected using structural or additional criteria and synthesized using automated equipment.

The first step in the process is the generation of a chemical template or scaffold that can be derivatized at multiple sites using reliable chemical reactions to produce a large combinatorial library (Figure 5). This custom chemical template is designed based on the structure of the target using the same heuristic set of rules used for traditional structure-based drug design. Useful 3-dimensional pharmacophore models are best derived from crystallographic or nuclear magnetic resonance structures of the target, but can also be derived from homology models based on the structures of related targets or 3-dimensional quantitative structure-activity relationships (3D QSAR) derived from a previously discovered series of active compounds [20]. In addition, the mechanism of action of the target or any other information that exists regarding the target or the target class can be used in the design to maximize the chances of finding hits [21,22]. The combinatorial libraries are designed so that a few thousand to millions of discrete molecules can be produced by reaction of the custom-designed template with appropriate proprietary and commercially available chemical building blocks. The next step in the process involves the implementation of the automated synthesis and generation of the library. Significant lead time is anticipated before a library can be produced because even the most reliable chemical reactions require optimization if they are to be carried out by a robot, particularly if

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the reactions are to be implemented with the template attached to a solid support. When the synthesis is optimized and fully automated, thousands to millions of compounds are accessible. One of the key features of this process is that all of the compounds that can potentially be made by elaboration of a custom scaffold are first “made” in virtual form in a computer. Each of the chemical reactions required to create the library is encoded in a program that then systematically combines the template and the building blocks to create a 2-dimensional representation of each member of the library. Next, 3-dimensional representations are created and molecular-property descriptors are calculated for each member of the library. Molecular property descriptors encompass molecular connectivity, dipole moments, calculated partition coefficients, and many other calculable molecular properties. The virtual library of compounds can then be computationally screened and the library members ranked according to their ability to interact with the target receptor or 3-dimensional pharmacophore model [23–25]. The compounds can also be ranked by their inability to interact with any number of alternative targets whose inhibition is undesirable, or their ability to meet any range of desired chemical or physical properties that may be important in drug pharmacology. Alternately or additionally, compounds can be ranked according to their ability to span or sample the physical-chemical property space to produce the most diverse set of compounds for initial screening [26,27]. Small sublibraries of the large virtual library that best satisfy the selection criteria are chemically synthesized using automated methods and then the biological and/or chemical properties of each compound are measured using automated assays. The SAR data that emerge from the assays are stored in a central database and used in the selection process to drive additional rounds of sublibrary selection, synthesis, and assay. Multiple mathematical models are developed to correlate the computed structure and properties of each synthesized library member with the biological, chemical, or physical properties that are measured during each cycle of testing [28]. A key feature of this approach is that compounds can be selected not only on the basis of which are predicted to perform the best in the target assay but also on the basis of their ability to perform the best in the target assay but also on the basis of their ability to distinguish between or validate the SAR models that are generated. The observed and predicted properties of a given sublibrary are compared so that the set of assumptions upon which property refinement is based is constantly updated. In principle, this process can become completely automated so that leads are discovered and refined with very little manual intervention [28]. To achieve the greatest improvements in drug discovery efficiency, empirical data of various kinds must be collected throughout the iterative refinement process. It is desirable to obtain more accurate dissociation constants rather than IC50 or single-point percent-inhibition values. In addition, the 3dimensional structures of interesting target—inhibitor complexes are determined

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Figure 6 The combination of structure-based design and combinatorial chemistry can facilitate the generation of recombined compounds to rapidly produce potent compounds with maximum chemical and structural diversity.

to provide information regarding how substituents are interacting with the target and when binding modes change. This information may be essential if SAR models are to remain predictive over large numbers of compounds. The integrated drug discovery process utilizes as much information as is available to find and optimize initial lead compounds. Because the automated synthesis of large libraries of compounds requires reliable and versatile chemical reactions, initial libraries are designed to discover new chemical lead classes or to develop SAR models. Both library and substituent designs evolve as hits are encountered and the structures of target—inhibitor complexes are determined. Single modifications on both the template and substituents may be made during the process based on the structures of target—lead complexes. When sufficient SAR data has accumulated and the structures of target complexes with key templates and substituents have been determined, potent compounds with the desired “drug like” compositions are designed and synthesized using a structure-based recombination strategy (Figure 6). The integration of these recent advances in drug discovery offers the possibility to substantially decrease the time required to find initial leads and develop them into prototype drugs. Similarly, the interval required to produce preclinical development compounds and backup candidates is also expected to shrink. Perhaps most interesting is the potential of this integrated technology, which is ultimately based on an abstract information model of the biological process, to dramatically increase the reliability of successful drug development

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by factoring in and refining important drug properties concurrently with the optimization of drug affinity and specificity for a specific molecular receptor. This objective is achieved by integrating assays and computational models that relate to important drug development issues (e.g., oral absorption, optimal pharmacokinetics, minimal toxicity, etc.) directly into the iterative design process. V. Chemi-Informatics The development of methods that can optimize the parallel refinement of drug potency and pharmacological properties is a key objective in enhancing the efficiency and productivity of drug discovery. To address this problem and to handle the dramatic increase in experimental data generated using robotic synthesis and assay methodologies, advanced informatics systems are required to collect and exploit data relating to the properties of the chemicals being produced. These systems will amplify the synergy between structure-based design and combinatorial chemistry and provide a means for reducing the aggregate failure rate of development candidates. The poor success rate for preclinical development candidates (about 1 in 20) results from our limited ability to predict such drug properties as intestinal absorption, excretion, metabolism, toxicity, efficacy, and side effects. Rendering the prediction more difficult is the certainty that these drug properties are composite properties that result from the operation of many biological processes. Recently, progress has been made towards understanding the underlying molecular basis for some of the individual components that contribute to the observed drug properties. For instance, the absorption of compounds into Caco-2 cells in culture may be predictive of intestinal absorption [29–31]. Retention times of compounds during artificial membrane chromatography has also been correlated with oral absorption [32]. Recently, molecular transporters have been identified, cloned, and characterized that are responsible for the absorption of dipeptides and the excretion of organic cations [33,34]. The enzymatic basis for the metabolism of xenobiotics (cytochrome P450, glutathione transferase, etc.) has been known for some time, at least in part. In fact, many of the individual components of the composite biological processes can be developed into high-throughput assays, providing the opportunity to collect SAR data and develop SAR models. These observations, together with the integrated structure-based design and combinatorial chemical technology described here, define a comprehensive new strategy for drug discovery that has the potential to reduce the aggregate failure rate for development candidates (Figure 4). The creation of virtual libraries of compounds that are readily accessible through the use of automated

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synthesis methods allows for the first time an extensive and systematic investigation of structure-activity relationships related to drug properties. Small sublibraries can be selected, synthesized, and screened in high-throughput assays with the goal of developing predictive SAR models for individual components of the biological processes responsible for pharmacokinetic and toxicological properties of drug candidates. Eventually, it will be possible to assemble the predictive SAR models for various component processes to produce predictive models for bioavailability and toxicology. At this point it will be possible to use these models to guide the selection of sublibraries directed against therapeutic targets so that library members with the most “drug like” properties and the fewest liabilities are chosen. Steering away from compounds with undesired properties will focus the selection process on molecules with an improved probability of successful development. This strategy permits the simultaneous refinement of multiple chemical properties in addition to target efficacy and will shorten the drug-discovery process. Developing a system capable of collecting multivariate SAR data and exploiting the data to produce predictive SAR models is a major systems integration task. However, recent advances in computers, operating systems, and computational chemical tools now enables the implementation of a system that can track compounds, store chemical property data in a comprehensive relational database, and operate on virtual libraries in an iterative fashion to develop SAR models and refine chemical properties [28]. Tools for the production of virtual libraries have been developed by several groups and large virtual libraries can be produced within a few days using high-power computer workstations. A variety of tools also exist for selecting compounds based on criteria such as the ability to fit a target receptor [23], similarity or diversity [26,27], or any number of other properties that might be important for interaction with the target receptor. Statistical programs also exist that are up to the task of developing SAR models [20]. To obtain SAR models that have utility in the drug-discovery process it will be necessary to collect data on the properties of large numbers of compounds so that the models are predictive across diverse chemical classes of lead molecules. This requires implementation of a comprehensive relational database to collect and correlate SAR data. The challenge is to integrate these tools into a system that can collect and operate on vast arrays of chemical-property data in an intelligent fashion to direct and refine the properties of robotically synthesizable compounds. One approach that has been used successfully is to create a hierarchical clientserver system with a powerful central selection algorithm (Figure 4) that chooses compounds from virtual libraries based on their computed properties, supervises data analysis, and integrates results from experimental measurements [28,35]. This system, which has been termed DirectedDiversity in the author's laborato-

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ries, holds the promise of a major reduction in the time required to produce new preclinical compounds with enhanced development potential. VI. Conclusion The preceding outlines a new drug-discovery paradigm that integrates structure-based design, directed strategies for combinatorial chemical synthesis, and a comprehensive chemi-informatics system for accumulating and analyzing information regarding chemical properties. Three-dimensional structures provide the information required to most efficiently direct the design and optimization of new lead compounds. High-throughput automated methods of chemical synthesis produce new classes of lead compounds and provide for the rapid generation of structure—activity data. Chemical informatics systems track chemical compounds, store chemical property data, develop predictive SAR models, and provide a means for intelligently directing the drug-discovery process. By applying this approach not only to the therapeutic target, but also to molecules involved in absorption, clearance, metabolism, or toxicology it will be possible to develop predictive models for bioavailability and toxicology. Ultimately this approach will greatly increase the cost effectiveness and efficiency of drug discovery by reducing the aggregate failure rate for development candidates. References 1. Appelt K, Bacquet RJ, Bartlett CA, Booth CL, Freer ST, Fuhry MAM, Gehring MR, Herrmann SM, Howland EF, Janson CA, Jones TR, Ka C-C, Kathardekar V, Lewis KK, Marzoni GP, Matthews DA, Mohr C, Moomaw EW, Morse CA, Oatley SJ, Ogden RC, Reddy MR, Reich SH, Schoettlin WS, Smith WW, Varney MD, Villafranca JE, Ward RW, Webber S, Welsh KM, White J. Design of enzyme inhibitors using iterative protein crystallographic analysis. J Med Chem 1991; 34:1925–1934. 2. Ealick SE, Babu YS, Bugg CE, Erion MD, Guida WC, Montgomery JA, Secrist III JA. Application of crystallographic and modeling methods in the design of purine nucleoside phosphorylase inhibitors. Proc Natl Acad Sci USA 1991; 88:11540–11544. 3. Varney MD, Marzoni GP, Palmer CL, Deal JG, Webber S, Welsh KM, Bacquet RJ, Bartlett CA, Morse CA, Booth CLJ, Herrmann SM, Howland EF, Ward RW, White J. Crystal-structure-based design of Benz[cd]indole-containing inhibitors of thymidylate synthase. J Med Chem 1992; 35:663–676. 4. Reich SH, Fuhry MAM, Nguyen D, Pino MJ, Welsh KM, Webber S, Janson CA, Jordan SR, Matthews DA, Smith WA, Bartlett CA, Booth CLJ, Herrmann SM, Howland EF, Morse CA, Ward RW, White J. Design and synthesis of novel 6,7-

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Imidazotetrahydroquinoline inhibitors of thymidylate synthase using iterative protein crystal structure analysis. J Med Chem 1992; 35:847–858. 5. Weber PC, Wendoloski JJ, Pantoliano MW, Salemme FR. Crystallographic and thermodynamic comparison of natural and synthetic ligands bound to streptavidin. J Amer Chem Soc 1992; 114:3197–3200. 6. Weber PC, Pantoliano MW, Simons DM, Salemme FR. Structure-based design of synthetic azobenzene ligands for streptavidin. J Amer Chem Soc 1994; 116:2717–2724. 7. Andrews PR, Craik DJ, Martin JL. Functional group contributions to drug-receptor interactions. J Med Chem 1984; 27:1648–1657. 8. Terrett NK, Gardner M, Gordon DW, Kobylecki RJ, Steele J. Combinatorial synthesis—the design of compound libraries and their application to drug discovery. Tetrahedron 1995; 51:8135–8173. 9. Baum RM. Combinatorial approaches provide fresh leads for medicinal chemistry. C and EN 1994; 7:20–26. 10. Gallop MA, Barrett RW, Dower WJ, Fodor SPA, Gordon EM. Applications of combinatorial technologies to drug discovery. 1. Background and peptide combinatorial libraries. J Med Chem 1994; 37:1234–1251. 11. Gordon EM, Barrett RW, Dower WJ, Fodor SPA, Gallop MA. Applications of combinatorial technologies to drug discovery. 2. Combinatorial organic synthesis, library screening strategies and future directions. J Med Chem 1994; 37:1385–1399. 12. Dooley CT, Chung NN, Wolkes BC, Schiller PW, Bidlack JM, Pasternak GW, Houghten RA. An all D-amino acid peptide with central analgesic activity from a combinatorial library. Science 1994; 266:2019–2022. 13. Kerr JM, Banville SC, Zuckermann RN. Encoded combinatorial peptide libraries containing nonnatural amino acids. J Amer Chem Soc 1993; 115:2529–2531. 14. Ohlmeyer MHJ, Swanson RN, Dillard LW, Reader JC, Asouline G, Kobayashi R, Wigler M, Still C. Complex synthetic chemical libraries indexed with molecular tags. Proc Natl Acad Sci USA 1993; 90:10922–10926. 15. Dankwardt SM, Newman SR, Krestenansky JL. Solid phase synthesis of aryl and benzylpiperidines and their application in combinatorial chemistry. Tetrahedron Letters 1995; 36:4923–4926. 16. Baldwin JJ, Burbaum JJ, Henderson I, Ohlmeyer MHJ. Synthesis of a small molecule combinatorial library encoded with molecular tags. J Amer Chem Soc 1995; 117:5588–5589.

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17. Zuckermann RN, Martin EJ, Spellmeyer DC, Stauber GB, Shoemaker KR, Kerr JM, Figliozzi GM, Goff DA, Siani MA, Simon RJ, Banville SC, Brown EG, Wang L, Richter LS, Moos WH. Discovery of nanomolar ligands for 7-transmembrane G-protein coupled receptors from a diverse N(substituted)glycine peptoid library. J Med Chem 1994; 37:2678–2685. 18. Burbaum JJ, Ohlmeyer MHJ, Reader JC, Henderson I, Dillard LW, Li G, Randle TL, Sigal NH, Chelsky D, Baldwin JJ. A paradigm for drug discovery employing encoded combinatorial libraries. Proc Natl Acad Sci USA 1995; 92:6027–6031. 19. Lam KS, Salmon SE, Hersh EM, Hruby VJ, Kazmierski WM, Knapp RJ. A new type of synthetic peptide library for identifying ligand-binding activity. Nature 1991; 354:82–84. 20. Loew GH, Villar HO, Alkorta I. Strategies for indirect computer-aided drug design. Pharmaceutical Res 1993; 10:475–486.

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21. Kick EK, Ellman JA. Expedient method for the solid-phase synthesis of aspartic acid protease inhibitors directed towards the generation of libraries. J Med Chem 1995; 38:1427–1430. 22. Campbell DA, Bermak JC, Burkoth TS, Patel DV. A transition state analog inhibitor combinatorial library. J Amer Chem Soc 1995; 117:5381–5382. 23. Kuntz ID, Meng EC, Shoichet BK. Structure-based molecular design. Acc Chem Res 1994; 27:117–123. 24. Bohm H-J. LUDI: Rule-based automatic design of new substituents for enzyme inhibitor leads. Journal of Computer-Aided Molecular Design 1992; 6:593–606. 25. Bohm H-J. The computer program LUDI: A new method for the de novo design of enzyme inhibitors. Journal of Computer-Aided Molecular Design 1992; 6:61–78. 26. Martin EJ, Blaney JM, Siani M, Spellmeyer DC, Wong AK, Moos WH. Measuring diversity: experimental design of combinatorial libraries for drug discovery. J Med Chem 1995; 38:1431–1436. 27. Sullivan M. Mass Screening: a new approach to chemical discovery. Today's Chemist at Work 1994;September:19–26. 28. Agrafiotis DK, Bone RF, Salemme FR, Soll RM. A system and method of automatically generating chemical compounds with desired properties. US Patent 5463564, October 31, 1995. 29. Hildalgo IJ, Raub TJ, Borchardt RT. Characterization of the human colon carcinoma cell line (Caco2) as a model system for intestinal epithelial permeability. Gastroenterology 1989; 96:736–749. 30. Gan L-S, Eads C, Niederer T, Bridgers A, Yanni S, Hsyu P-H, Pritchard FJ, Thakker D. Use of Caco2 cells as an in vitro intestinal absorption and metabolism model. Drug Development and Industrial Pharmacy 1994; 20:615–631. 31. Conradi RA, Hilgers AR, Ho NFH, Burton PS. The influence of peptide structure on transport across Caco-2 cells. Pharmaceutical Research 1991; 8:1453–1460. 32. Pidgeon C, Ong S, Liu H, Qiu X, Pidgeon M, Dantzig AH, Munroe J, Hornback WJ, Kasher JS, Glunx L, Szczerba T. IAM chromatography: an in vitro screen for predicting drug membrane permeability. J Med Chem 1995; 38:590–594. 33. Grundermann D, Gorboulev V, Gambaryan S, Veyhl M, Koepsell H. Drug excretion mediated by a new prototype of polyspecific transporter. Nature 1994; 372:549–552.

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34. Fei Y-J, Kanai Y, Nussberger S, Ganapathy V, Leibach FH, Romero MF, Singh SK, Boron WF, Hediger MA. Expression and cloning of a mammalian proton-coupled oligopeptide transporter. Nature 1994; 368:563–566. 35. Agrafiotis DK, Bone R, Jaeger EP, Rhind AW, Salemme FR, Soll RM. Directed Diversity®: a new paradigm for drug discovery. 1996; in preparation.

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21 Structure-Based Combinatorial Ligand Design Amedeo Caflisch University of Zürich, Zürich, Switzerland Claus Ehrhardt Novartis Pharma Inc., *Basel, Switzerland I. Introduction Structure-based ligand design is fascinating and challenging. Whenever it is possible to determine the three-dimensional structure of a pharmacologically relevant enzyme or receptor, computational approaches can be used to design novel high-affinity ligands. These methods can complement the broad screening efforts, which represent traditional lead discovery. In this chapter we focus on our approach to computer-aided ligand design. It is based on the docking of a diverse set of molecular fragments into the active site of a macromolecular target and on the use of a combinatorial strategy to connect them to form candidate ligands. The methodology is illustrated by an application to human thrombin, a trypsin-like serine protease fulfilling a central role in both hemostasis and thrombosis. The selective inhibition of thrombin is expected to prevent thrombotic diseases. Ligand-design programs are being developed at an ever increasing rate and some are related to various aspects of our ligand design approach. The LEGO software tool is based on the combination of multiple fragment docking, automatic connection by small linker units (one to four atom chains), and searching of three-dimensional databases for complementary molecules [1,2]. It has been implemented within the MOLOC molecular modeling system [3], which allows the visualization of the functionality maps and interactive model building of the growing ligands. Another related approach is that embodied in the program LUDI [4–8]. It makes use of statistical data from small-molecule * Formerly Sandoz Pharma Ltd.

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crystal structures to determine binding sites of molecular fragments, i.e., discrete positions on the binding site surface suitable to form hydrogen bonds and/or to fill hydrophobic sites of the receptor. Alternatively, it uses simple rules or the output of the program GRID [9–12] to generate the interaction sites. Finally, the fragments fitted in the interaction sites are connected by linker groups. Other fragmentbased programs are GROUPBUILD [13]; GROW [14], HOOK [15], NEWLEAD [16], SPROUT [17], and TORSION [18]. These and other strategies for computer-aided structure-based ligand design have been reviewed by several contributors [13,19,20]. II. Docking Molecular Fragments A. Multiple Copy Simultaneous Search The present approach for ligand design is based on the combinatorial selection of molecular fragments optimally docked on the protein binding site to form a population of diverse candidate ligands. The multiple copy simultaneous search (MCSS) procedure combines the advantages of random distribution and simultaneous minimization of a set of replicas of a chemical fragment to obtain maps of energetically favorable positions and orientations (local energy minima) [21,22]. These maps, which contain all possible low-energy minima of a fragment-protein complex, are called functionality maps. A plethora of structural and thermodynamic data on inhibitor-enzyme complexes [23–26] suggest that the burial of nonpolar groups of the ligand in hydrophobic pockets of the protein is important for binding affinity and that intermolecular electrostatic interactions determine selectivity. For this reason, and because most of the known enzymes' binding sites have both hydrophilic and hydrophobic character, very diverse functional groups are used in MCSS. Representative examples include charged (e.g., acetate, benzamidine, methylammonium, methylguanidinium, pyrrolidine); polar (e.g., methanol, 2propanone, N-methylacetamide); aromatic (e.g., benzene, pyrrole, imidazole, phenol); and aliphatic (e.g., propane, isobutane, cyclopentane, cyclohexane) groups. Although most of these fragments are rigid, MCSS can also generate the functionality maps of flexible medium-size fragments, e.g., the amino acid side chains. Additional functional groups and more complex heterocyclic systems are currently being introduced to increase the diversity of the resulting ligands and to better characterize the specificity of the binding pockets (A. Caflisch and C. Ehrhardt, unpublished results). As shown by a flowchart in Figure 1, the method is fully automated, although certain critical parameters (e.g., number of replicas, radius of the sphere for random distribution, CHARMM parameters for the minimization) can be adjusted by the user to optimize it for specific applications. Several thousand replicas of a given group are randomly distributed inside a sphere

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Figure 1 Schematic representation of the MCSS procedure. Conditional statements are enclosed by diamonds.

whose radius is chosen large enough to cover the entire region of interest. This can be a known binding site or the entire protein, if one wants to explore alternative binding pockets. The initial random distribution also can be performed inside a parallelepiped if the region of interest is elongated in one or two directions. A minimal distance can be given as input to avoid bad contacts between functional group atoms and protein atoms for the initial distribution. Subsets of between 500 and 3000 randomly distributed replicas of the same group are simultaneously minimized in the force field of the protein. The

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interactions between the group replicas are omitted. The polar-hydrogen approximation (PARAM19) of the CHARMM force field is used [27]. In the application of the MCSS method to the sialic acid binding site of the influenza coat protein hemagglutinin [21], HIV-1 aspartic proteinase [22], and thrombin [20,28] the protein was kept fixed; hence, the forces on each replica consist of its internal forces and those due to the protein, which has unique conformation and, therefore, generates a unique field. The positions are compared every 1000 steps to eliminate replicas converging toward a common minimum. Further details concerning the methodology are given in References 21 and 22, while a critical assessment has been presented in Reference 20. B. Simple Approximations of Solvation Effects In previous applications of MCSS [21,22] the effects of the solvent were neglected, i.e., all proteinfragment interactions were calculated with a vacuum potential [27]. This choice was based on the principle that fast methods are necessary to perform effective searches of the binding site and that good candidate ligands subsequently can be ranked in terms of their binding free energy [20,28]. A possible difficulty with this approach is that minimized positions may be missed or misplaced due to the lack of a solvation correction during the MCSS minimization. Electrostatic Shielding In MCSS studies of thrombin [20,28], it was observed that minima of charged groups tend to cluster in the vicinity of charged side chains on the thrombin surface and in the S1 (basic groups) or S1' pocket (acidic groups). It is then necessary to estimate the electrostatic desolvation of both protein and fragment to obtain a realistic ranking of the minima [28]. As a simple test of the importance of electrostatic shielding, a distance-dependent dielectric function [29] was introduced instead of the unit dielectric constant in the vacuum potential. The overall shape of the acetate map did not change, but three more minima were found close to the Lys60F side chain in S1' [20]. It is difficult to find a physical meaning in favor of the distance-dependent dielectric function. Nevertheless, it is a simple and useful approximation, since it yields a smoother and more realistic potential surface than the vacuum Coulombic interaction. Hydrophobic Effect Aliphatic and aromatic fragments do not bear any partial charges in the polar-hydrogen approximation. Hence, MCSS determines their optimal position in the

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protein binding site exclusively by van der Waals interactions. Minima of nonpolar fragments may be found in hydrophilic pockets because of the lack of an energy penalty for protein desolvation. A representative example of a cluster of propane minima in a mainly hydrophilic region of the HIV-1 aspartic proteinase binding site is shown in Reference 20. In a simple attempt to approximate desolvation of polar regions of the protein, the attractive contribution of the van der Waals interaction energy was switched off between atoms of nonpolar MCSS fragments and all protein polar hydrogen, nitrogen, oxygen, and sp2 carbon atoms. In addition, the van der Waals radius of nitrogen and oxygen atoms was increased from the PARAM19 default value of 1.6 Å to 2.2 Å and the van der Waals radius of the aliphatic carbons was reduced by 0.1 Å to avoid the too large van der Waals distance between carbons often produced by PARAM19. The modified force field yields thrombin functionality maps of propane, cyclopentane, cyclohexane, and benzene in agreement with structural data of known inhibitors (see Section II. C). In addition, these nonpolar groups are prevented from occupying hydrophilic pockets. As a further test of the modified force field, the MCSS procedure was used to generate the propane functionality map on the surface of the A peptide chain of the leucine zipper, i.e., residues 249–281 from the yeast transcriptional activator protein GCN4. Leucine zippers are composed of amphipathic α helices containing heptad repeats (abcdefg) in which hydrophobic residues are frequent at a and d. As the x-ray structure indicates [30,31], the two amphipathic α helices are held together by hydrophobic interactions between residues in the a and d positions (Figure 2). The B peptide chain was removed and MCSS was run separately with the original and the modified force field starting from the same random distribution of 5000 propane replicas around helix A. The sixty most favorable minima obtained with the modified force field are distributed in twelve clusters, seven of which match the Val and Leu side chains of the B helix involved in the interhelical interactions (Figure 2). Of the remaining five clusters, labeled A to E in Figure 2, B has minima in contact with Val and Ala side chains, D with Leu and Tyr side chains, while A, C, and E with both a Val or Leu side chain and the alkyl part of Lys side chain. The sixty most favorable minima obtained with the original force field form sixteen clusters (not shown). Only four of these match Val and Leu side chains of the B helix, while eight clusters desolvate one (or more) polar group on the hydrophilic (exposed) surface of the A helix. The functionality maps of polar groups generated with a distance-dependent dielectric and those of nonpolar groups obtained with the modified force field are closer to those obtained by using a continuum dielectric model [32–35] to postprocess the MCSS minima and estimate solvation effects [28]. Hence, simple modifications of the force field may result in more accurate projections of the binding free energy. Since these modifications are used during the

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Figure 2 Stereo view of the sixty propane minima (thick lines) obtained with the modified force field (see text) on the surface of the A peptide chain (medium lines) of the GCN4 leucine zipper (PDB code 2ZTA). Although the B peptide chain was removed during the MCSS procedure, its backbone and hydrophobic side chains are also drawn (thin lines) to show how the propane minima match the aliphatic groups of chain B. Hydrophobic residues are labeled at their Cα atom.Five clusters of propane minima that do not match the hydrophobic side chain of the B helix involved in the interhelical interactions are labeled from A (top right) to E (bottom center) and discussed in the text.

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minimization phase, a more realistic distribution of functional group minima is generated. C. Thrombin Functionality Maps Human thrombin is one of the best characterized enzymes from a structural point of view (Figure 3). It binds a series of diverse inhibitors without major rearrangements in its conformation, as shown by a number of x-ray crystallography studies [26,36–39]. Its S3 and S2 precleavage subpockets have hydrophobic character, whereas at the bottom of the S1 or recognition pocket the carboxy group of Asp189 is a salt bridge partner for basic side chains (Figure 3). D-phenylalanyl-L-prolyl-L-arginine chloromethane, PPACK, (Figure 4a), and Nα-((2-naphthylsulfonyl)glycyl)-DL-pamidinophenylalanylpiperidine, NAPAP, (Figure 4b) are the archetypal active-site inhibitors of thrombin. The crystal structure of the thrombin-NAPAP complex is shown in Figure 3. The PPACK and NAPAP inhibitors bind to the thrombin active site by occupying the S3 and S2 pockets with their hydrophobic moieties and by positioning their basic group (guanidinium of PPACK, benzamidine of NAPAP) into S1 to form a salt bridge with Asp189. In continuation of a project aiming at the structure-based design of low molecular weight, active-site directed inhibitors of human thrombin [40], MCSS was used to generate a series of functionality maps of the thrombin S3 to S2' pockets [20,28]. A detailed description of the thrombin functionality maps and the continuum approximation used to postprocess the MCSS minima is given in Reference [28]. From the analysis of the results for the nonpolar groups it is evident that hydrophobic moieties prefer to bind to the S3 and S2 pockets (Figure 5). The solvent exposed face of the Trp60D indole is another favorable site, though the intermolecular van der Waals interactions are much smaller. Binding to the S2' region is favored by interactions with the Leu40 side chain but implies a desolvation penalty because of the burial of part of the Arg73 guanidinium and/or the Gln151 side chain. The latter might be an artifact of the rigid protein structure used in the minimization, since the side chains of Arg73 and Gln151 are flexible enough to displace their polar groups towards a more exposed region. Binding to the neighboring Leu41 side chain in S1' is highly unfavorable because of the concomitant desolvation of Lys60F. For polar groups with zero net charge there are several hydrophylic groups on the thrombin main chain that might be involved in strong hydrogen

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Figure 3 (a) Stereo view of the thrombin molecule in its complex with NAPAP. The side chains of thrombin involved in the binding of NAPAP and the disulfide bridges are shown in stick-and-ball representation with black sticks. NAPAP is shown in stick-and-ball representation with white sticks. (b) Zoom image of the active site region. Figures made with the program MOLSCRIPT [42].

bonds. These are 214CO, 216NH, 216CO, and 219NH in S1; 193NH and 195NH in the oxyanion hole; 41CO in S1'; 40CO in S2'; and 147NH and 148NH on the autolysis loop, whose exposure is dependent on crystallization conditions and inhibitor type. Two main conclusions can be drawn from the analysis of the minimized positions of the charged functional groups. First, the minima with the lowest binding free energy have optimal hydrogen bonds with the Asp189 side chain in the S1 pocket. Representative examples are the lowest energy minimum of benzamidine (Figure 5) and the lowest energy minima of methylguanidinium and methylammonium (Figure 4 of Reference 28). Since the Asp189 side chain http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_548.html (1 of 2) [4/9/2004 12:52:27 AM]

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Figure 4 Chemical structure of PPACK (a) and NAPAP (b).

is more buried than the side chain of Lys60F, the minima of positively charged groups interacting with the former have a more favorable binding free energy than those of the negatively charged groups close to the latter. This is due to reduced shielding of the charge—charge interaction and the smaller desolvation of the carboxylate oxygens of Asp189 compared to the amino group in Lys60F [28]. Second, polar groups on the protein surface are not ideal partners for a charged functional group because the high desolvation penalty for these groups might not be completely compensated for by the favorable electrostatic interaction energy. This finding is analogous to the results for the polar functional groups, i.e., their binding to a partially exposed charged side chain of the protein may result in an unfavorable total binding free energy.

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Figure 5 Stereo view of the three lowest energy minima of benzene obtained with the modified force field and the lowest energy minimum of benzamidine (thick lines for heavy atoms and thin lines for polar hydrogens) in the thrombin active site (thin lines). The inhibitor PPACK is also shown (medium lines), though it was removed during the MCSS procedure. Some Cα atoms of thrombin are labeled.

III. Connecting Molecular Fragments A. Computational Combinatorial Ligand Design Overview The recently developed program for computational combinatorial ligand design (CCLD) requires as input atomic coordinates and partial charges of the protein atoms, as well as the coordinates of the MCSS minima and the individual contributions to the free energy of binding [28]. An additional file contains a number of control parameters and, for each functional group used for MCSS, a list of atoms which can be used for connection (linkage atoms). The following procedures are performed during a regular execution of CCLD (Figure 6): The MCSS minima are first sorted according to their approximated binding free energies [28]; then a list of bonding fragment pairs and a list of overlapping fragment pairs are generated (see below). This is followed by the combinatorial generation of putative ligands (see below).

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Figure 6 Schematic representation of the CCLD program. Variable assignments are symbolized by :=. Conditional statements are enclosed by diamonds. [From Reference 28.]

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Lists of Bonding Fragment Pairs and Overlapping Fragment Pairs The user has to specify for each functional group type which atoms are to be used for connection to other fragments. For each linkage atom CCLD generates a set of possible linkage points, i.e., points that will be used to determine the position and orientation of the link. All possible pairs of minimized positions are then analyzed and added to the list of bonding fragment pairs if they can be linked; otherwise, if two fragments have bad contacts they are added to the list of overlapping fragment pairs. A pair of bonding fragments may be connected by a linker unit, by a single covalent bond (1-bond), or by fusing two overlapping atoms belonging to different fragments (0-bond). The linker units are small since their function is to optimally connect two fragments without adding considerably to the molecular weight. The following linker elements have been implemented so far: Keto and methylene (2-bond), amide and ethylene (3-bond). The user is free to choose minimal and maximal values for the distance (d) between linkage atoms for each connection type. In the application to thrombin the following values in Ångstroms were used: d<0.43, 0-bond; 1.2< d<1.8, 1-bond; 2.2
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B. Candidate Ligands of Thrombin To test if CCLD is able to reproduce the known thrombin inhibitors, the functional groups of PPACK and NAPAP were given as input molecular fragments in the orientation derived from the crystal structures of the complexes. The CCLD program generated a set of candidate ligands that not only contained the PPACK and NAPAP structures but also a number of interesting hybrid molecules consisting of fragments from both inhibitors. A representative example is shown in Figure 7. This putative ligand consists of the C-terminal part of NAPAP, whose piperidine ring is connected at the 3position to the PPACK D-Phe by an amide linker. The latter has its carbonyl oxygen involved in a hydrogen bond with the Gly216 NH. In another run, the MCSS minima of benzamidine (Figure 5), benzene (Figure 5), cyclopentane, and cyclohexane [28] were used as starting molecular fragments. In a few seconds of CPU time of an SGI Indigo2 (R4400 processor), CCLD generated a series of molecules showing the same interaction patterns as those of known thrombin inhibitors, i.e., hydrophobic moieties in S3 and S2, hydrogen bonds with the polar groups of Gly216, and benzamidine in S1. One of these putative ligands is shown in Figures 8 and 9. It is involved in the same interactions as in the NAPAP-thrombin complex except for the hydrogen bond with the CO of Gly216. Its cyclohexane ring in S3 is connected to the

Figure 7 PPACK-NAPAP hybrid ligand (thick lines) generated by CCLD starting from the functional groups of PPACK and NAPAP (thin lines). The amide linker connecting the piperidine in 3-position to the D-Phe was created by CCLD.

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Figure 8 Minimized structure of a putative ligand suggested by CCLD (thick lines). The CCLD run used MCSS minima of benzamidine, benzene, cyclopentane, and cyclohexane. The putative ligand was minimized in the thrombin active site, whose residues within 8 Å of any atom of the ligand were allowed to move. The remaining residues of thrombin were kept rigid. The NAPAP structure is also shown (thin lines) as a basis of comparison.

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Figure 9 Stereo view of the minimized complex between the putative ligand shown in Figure 8 and thrombin (thin lines). Intermolecular hydrogen bonds are drawn by dashed lines.

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N-acylpyrrolidine ring in S2 by a single methylene linker. This is a novel design and the candidate ligand appears to be more rigid than NAPAP, since it has a smaller number of rotatable bonds. Hence, the penalty paid for the loss in entropy upon binding should be smaller for this CCLD hit than for NAPAP. In a study with a more diverse set of starting molecular fragments, ligands containing both a “core” similar to known inhibitors and additional intermolecular hydrogen bonds and/or van der Waals interactions were generated [28]. IV. Conclusion A combinatorial approach for the computer-aided design of putative ligands of proteins or receptors of known three-dimensional structure has been presented. Diversity of these candidate ligands is provided by first docking a set of diverse molecular fragments. For aliphatic functional groups a modified force field (switching off the attractive part of the van der Waals interaction with polar atoms of the protein) was introduced to better approximate solvation effects, thereby avoiding the docking of apolar fragments into hydrophilic cavities of the macromolecular target. The second part of the present ligand design approach consists of a combinatorial strategy for the connection of optimally docked molecular fragments by small linkage elements having optimal interactions with the target molecule. An application was presented in which candidate inhibitors of thrombin were found. It is important to note that the most difficult part of ligand design is the prediction of binding affinity. A method to estimate relative binding constants has recently been applied to a series of similar inhibitors of HIV-1 aspartic proteinase [41]. Our own efforts are concentrated on the development of an approach that will predict relative binding affinities and will be general enough to be used for any enzyme or receptor of known structure (A. Caflisch, D. Arosio, and C. Ehrhardt, work in progress). One of the purposes of this chapter was to show that structure-based ligand design is a fascinating and progressing research field. It is fascinating not only for its ultimate goal, i.e., the discovery of ethical drugs, but also because it is based on, and thereby increases, our understanding of molecular interactions and recognition phenomena on an atomic level. Another reason is its multidisciplinary character, which requires skills in different branches of science, from theoretical physics and chemistry to computer science and statistics. That structure-based computer-aided ligand design is a progressing field is evident in many chapters of this book. This is mainly a consequence of the methodological developments and the ever-increasing performance of computers.

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Acknowledgments We thank J. Apostolakis and Professor A. Plückthun for helpful discussions. The calculations were performed on an SGI Indigo2 and an eight-processor SGI Challenge (R4400 processors). The CHARMM program within the version 4.0 of the QUANTA software package (Biosym-MSI Inc) was used for some of the minimization performed in this work. The CCLD program is available from A. Caflisch. This work was supported by the Swiss National Science Foundation (Schweizerischer Nationalfonds grant nr. 3100-043423.95) and by Novartis Pharma Inc. References 1. Gubernator K, Broger C, Bur D, Doran DM, Gerber PR, Müller K, Schaumann TM. Structure-based ligand design. In: Hermann EC, Frankle R, eds. Computer-Aided Drug Design in Industrial Research 1995:61–77. 2. Müller K. Paradigms of rational molecular design. In: Schwartz TW, Hjorth SA, Kastrup JS, eds. Structure and Function of 7TM Receptors, in press. 3. Gerber PR, Gubernator K, Müller K. Generic shapes for the conformational analysis of macrocyclic structures. Helv Chim Acta 1988; 71:1429–1441. 4. Böhm HJ. The computer program LUDI: a new method for de novo design of enzyme inhibitors. J Comput-Aided Mol Design 1992; 6:61–78. 5. Böhm HJ. LUDI: rule-based automatic design of new substituents for enzyme inhibitor leads. J Comput-Aided Mol Design 1992; 6:593–606. 6. Böhm HJ. The development of a simple empirical scoring function to estimate the binding constant for a protein-ligand complex of known three-dimensional structure. J Comput-Aided Mol Design 1994; 8:243–256. 7. Böhm HJ. On the use of LUDI to search the fine chemicals directory for ligands of proteins of known three-dimensional structure. J Comput-Aided Mol Design 1994; 8:623–632. 8. Böhm HJ. Site-directed structure generation by fragment-joining. Perspectives in Drug Discovery and Design 1995; 3:21–33. 9. Goodford PJ. A computational procedure for determining energetically favorable binding sites on biologically important macromolecules. J Med Chem 1985; 28:849–857.

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10. Bobbyer DNA, Goodford PJ, McWhinnie PM, Wade RC. New hydrogen-bond potentials for use in determining energetically favorable binding sites on molecules of known structure. J Med Chem 1989; 32:1083–1094. 11. Wade RC, Clark KJ, Goodford PJ. Further development of hydrogen bond functions for use in determining energetically favorable binding sites on molecules of known structure. 1. Ligand probe groups with the ability to form two hydrogen bonds. J Med Chem 1993; 36:140–147.

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12. Wade RC, Goodford PJ. Further development of hydrogen bond functions for use in determining energetically favorable binding sites on molecules of known

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structure. 2. Ligand probe groups with the ability to form more than two hydrogen bonds. J Med Chem 1993; 36:148–156. 13. Rotstein SH, Murcko MA. GroupBuild: A fragment-based method for de novo drug design. J Med Chem 1993; 36:1700–1710. 14. Moon JB, Howe WJ. Computer design of bioactive molecules: a method for receptor-based de novo ligand design. Proteins: Structure, Function and Genetics 1991; 11:314–328. 15. Eisen MB, Wiley DC, Karplus M, Hubbard RE. HOOK: A program for finding novel molecular architectures that satisfy the chemical and steric requirements of a macromolecule binding site. Proteins: Structure, Function and Genetics 1994; 19:199–221. 16. Tschinke V, Cohen NC. The NEWLEAD program: a new method for the design of candidate structures from pharmacophoric hypotheses. J Med Chem 1993; 14:3863–3870. 17. Gillet VJ, Myatt G, Zsoldos Z, Johnson AP. SPROUT, HIPPO and CAESA: Tools for de novo structure generation and estimation of synthetic accessibility. Perspectives in Drug Discovery and Design 1995; 3:34–50. 18. Lewis RA, Roe DC, Huang C, Ferrin TE, Langridge R, Kuntz ID. Automated site-directed drug design using molecular lattices. J Mol Graphics 1992; 10:66–78. 19. Kuntz ID. Structure-based strategies for drug design and discovery. Science 1992; 257:1078–1082. 20. Caflisch A, Karplus M. Computational combinatorial chemistry for de novo ligand design: Review and assessment. Perspectives in Drug Discovery and Design 1995; 3:51–84. 21. Miranker A, Karplus M. Functionality maps of binding sites: a multiple copy simultaneous search method. Proteins: Structure, Function, and Genetics 1991; 11:29–34. 22. Caflisch A, Miranker A, Karplus M. Multiple copy simultaneous search and construction of ligands in binding sites: application to inhibitors of HIV-1 aspartic proteinase. J Med Chem 1993; 36:2142–2167. 23. Appelt K. Crystal structures of HIV-1 protease-inhibitor complexes. Perspectives in Drug Discovery and Design 1993; 1:23–48. 24. Wlodawer A, Erickson JW. Structure-based inhibitors of HIV-1 protease. Annu Rev Biochem 1993; 62:543–585. 25. Stubbs MT, Bode W. Crystal structures of thrombin and thrombin complexes as a framework for antithrombotic drug design. Perspectives in Drug Discovery and Design 1993; 1:431–452.

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26. Hilpert K, Ackermann J, Banner DW, Gast A, Gubernator K, Hadvary P, Labler L, Müller K, Schmid G, Tschopp T, van de Waterbeemd H. Design and synthesis of potent and highly selective thrombin inhibitors. J Med Chem 1994; 37:3889–3901. 27. Brooks BR, Bruccoleri RE, Olafson BD, States DJ, Swaminathan S, Karplus M. CHARMM: A program for macromolecular energy, minimization, and dynamics calculations. J Comput Chem 1983; 4:187–217. 28. Caflisch A. Computational combinatorial ligand design: Application to human α-thrombin. J Computer-Aided Molec Design 1996; 10:372–396. 29. Gelin BR, Karplus M. Sidechain torsional potentials and motion of amino acids in proteins: bovine pancreatic trypsin inhibitor. Proc Natl Acad Sci USA 1975; 72:2002–2006.

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30. O'Shea EK, Rutkowski R, Kim PS. Evidence that the leucine zipper is a coiled coil. Science 1989; 243:538–542. 31. O'Shea EK, Klemm JD, Kim PS, Alber T. X-ray structure of the GCN4 leucine zipper, a twostranded, parallel coiled coil. Science 1991; 254:539–544. 32. Warwicker J, Watson HC. Calculation of the electric potential in the active site cleft due to α-helix dipoles. J Mol Biol 1982; 157:671–679. 33. Gilson MK, Honig BH. Calculation of the total electrostatic energy of a macromolecular system: solvation energies, binding energies, and conformational analysis. Proteins: Structure, Function, and Genetics 1988; 4:7–18. 34. Bashford D, Karplus M. pKas of ionizable groups in proteins: atomic detail from a continuum electrostatic model. Biochemistry 1990; 29:10219–10225. 35. Davis ME, Madura JD, Luty BA, McCammon JA. Electrostatics and diffusion of molecules in solution: simulations with the University of Houston Brownian dynamics program. Comp Phys Comm 1991; 62:187–197. 36. Bode W, Mayr I, Baumann U, Huber R, Stone SR, Hofsteenge J. The refined 1.9-Å crystal structure of human α-thrombin: interaction with D-Phe-Pro-Arg chloromethylketone and significance of the TyrPro-Pro-Trp insertion segment. EMBO J 1989; 8:3467–3475. 37. Banner DW, Hadvary P. Crystallographic analysis at 3.0-Å resolution of the binding to human thrombin of four active site-directed inhibitors. J Biol Chem 1991; 266:20085–20093. 38. Obst U, Gramlich V, Diederich F, Weber L, Banner DW. Design neuartiger, nichtpeptidischer Thrombin-Inhibitoren und Struktur eines Thrombin-Inhibitor-Komplexes. Angew Chem 1995; 107:1874–1877. 39. Håkansson K, Tulinsky A, Abelman MM, Miller TA, Vlasuk GP, Bergum PW, Lim-Wilby MSL, Brunck TK. Crystallographic structure of a peptidyl keto acid inhibitor and human α-thrombin. Bioorganic and Medicinal Chemistry 1995; 3:1009–1017. 40. Tapparelli C, Metternich R, Ehrhardt C, Cook NS. Synthetic low-molecular weight thrombin inhibitors: molecular design and pharmacological profile. TIPS 1993; 14:366–376. 41. Holloway MK, Wai JM, Halgren TA, Fitzgerald PMD, Vacca JP, Dorsey BD, Levi 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 for HIV-1 protease inhibitors employing energy minimization in the active site. J Med Chem 1995; 38:305–317.

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42. Kraulis P, Molscript, a program to produce both detailed and schematic plots of protein structures. J Appl Crystallogr 1991; 24:946–950.

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22 Peptidomimetic and Nonpeptide Drug Discovery: Impact of StructureBased Drug Design Tomi K. Sawyer Parke-Davis Pharmaceutical Research, Ann Arbor, Michigan I. Introduction Peptidomimetic and nonpeptide drug discovery has evolved to become an extraordinarily intriguing area of interdisciplinary research. It challenges synthetic, computational, and biophysical chemists, biochemists, pharmacologists, and drug delivery scientists to collaboratively discover lead compounds that exhibit sufficient potency, selectivity, metabolic stability, and in vivo pharmacological efficacy to warrant further development as drug candidates. Over the past two decades a highly focused effort in both industry and academia has advanced the rational transformation of “first generation” peptide lead compounds to significantly modified analogs having minimal peptide-like chemical structure [1–18]. Such work is typically illustrated by systematic backbone and/or side chain modifications, transformation into macrocycles, structure-conformation analysis (x-ray, NMR, CD), and computerassisted molecular modeling. These extensive structure-activity and structure-conformation studies have enabled the creation of prototype peptidomimetic “second-generation” analogs from initial peptide leads. Over recent years the emergence of 3D structural information on target proteins has significantly impacted peptidomimetic drug discovery strategies. Particularly noteworthy has been the advancement of the de novo design of chemically novel compounds that possess very limited peptide-like substructure, but include structure-based functional group modifications which may make new intermolecular interactions with the target protein (versus a “first generation” peptide lead compound). Furthermore, the integration of such structure-based drug design efforts with high-throughput

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screening and synthetic chemical library generation is markedly reshaping peptide, peptidomimetic, and nonpeptide drug discovery research. In this chapter a few examples of peptidomimetic and nonpeptide drug discovery are detailed to highlight the scope of such work relative to a few specific targets (e.g., receptors, proteases, and signal transduction proteins) in which structure-based drug design has contributed in significant ways. A. Peptides: Molecular Diversity and φ-Ψ-χ Space Peptides exhibit extraordinary molecular diversity by virtue of their varying primary structures (Figure 1). For many peptide hormones, neurotransmitters, and releasing factors the substructure of amino acids that contribute to molecular recognition (binding) and biological activity (signal transduction) at their target receptors have been determined [2]. Such work has led to the generation of pharmacophore models of either agonist or antagonist analogs and, in some cases, the design of peptidomimetics. Yet, for most peptide growth factors, cytokines, and large-sized (>50 amino acids) peptide hormones, the identification of the amino acid substructure which accounts for molecular recognition and signal transduction has been a difficult task, and proposals for pharmacophore models remain significant challenges. In this regard the term “pharmacophore” is defined as the collection of relevant groups (substructure) of a ligand which are arranged in three-dimensional space in a manner complementary to the target protein and are responsible for the biological property of the ligand as a result of binding of the ligand to its target protein [8b]. The three-dimensional structural properties of peptides (Figure 2) are defined in terms of torsion angles (Ψ, φ, ω, χ) between the backbone amine nitrogen (Nα), backbone carbonyl carbon (C'), backbone methine carbon (Cα), and side chain hydrocarbon functionalization (eg., Cβ, Cγ, Cδ, Cε of Lys) derived from the amino acid sequence. A Ramachandran plot (Ψ versus φ) may define the preferred combinations of torsion angles for ordered secondary structures (conformations), such as α helix, β turn, γ turn, or β sheet. With respect to the amide bond torsion angle (ω) the trans geometry is more energetically favored for most typical dipeptide substructures, however, when the C-terminal partner is Pro or other N-alkylated (including cyclic) amino acids the cis geometry is possible and may further stabilize β-turn or γ-turn conformations. Molecular flexibility is directly related to covalent and/or noncovalent bonding interactions within a particular peptide. Even modest chemical modifications by Nα-methyl, Cα-methyl or Cβ-methyl can have significant consequences on the resultant conformation [6; also, see Phe analogs in Figure 2]. The Nα-Cα-C' scaffold may be further transformed by introduction of olefin substitution (e.g., CαCβ rarrow.gif C=C or dehydroamino acid [19]) or insertion (e.g., Cα-C' rarrow.gif Cα-C=C-C' or vinylogous amino acid [20]). Also the Cβ carbon may be substituted to advance the design of so-called “chimeric” amino acids

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Figure 1 Examples of native peptide hormones, neurotransmitters, and releasing factors

[9]. Finally, with respect to N-substituted amides it is also noteworthy to mention the intriguing approach [21] of replacing the traditional peptide scaffold by achiral N-substituted glycine building blocks. Overall, such Nα-Cα-C scaffold or Cα-Cβ side chain modifications provide significant opportunities for expanding peptide-based molecular diversity (i.e., so-called “peptoid” libraries) as well as to extend our 3D structural knowledge of traditional φ-Ψ-χ space.

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Figure 2 Three-dimensional structural properties of peptides: backbone and side chain

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B. Peptidomimetic Drugs: Concepts, Strategies, and Technologies Historically, the major focus of peptidomimetic design has evolved from receptor-targeted drug discovery research and has not been directly impacted by an experimentally-determined 3D structure of the target protein. Nevertheless, a hierarchial approach of peptide rarrow.gif peptidomimetic molecular design and chemical modification has evolved over the past two decades, based on systematic transformation of a peptide ligand and iterative analysis of the structure-activity and structureconformation relationships of “second generation” analogs (Figure 3). Such work has typically integrated biophysical techniques (x-ray crystallography and/or NMR spectroscopy) and computerassisted molecular modeling with biological testing to advance peptidomimetic drug design. A plethora of sophisticated synthetic chemistry approaches have entered into the arena of peptide-based molecular design, including well-established applications of unusual amino acids and dipeptide surrogates, among other types of chemical modifications. Such backbone or side chain modifications may afford stability of the parent peptide to peptidases and have provided conceptual impetus for yet more sophisticated molecular design and peptidomimetic chemistry studies [1-18]. For example, a few of the known amide bond replacements (Figure 4) include: aminomethylene or Ψ [CH2NH], 1; ketomethylene or Ψ[COCH2], 2; ethylene or Ψ[CH2CH2], 3; olefin or Ψ[CH=CH], 4; ether or

Figure 3 Hierarchial approach in peptide rarrow.gif peptidomimetic structure-based drug design

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Ψ[CH2O], 5; thioether or Ψ[CH2S], 6; tetrazole or Ψ[CN4], 7; thiazole or Ψ[thz], 8; retroamide or Ψ[NHCO], 9; thioamide or Ψ[CSNH], 10; and ester or Ψ[CO2], 11. These amide bond surrogates provide insight into the conformational and H-bonding properties that may be requisite for peptide molecular recognition and/or biological activity at receptor targets. Furthermore, such backbone replacements can impart metabolic stability towards peptidase cleavage relative to the parent peptide. The discovery of yet other nonhydrolyzable amide bond isosteres has particularly impacted the design of protease inhibitors, and these include: hydroxymethylene or Ψ[CH(OH)], 12; hydroxyethylene or Ψ[CH(OH)CH2] and Ψ[CH2CH(OH)], 13 and 14, respectively; dihydroxyethylene or (Ψ[CH(OH)CH(OH)], 15, hydroxyethylamine or Ψ[CH(OH)CH2N], 16, dihydroxyethylene 17 and C2symmetric hydroxymethylene 18. In the specific case of aspartyl protease inhibitor design (see below) such backbone modifications have been extremely effective, as they may represent transition state mimics or bioisosteres of the hypothetical tetrahedral intermediate (e.g., Ψ[C(OH)2NH] for this class of proteolytic enzymes. Both peptide backbone and side chain modifications may provide prototypic leads for the design of secondary structure mimicry [11, 22–31] as typically suggested by the fact that substitution of D-amino acids, Nα-Me-amino acids, Cα-Me-amino acids, and/or dehydroamino acids within a peptide lead may induce or stabilize regiospecific β-turn, γ-turn, β-sheet, or α-helix conformations. To date, a variety of secondary structure mimetics have been designed and incorporated in peptides or peptidomimetics (Figure 5). The β-turn has been of particular interest to the area of receptor-targeted peptidomimetic drug discovery. This secondary structural motif exists within a tetrapeptide sequence in which the first and fourth Cα atoms are < 7 Å separated, and they are further characterized as to occur in a nonhelical region of the peptide sequence and to possess a ten-membered intramolecular H-bond between the i and i+4 amino acid residues. On of the initial approaches of significance to the design of β-turn mimetics was the monocyclic dipeptide-based template 19 [22] which employs side chain to backbone constraint at the i+1 and i+2 sites. Over the past decade a variety of other monocyclic or bicyclic templates have been developed as β-turn mimetics, and specific examples include 20 [23], 21 [24], 22 [25], 23 [26] and 24 [27]. Most recently, the monocyclic β-turn mimetic 25 has been described [28] and illustrates the potential opportunity to design scaffolds that may incorporate each of the side chains (i, i+1, i+2 and i+3 positions), as well as five of the eight NH or C=O functionalities, within the parent tetrapeptide sequence. tetrapeptide sequence modeled in type I–IV β-turn conformations. Similarly, the benzodiazepine template 26 has shown [29, 30] utility as a β-turn mimetic scaffold which also may be multisubstituted to simulate side chain functionalization, particularly at the i and i+3 positions of the corresponding tetrapeptide sequence modeled in type I–VI β-turn conformations. A recently reported [31]

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Figure 4 Backbone amide bond surrogates: Ψ[CONH] replacements

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Figure 5 Secondary structure modifications using β-turn α-turn, and β-sheet scaffolds.

γ-turn mimetic, 27, illustrates an innovative approach to incorporate a retroamide surrogate between the i and i+1 amino acid residues with an ethylene bridge between the N' (i.e., nitrogen replacing the carbonyl C') and N atoms of the i and i+2 positions, and this template allows the possibility for all three side chains of the parent tripeptide sequence. Finally, the design of a β-sheet mimetic, 28, provides an attractive template to constrain the backbone of a peptide to that simulating an extended conformation [32]. The β-sheet is of particular interest to the area of protease-targeted peptidomimetic drug discovery.

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12640-0567a.gif Figure 6 Contemporary drug discovery: integration of structure-based drug design, synthetic chemical libraries, and high-throughput mass screening technologies

Finally, the convergence of structure-based drug design (biophysical and computational chemistry), synthetic chemical libraries, and high-throughput screening technologies have established a new paradigm for drug discovery (Figure 6). This powerful alliance of scientific disciplines is accelerating the identification of lead compounds and/or the optimization of drug candidates. The impact to academic, biotechnological and pharmaceutical research will be immense. C. Receptor, Protease, and Signal Transduction-Protein Targets Structure-based drug design and peptidomimetic drug discovery has emerged as a powerful approach in many areas of pharmaceutical research, including receptor agonists and antagonists, protease inhibitors, and, more recently, in the rapidly developing area of signal transduction, in which the protein targets include catalytic and noncatalytic domains of a particular intracellular macro

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molecule. A noncomprehensive listing of such receptor, protease, and signal-transduction protein targets is shown in Table 1. With respect to receptor-targeted peptidomimetic drug discovery, the most noteworthy success has been attained for G-protein-coupled receptor agonists and antagonists as well as, more recently, cell-adhesion integrin receptor antagonists (see below). However, it is important to state that the impact of highthroughput screening in the discovery of nonpeptide ligands (typically antagonists) at G-protein-coupled receptors has yielded extraordinary success. Although screening-based nonpeptide drug discovery will not be extensively reviewed here, the possibility of common pharmacophores between peptide and nonpeptide ligands may exist (limited cases) in relation to receptor binding. Nevertheless, in most cases receptor mutagenesis studies suggest the existence of different binding pockets for peptide and nonpeptide ligands, regardless of whether they both are functionally similar as related to agonism or antagonism [33]. With respect to both protease-and signal-transduction protein-targeted peptidomimetic drug discovery, the emergence of 3D structural information to provide high resolution molecular “maps” of the catalytic (or noncatalytic) domain has provided incredible opportunities for structure-based drug design. II. Peptide Ligand Lead Discovery and Structure-Based Drug Design As stated previously, peptidomimetic drug discovery was first advanced by molecular design concepts and chemical modification strategies focused on

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Figure 7 Convergent pathways in peptide, peptidomimetic, and nonpeptide drug discovery

using receptor-targeted peptide ligands or “second generation” agonist or antagonist analogs to develop pharmacophore models (see Figure 3). However, this represents only a part of the sophisticated convergent pathways that exist currently to advance both peptidomimetic and nonpeptide drug discovery (Figure 7). In this scenario both native and foreign (biological or chemical origin) peptides may provide the opportunity for ligand structure-based drug design. From pharmacophore models of key peptide leads the iterative transformation to peptidomimetic “second generation” analogs may proceed through either peptide scaffold-or nonpeptide template-based approaches. Examples of such work are detailed below. Independent of the hierarchial approach to peptide ligand structure-based design of peptidomimetics is the work that encompasses screening-based nonpeptide lead discovery. Such nonpeptides may be of chemical or biological origin (see below), and historically have been identified by targeted or random screening approaches. Although they are not designed to be peptidomimetics in the chemical structure sense, many appear to be biological mimics (e.g., receptor agonists or antagonists, protease inhibitors, signal-transduction protein inhibitors, or antagonists) of a known native or foreign peptide. And, in few

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cases, a pharmacophore model of a nonpeptide lead (or series) may show similarity to that of a pharmacophore model of a peptide ligand. Examples of such work are detailed below. A. Peptidomimetics: Receptor Agonists and Antagonists Specific examples that illustrate peptide scaffold-and nonpeptide template-directed drug design strategies are shown in Figure 8 and include µ-opioid endorphin (END) agonist, 29 [34]; thyrotropinreleasing hormone (TRH) agonist, 30 [13]; fibrinogen (GPIIa/IIIb) antagonists, 31 [35] and 32 [36]; CCKA antagonist, 33 [37]; CCKB/gastrin antagonist, 34 [38]; endothelin antagonist, 35 [39]; growth hormone secretagogue (GHRP), 36 [40]; somatostatin agonist (partial), 37 [41], substance-P (NK1) antagonists, 38 [42]; neurokinin-A (NK2) antagonist, 39 [43]; and neurokinin-B (NK3) antagonist, 40 [44]. For the most part, the above compounds have been advanced as the result of extensive structureactivity studies and, typically, more focused structural studies (NMR) on a conformationally constrained, linear or cyclic peptide lead or series. Such structure-conformation activity studies have led to the development of pharmacophore models to guide iterative structure-based design strategies. One example is that of integrin receptor gpIIb/IIIa antagonists that are structurally derived from the tripeptide sequence Arg-Gly-Asp, which is common to gpIIb/IIIa protein ligands such as fibrinogen, vitronectin, fibronectin, von Willebrand factor, osteopontin, thrombospondin, and the collagens [45]. As shown in Figure 9, transformations of the linear peptide ligand Arg Gly-Asp-Phe by both peptide scaffold (at the Arg-Gly backbone) modification and substitution of the Arg side chain by a benzamidine moiety provided the peptidomimetic lead 31 that is active in vivo as an antiplatelet agent [35]. On the other hand peptidomimetics such as 32 illustrate nonpeptide template-based design strategies derived from iterative transformations of a cyclic peptide lead in which a γ-turn about the Asp residue was implicated in a pharmacophore model for the bioactive conformation [36]. Specifically, the benzodiazepinone substructure of 32 may effectively replace this predicted γ-turn conformation about the Asp, and the N-Me-Arg replacement with piperidine moiety was also compatible to high-affinity receptor binding. A second example is that involving the use of a glucopyranoside nonpeptide template by Hirschmann and coworkers [41, 46] for systematic functionalization to create novel peptidomimetics for the somatostain (SRIF) and substance-P (NK1) receptors. As illustrated in Figure 10, the cyclic hexapeptide SRIF agonist provided a macrocyclic lead structure that was transformed to a glucopyranoside template designed to substitute for a postulated β turn about the Tyr-D-Trp-Lys-Thr substructure of the parent peptide ligand. The prototype

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Figure 8 Receptor-targeted peptidomimetics exemplifying ligand structure based drug design

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Figure 9 Peptide scaffold- and nonpeptide template-based design strategies: gpIIb/IIIa antagonists

peptidomimetic 37 was found to be a moderately potent SRIF-like agonist (partial) in cellular assays [41]. This discovery extends previous studies on TRH (see peptidomimetic 30, Figure 8) which utilized a cyclohexane ring system as a nonpeptide template to functionalize with the pyroglutamic acod and His side chains as well as the C-terminal carboxamide group of the parent peptide ligand [13]. However, in the comparative analysis of analogs of the SRIF-mimetic 37 it was also found that N-acetylation of the Lys side chain moiety yielded a potent antagonist of substance-P (NK1 receptor). This indicated that slightly different functionalization of the nonpeptidic glucopyranoside template was quite compatible with NK1 receptor molecular recognition. Intuitively, a “reversed design” strategy to convert the latter glucopyranoside-based NK1 ligand to a cyclic peptide was next investigated (Figure 10), and successfully led to the discovery of a novel cyclic peptide ligand also exhibiting potent NK1 receptor binding and antagonism [46]. The above examples of peptide scaffold- or nonpeptide template-based peptidomimetic agonists or antagonists illustrate various strategies to elaborate bioactive conformation and/or pharmacophore models of peptide ligands at their receptors. In many cases, receptor subtype selectivity has also been achieved by systematic structural modifications of prototypic leads of peptidomimetics. Thus, although the 3D structures of G-protein-coupled receptors (GPCRs) remain as elusive (except for models constructed from homology-based low-

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Figure 10 Peptide scaffold- and nonpeptide template-based design strategies: somatostatin agonist and substance-P (NK1) antagonists

resolution 3D structures of bacteriorhodopsin or rhodopsin, see below) the development of pharmacophore models using the hierarchial approach in peptide rarrow.gif peptidomimetic structurebased drug design (see above, Figure 3) remains quite promising to creative science. Indeed, targetedreceptor screening of synthetic chemical libraries of highly modified peptide molecules (e.g., Nsubstituted Gly “peptoids” [21] is rapidly expanding our database of structurally diverse ligands. Such structurally unique lead compounds will provide the opportunity for further pharmacophore modeling strategies to be used in the discovery of novel agonists or antagonists at GPCR and other receptor types. B. Peptidomimetics: Protease Inhibitors Specific examples of peptidomimetics which illustrate peptide scaffold- and nonpeptide templatedirected drug-design strategies as applied to protease in-

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Figure 11 Protease-targeted peptidomimetics derived by ligand structure-based drug design

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hibitors (Figure 11) include renin inhibitors, 41 [47] and 42 [48]; HIV protease inhibitors 43 [49] and 44 [50]; angiotensin-converting enzyme inhibitors 45 [51] and 46 [52]; collagenase inhibitor, 47 [53]; gelatinase inhibitor, 48 [54], stromelysin inhibitor, 49 [55]; elastase inhibitor, 50 [56]; thrombin inhibitors, 51 [57] and 52 [58]; and interleukin-converting enzyme inhibitor, 53 [59]. In general, the design of protease inhibitors has focused on both the natural substrate structure and the mechanism of substrate cleavage to provide “first-generation” inhibitors. Also, these initial leads are typically peptide scaffold-based to provide the possibility for β-sheet conformation, which may permit “extensive” Hbonding between the backbone amide groups of the inhibitor and complementary H-bond donor or acceptor groups of the enzyme active site. Furthermore, the traditional approach to designing protease inhibitors includes the substitution of nonhydrolyzable amide surrogates (see below Figure 4) at the P1P1' cleavage site. Specificity to a particular protease may sometimes be extrapolated directly from the primary structure of the substrate (e.g., human renin substrate specificity is conferred from the Val-Ile-His13~ in which angiotensinogen N-terminal octapeptide sequence ~His6-Pro-Phe-His-Leu refers to the cleavage site). In many cases substitution of the scissile amide (substrate) by a “transition state” bioisosteres or an electrophilic ketomethylene moiety have provided tight-binding “first generation” pseudopeptide inhibitor leads. One example of protease inhibitor design that illustrates the peptide scaffold-based approach is that for HIV protease inhibitors. Albeit over the past several years HIV protease inhibitor research has become a highly advanced example of iterative structure-based drug design (see below), the first discoveries of pseudopeptide and peptidomimetic inhibitors of this aspartyl protease were not made with knowledge of the 3D structure of the target enzyme. Specifically, the natural product pseudopeptide pepstatin (Figure 12), a typical inhibitor of the aspartyl protease family of enzymes, was determined to be weakly potent against HIV-1 protease [60]. Relative to pepstatin, the central P1-P1' statine (i.e., Sta or LeuΨ[CH(OH)]Gly) moiety was further evaluated within the context of an “optimized” N- and Cterminal amino acid sequence using a chemical-library strategy [61]. As shown in Figure 12, a resultant pseudo-tetrapeptide Ac-Trp-Val-Sta-D-Leu-NH2 was found to be a relatively potent HIV protease inhibitor. In another approach, a designed renin inhibitor (55; Figure 12) was determined to be a highly potent HIV protease inhibitor [62]. Further optimization studies led to the discovery [49] of the first bonafide peptidomimetic inhibitor of HIV protease (Tba-ChaΨ[CH(OH)CH2] Val-Ile-Amp; 43) of HIV protease. This compound (43) provided the first evidence of cellular anti-HIV activity and, therefore, supported proof-of-concept studies related to the therapeutic significance of targeting HIV-1 protease. Replacement of the peptide scaffold by the pyrrolidinone-type β-sheet mimetic 28 in a

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Figure 12 Peptide scaffold-based design strategies: HIV protease inhibitors

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chemically-related P1-P1' PheΨ[CH(OH)CH2] Phe-modified lead has been reported [63] to yield effective peptidomimetic inhibitors of the HIV-1 protease (56; Figure 12). The pyrrolidinone-type lead has shown enhanced cellular permeability relative to its peptide backbone-type counterparts. In a third approach guided by HIV substrate-based design, the cleavage site dipeptide Phe-Pro was substituted by the “transition state” bioisostere to provide the highly potent and selective HIV protease inhibitor 57, a P1-P1' PheΨ[CH(OH)CH2N] Pro-modified heptapeptide [64]. As compared to this pseudopeptide, a pioneering effort focused on peptide ligand structure-based design provided a second series of highly potent, selective, and cellularly-active HIV protease inhibitors [50] as represented by the recently FDAapproved anti-HIV drug 44 (Saquinavir). The design of still more HIV protease inhibitors having novel chemical structures (e.g., C2-symmetric scaffolding, P1-P1' “transition state” bioisostere cyclization, and achiral nonpeptide template replacement) has progressed at an extraordinary pace (for reviews see Reference 65), and in a majority of cases such work has been strongly impacted by knowledge of the 3D structure of the target enzyme and/or inhibitor complexes of it (see below). A second example of protease inhibitor design that properly illustrates the peptide scaffold-based approach is that of thrombin inhibitors. Work with these compounds led to the identification of highly potent, selective, and in vivo-effective lead compounds. A member of the serine protease family, thrombin cleaves a number of substrates (e.g., fibrinogen) and activates its platelet receptor (a G-proteincoupled receptor) by proteolysis of the extracellular N-terminal domain which results in self-activation (for a review see Reference 66). Initial lead inhibitors of thrombin were substrate-based, including the fibrinogen P3-P1 Phe-Pro-Arg sequence [67] and simple Arg derivatives such as Tos-Arg-OMe [68]. Also, the natural product cyclothreonide-A, a macrocyclic peptide containing a Pro-Arg ketoamide sequence, provided an inhibitory peptide ligand lead [69]. As shown collectively in Figure 13, compounds 52 and 58–60 provided the opportunity to try different strategies to advance the design of thrombin inhibitors. Particularly noteworthy from these early peptidomimetic lead discovery studies was the design effort [70] that led to the highly potent thrombin inhibitor 52 (Agatroban), a sulfonamidemodified Arg derivative, which incorporated an unusual cyclic amino acid substituent C-terminal to the P1 moiety as opposed to reactive electrophilic groups (e.g., ketone, aldehyde, or boronic acid). Interestingly, replacement of the Arg side chain moiety within a structurally similar analog 60 by a amidinobenzyl group was shown [71] to be optimal when the stereochemistry at the P1 α-carbon has a Dconfiguration suggesting that the mode of binding may be different for 52 versus 60. In this regard, xray crystallographic analysis of thrombin-inhibitor complexes (see below) have provided insight in the interpretation of the structure-activity relationships of the aforementioned lead compounds.

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Figure 13 Peptide scaffold-based design strategies: thrombin and tripeptidyl peptidase II inhibitors

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The recent discovery of peptidomimetic inhibitors of the serine protease TTP-II (tripeptidyl peptidase-II) further illustrates the peptide scaffold-based design approach [72]. Specifically, relative to a known TTPII cleavage site on the endogenous neuropeptide CCK-8 (i.e., Asp1-Tyr[SO H]-Met arrowd.gif Gly-Trp3

Met-Asp-Phe8-NH

) the design of a highly potent inhibitor 61 (Figure 13) was successfully achieved by iterative structure-based optimization of the P3-P1 sequence. Noteworthy of the relatively simple structure of the TTP-II inhibitor 61 was that it contains no functional group C-terminal to the P1 αcarbon. Such absence of an electrophilic moiety, “transition state” bioisostere, or other type of nonhydrolyzable amide substituent is rather unique relative to most examples of substrate-based protease inhibitors. 2

C. Peptidomimetics: Signal-Transduction Protein Inhibitors and Antagonists Beyond receptors and proteases exists the rapidly emerging area of signal-transduction protein-targeted drug discovery research. To date, a multitude of catalytic and noncatalytic proteins have been identified which are critical components of intracellular signal-transduction pathways. These signal-transduction proteins provide the molecular basis for communication from extracellular “effectors” (e.g., hormones, neurotransmitters, growth factors, and cytokines) to stimulate cells in specific and regulated manner. Signal-transduction pathways often involve protein-protein interactions, including examples of enzymesubstrate (e.g., kinases, phosphatases, transferases, and isomerases) as well nonenzymatic complex formation (e.g., “adapter” proteins, exchange factors, and transcription factors). As compared to receptoror protease-targeted peptidomimetic drug discovery, there are significantly fewer examples reported in the field of signal-transduction research. Thus, in several cases peptide ligands (as “prototype peptidomimetic leads”) will be described to illustrate opportunities for structure-based drug design. Specific examples which illustrate peptide scaffold- and nonpeptide template-directed drug-design strategies are shown in Figure 14 and include: Ras farnesyl transferase inhibitors, 62 [73], 63 [74], 64 [75], and 65 [76], Src SH2 domain antagonists, 66 [77], 67 [78], and 68 [79]; and the protein tyrosine phosphatase PTP1B inhibitor 69 [80]. An example of the signal-transduction protein-targeted inhibitor design which illustrates both peptide scaffold- and nonpeptide template-based approaches is that for the Ras farnesyl transferase inhibitor discovery. Such compounds show potential as new therapeutic agents for Ras-related carcinogenesis [81]. Substrate sequences for farnesyl transferase have the consensus ~Cys-AA1-AA2-Met motif (AA refers to Val or Ile). Both substrate-based

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Figure 14 Signal-transduction protein-targeted peptidomimetics derived by structure-based drug design

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inhibitors [73–75,82,83] and, more recently, a novel non-Cys containing peptide inhibitors [76,84] have led to potent and cellularly active compounds. As illustrated in Figure 15, the “collected” substratebased inhibitor 70 was designed to covalently attach farnesyl to a peptide via a phosphinic acid linker replacement for [82], and this compound has been determined to be both potent against the target enzyme and cellularly effective. Relative to peptide substrate structure-based design efforts, peptidomimetics incorporating Ψ[CH2NH]-substitutions (e.g., 62, [73]) or a benzodiazepinone replacement of the central dipeptide moiety (e.g., 63, [74]) have yielded high affinity inhibitors. Another series of very potent Ras farnesyl transferase inhibitors have been designed in which the central dipeptide has been substituted by various isomeric and/or homologated derivatives of amino benzoic acid (e.g., 64 [75]), including a particularly effective analog biphenyl derivative 71 [83]. The above studies indicated that both conformationally flexible or constrained peptide scaffolds as well as nonpeptide template replacements can be used to “link” the Cys and Met substructures. It is also important to point out that although compounds such as 62–64 have “free” sulfhydryl groups (Cys) there is no evidence that they become farnesylated, and therefore the binding mode and effect on catalytic function of the target enzyme are unique relative to their peptide substrate counterparts. Recently, a novel peptide inhibitor series, as exemplified by Cbz-His-Try(O-benzyl)-Ser(O-benzyl)-Trp-D-Ala-NH2, has been discovered [76]. These inhibitors do not contain a Cys residue and structure-based design efforts have successfully led to a series of peptidomimetics (e.g., 65) having only one chiral center. Interestingly, this novel inhibitor series has been determined to competitively inhibit farnesyl pyrophosphate binding rather than the binding of peptide substrate to the target enzyme. Another Hissubstituted peptidomimetic inhibitor of Ras farnesyl transferase has been recently reported [84] as exemplified by 73, which was designed relative to a peptide substrate-based parent analog (72, Figure 15). Although the 3D structure of Ras farnesyl transferase is not known, biochemical studies suggest that a divalent metal ion (e.g., Zn2+) may coordinate with the above inhibitor sulfhydryl or imidazole groups at their corresponding binding sites on the target enzyme. Another example of the signal-transduction protein-targeted drug design that illustrates peptide scaffoldbased approaches is that for Src SH2 domain antagonist discovery. Such compounds show promise as new therapeutic agents for Src-related carcinogenesis, osteoporosis, and immune diseases [85]. The Src SH2 domain is a prototype example of a superfamily of intracellular signal-transduction proteins that possess structurally homologous SH2 domains that specifically recognize cognate phosphoproteins in a sequence-dependent manner relative to a critical phosphotyrosine (pTyr) residue (i.e., ~pTyr-AA1-AA2AA3-AA4~ Furthermore, recent x-ray crystallographic studies of a several SH2 protein targets (e.g., phosphopeptide complexes) is greatly impacting the

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Figure 15 Peptide scaffold- and nonpeptide template-based design strategies: farnesyl transferase inhibitors

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opportunity for iterative structure-based drug design in this field or research (see below [86]). Relative to Src SH2 domain antagonist lead discovery, peptide library studies [87] have shown the ~pTyr-GluGlu-Ile~ as a preferred consensus sequence. Peptide scaffold-based approaches to replace the internal dipeptide, Glu-Glu, by both flexible and rigid linkers have been explored [88] but were unsuccessful in yielding potent analogs. As shown in Figure 16, prototype peptidomimetics 66 [77] and 67 [78] illustrate a successful approach in which stereoinversion at the second residue (P+2 relative to the pTyr) to the Dconfiguration and side chain substitution to hydrophobic functionalities (e.g., cyclohexyl and naphthyl) which provides accessibility to the known hydrophobic binding pocket for the P+3 Ile side chain. Indeed, such compounds showed binding affinities essentially identical to that of the N- and Cterminally extended phosphopeptides containing the pTyr-Glu-Glu-Ile sequence. Substitution of the pTyr residue of 66 by the difluoromethyl-phosphonate modified analog F2Pmp provides a more metabolically stable derivative 74 (Figure 16) and a prototype lead to advance the design of cellularly active second-generation compounds. More recently, structure-based drug design studies of compound 65 have led to the discovery of potent and Src SH2-selective peptidomimetic lead compounds [89], and this is further detailed below as related x-ray crystallographic structures of Src SH2-phosphopeptide complexes. III. Nonpeptide Ligand Lead Discovery and Structure-Based Drug Design As previously illustrated in Figure 7, the convergent pathways to design drugs which act as mimics (agonists), antagonists, or inhibitors of native peptide (protein) ligands at their target receptors, proteases, signal-transduction proteins, and so forth, include “foreign” nonpeptides. The origin of such nonpeptides may be either biological (e.g., natural product) or chemical (synthetic compound collection or libraries) that have been subject to biochemical screening to identify leads for further molecular design and structure-activity studies. Over recent years the success of screening-derived nonpeptide lead discovery and iterative transformation to drug candidates has been quite impressive, and many aspects of this area of research have been reviewed [12, 15]. Nevertheless, it is intriguing to explore the potential relationship between peptide ligands (including peptidomimetic derivatives) and such screening-derived nonpeptide ligands as related to pharmacophore modeling and structure-based drugdesign studies.

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Figure 16 Peptide scaffold-based design strategies: Src SH2 domain antagonists

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A. Molecular Diversity and Screening-Based Identification of Nonpeptide Ligands Precedence for the success of nonpeptide drug discovery can be traced to the identification of morphine (75; Figure 17) as nonpeptide natural product agonist at µ-opioid peptide receptors [90]. To date, the robust momentum of nonpeptide drug discovery continues to be accelerated by sophisticated biochemical screening technologies. The scope of molecular diversity as well as therapeutic targets for such screening-based nonpeptide ligand lead compounds includes the following examples (Figure 17): substance P (NK1 antagonist, 76 [91]; angiotensin AT1 antagonist, 77 [92]; growth hormone-releasing peptide (GHRP) agonist, 78 [93]; cholecystokinin CCKA antagonist, 79 [94]; CCKB/gastrin antagonist, 80 [95]; CCKA agonist, 81 [96]; endothelin antagonist, 82 [97]; gonadotropin-releasing hormone (GnRH) antagonist, 83 [98]; vasopressin V1 antagonist, 84 [99]; gastrin-releasing peptide antagonist, 85 [100]; glucagon antagonist, 86 [101], neurotensin antagonist, 87 [102]; angiotensin AT1 agonist, 88 [103]; oxytocin antagonist, 89 [104]; and HIV protease inhibitor, 90 [105]. A significant number of screening-derived nonpeptide leads have been identified for G-protein-coupled receptors (GPCRs), and in a majority of cases these compounds have been determined to be competitive antagonists. Albeit the 3D structures of this receptor superfamily have not been directly determined, homology model-building and site-directed mutagenesis studies are impacting structure-activity analysis of agonist and antagonist ligands (peptide, peptidomimetic, and nonpeptide) for several GPCR targets (see below). B. Nonpeptides: Exploring Pharmacophore Relationships to Peptide Ligands Relative to a number of GPCR targets, there exists significant opportunity to compare chemical structures and 3D pharmacophore models of both peptide and nonpeptide ligands. Such comparative analyses can explore the possibility of similar 3D substructural elements that may account for their molecular recognition at the binding site(s) of the receptor. The fact that a vast number of screeningderived nonpeptide leads are multifunctionalized 5- to 7-membered ring heterocycles (e.g., alkaloid and benzodiazepine) and contain conformationally rigid substructural elements (e.g., biphenyl, spiro-bicyclic rings, and N-substituted amide or amine linkages) suggests the likelihood that such compounds are binding with highly favorable entropic driving forces as compared to the more conformationally flexible peptide ligands. In this regard, efforts to “rigidify” peptide-based scaffolding or replace it by nonpeptide templates has been the underlying theme of peptidomimetic design strategies, and concepts for the latter approach date back to the proposed used of cycloaliphatic ring systems that might be multifunctionalized to create “topographi-

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Figure 17 Nonpeptide drug discovery: examples from screening-based approaches

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cally-designed” peptidomimetics (or, as also defined, “peptoids” [106]). Nevertheless, screening-based nonpeptide drug discovery has advanced a treasure of structure-function information to provide insight into both structure-based design and molecular recognition [12,15,107]. In a few (limited) cases, there exists a likely possibility of similar pharmacophoric features or substructural elements between nonpeptides and their peptide-ligand counterparts (Figure 18). Historically, drug discovery research on opioid GPCR receptor targets (e.g., µ, δ, κ) has provided insight to explore the pharmacophores of both agonist and antagonists derived from endogenous peptides (e.g., endorphin, endorphin, and dynorphin) versus nonpeptides (e.g., the µ-receptor selective agonist morphine and its N-allyl-substituted antagonist derivative naloxone). Relative to the N-terminal Tyr moiety (side chain and α-amino functionalities) of the endogenous opioid peptides [108], the N-methyltyramine substructure of morphine represents a likely common pharmacophore for agonist ligand binding to the µ-receptor (Figure 18). In the case of angiotensin II receptor antagonist drug discovery, it has been proposed [109] that a common pharmacophore may exist relative to the C-terminal His-ProPhe-OH sequence of angiotensin II and nonpeptide 91 (Figure 18). In fact, these studies provided design insight leading to the discovery of the drug candidate 77 (Lorsartan). A third example in which correlation between peptide and nonpeptide pharmacophore models becomes apparent is that of neuropeptide-Y (NPY) versus the benextramine-based derivative 92 [110] or arpromidine-based derivative 93 [111] as illustrated in Figure 18. In both cases, the C-terminal Arg-Gln-Arg-Tyr-NH2 sequence of NPY was modeled relative to the nonpeptide structures such that the guanido functionalities were superimposed upon the corresponding basic (i.e., guanido or imidazole) substructural elements of either 92 or 93. In some cases, the availability of x-ray crystallographic information of both the peptide and nonpeptide ligands may provide insight into the pharmacophore modeling studies. An example of this exists for oxytocin antagonist structure-based drug design studies [112]. As shown in Figure 19, pharmacophore models of both a cyclic hexapeptide oxytocin antagonists and conformationally-constrained, tolylpiperazine camphorsulfonamide nonpeptide antagonist (89) suggest the likelihood of common substructural elements that were key for molecular recognition at the oxytocin receptor, and led to the design of a highly potent derivative 94. Nevertheless, such comparative pharmacophore “mapping” studies are very simplistic since the 3D structures of the target receptors are not known. Furthermore, site-directed mutagenesis studies

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Figure 18 Comparative substructural elements of peptide and nonpeptide ligands: µ-opioid receptor agonists, angiotensin, and NPY receptor antagonists

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Figure 19 Comparative substructural elements of peptide and nonpeptide ligands: examples of oxytocin receptor antagonists and HIV protease inhibitors

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often suggest that peptide and nonpeptide ligands have different modes of binding to their receptors (see below). It is also important to point out that the discovery of nonpeptide agonists will likely provide important structural information to advance our understanding of ligand binding and activation of receptors as well as insight for pharmacophore modeling. In this regard, nonpeptide agonists 78 (growth hormone-releasing peptide receptor) and 81 (cholecystokinin receptor CCKA subtype) are noteworthy exceptions to the rule that nonpeptide screening-based leads are antagonists (see above, Figure 17). Finally, beyond receptor targets the advantages of high-throughput screening of chemical files, natural products, and synthetic libraries are increasing for proteases as well as other enzyme and noncatalytic targets. An recent example of such efforts is the nonpeptide HIV protease inhibitor 90 [105], which was originally identified from screening a chemical file. Through iterative structure-based drug design studies, including x-ray crystallographic analysis of both ligand and inhibitor-enzyme complexes, the pyrone template has led to the discovery of highly potent, selective, and cellularly active lead compounds (see below). In retrospect, the original concept of superimposing the key substructural elements of the nonpeptide ligand 90 to a known peptidomimetic inhibitor of HIV protease is illustrated in Figure 19. A more detailed account of this effort and successful elaboration of the nonpeptide lead structure is described below. IV. Protein Target 3D Structural Models and Structure-Based Drug Design A significant impact in both peptidomimetic and nonpeptide drug discovery has emerged over recent years as the result of the determination of the 3D structures of protein targets by x-ray crystallography or NMR spectroscopy [113–119]. In addition, computational methodologies such as QSAR, 3D QSAR/CoMFA, homology-modeling, ligand docking, molecular dynamics, and mechanics, and solventaccessible surface visualization have greatly impacted such research. Furthermore, programs such as GRID, GROW, GrowMol, LEGEND, BUILDER, LUDI, FOUNDATION-SPLICE, and CONCERTS have enabled 2D and 3D database searching and de novo design [115,120,121]. Overall, the iterative cycle of structure-based drug design (Figure 20) has evolved to be an “engine of invention” for several examples of peptidomimetic and nonpeptide drug discovery.

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Figure 20 Iterative cycle(s) for structure-based drug design

A. Receptor Targets Amongst the known superfamilies of cell membrane anchored receptors, significant research has been focused on GPCR targets (Table 1). The GPCR superfamily of receptors consist of seven transmembrane-spanning (TM) helices. Sequence homology among them varies from 25–70% (for reviews see References 33, 122–132). The initial development of 3D structural models of GPCR targets has been developed from homology-building methodologies based on a low-resolution structure of bacteriorhodopsin [133]. Representative examples of recent studies that provide insight to pharmacophore modeling and structure-based drug design of peptide, peptidomimetic, and nonpeptide ligands are discussed below. 3 G-Protein-Coupled Receptors Recent studies have explored several GPCR targets with respect to ligand-receptor binding interactions using site-directed mutagenesis to explore agonist

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versus antagonist ligands, and such work provides insight to pharmacophore modeling. Specific examples of such work as focused on peptide, peptidomimetic, and/or nonpeptide ligands, and working 3D structural models of their GPCR targets include angiotensin II AT1 and AT2 subtypes [134], neurokinin NK1 and NK2 subtypes [135], cholecystokinin/gastrin CCKA and CCKB subtypes [136], opioid µ-, δ-, and κ-subtypes [137], vasopressin V1A subtype [138], bradykinin B2 subtype [139], neurotensin [140] and α-melanotropin MC1 subtype [141]. From such work it has been inferred that different binding-site interactions may exist for peptide versus nonpeptide ligands as based on their differential sensitivities to site-directed mutants of the native GPCR. The recent development of 3D structural models of the neurotensin [140] and α-melanotropin MC1 subtype [141] GPCRs provide interesting case studies. Both examples provide the correlation of significant structure-activity databases and experimentally determined (NMR) structures of key peptide analogs with predicted molecular contacts at their respective target receptors. As illustrated in Figure 21, the proposed peptide agonist binding interactions for neurotensin and α-melanotropin analogs at their human GPCR targets may be used to further guide the molecular design and synthesis of “second-generation” peptidomimetic derivatives. In the first example, the neurotensin C-terminal octapeptide was subject to conformational searching (~Arg-Pro-Tyr~ sequences from the Brookhaven Protein Databank), manual docking to the homologybuilt neurotensin GPCR receptor model, and constrained molecular dynamics simulation to provide a 3D structure of the ligand-receptor complex [140]. A compact structure of the peptide in its complexed conformation was consistent with a Type-1 β-turn as previously determined by structural and structureactivity studies. Key molecular contacts predicted from this neurotensin GPCR model include hydrophobic interactions with the C-terminal Ile and Leu side chains, π-cation interactions with each Arg residue side chain, and a “cluster” of aromatic-aromatic interactions with the Tyr side chain. No electrostatic interactions were predicted, and the primary contact residues on the neurotensin GPCR model were those comprising the third extracellular loop. In the second example, the α-melanotropin (MC1) GPCR model was constructed [141] by homologybuilding methods relative to both bacteriorhodopsin and rhodopsin fingerprint maps, and the MSH superagonist peptides [Nle4, D-Phe7]-MSH and Ac-cyclo[Nle4, Asp5, D-Phe7, Lys10]-MSH4–10-NH2 were modeled in conformations derived from previous experimental studies (i.e., a Type-II β-turn at the common tetrapeptide sequence ~His-D-Phe-Arg-Trp~). Of the alternative binding modes that were described for the above the two MSH peptide ligands, one predicts the possibility of a network of aromatic-aromatic and hydrophobic interactions between the MC1 receptor and the D-Phe and Trp side chains of the MSH ligand (Figure 21). In addition, this MC1 receptor model predicts multiple electrostatic and π-cation interactions between

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Figure 21 GPCR 3D structural models for neurotensin and α-melanotropin agonists

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the MC1 receptor and the Arg side chain of the peptide ligand. The primary contact residues of this particular MC1 receptor model were all transmembrane domain derived and lie within 4–7.5 angstrom (centroid to centroid). In conclusion, the development and refinement of 3D structural models of GPCR targets, iterative site-directed mutagenesis studies, and systematic testing of key agonists and/or antagonists will undoubtedly make a significant impact in the structure-based drug design of peptidomimetic and nonpeptide ligands at these receptors. Such work may be expected to synergize well with ligand-based pharmacophore modeling strategies which have become quite sophisticated in recent years as the results of advanced computational chemistry methodologies. B. Protease Targets Protease-inhibitor drug discovery illustrates significant success in both mechanistic and 3D structurebased drug discovery for each of the representative classes (i.e., aspartyl, serinyl, metallo, and cysteinyl) as exemplified in Table 2 (for a review see Reference 142a). In retrospect, pioneering achievements in the design of peptidomimetic inhibitors of angiotensin-converting enzyme (for reviews see References 142b,142c) to have led to 45 (Captopril [51]) and 46 (Enalapril [52]). Such work has provided great impetus to the area of proteasetargeted drug discovery. Over the past two decades a pervasive effort integrating substrate-based inhibitor design, x-ray crystallography or NMR spectroscopy of inhibitorprotease complexes, high-throughput mass screening, and combinatorial chemical technologies has evolved to further advance this area of research. Aspartyl Proteases The aspartyl proteases include pepsin, renin, cathepsin-D, chymosin, and gastricsin as well as microbial enzymes (e.g., penicillopepsin, rhizopuspepsin, and endothiapepsin) and retroviral proteases (e.g., HIV1 protease). The first high-resolution x-ray crystallographic structures of this protease family were determined for penicillopepsin [143], rhizopuspepsin [144], endothiapepsin [145], pepsinogen [146], and pepsin [147]. Based on homology-building strategies, 3D structural models of renin were subsequently constructed [148] to first guide the structure-based design of peptidomimetic inhibitors (for a review see Reference 149). Furthermore, in several cases the x-ray crystallographic structures of renin inhibitors were determined [150] as ligand-enzyme complexes with rhizopuspepsin, endothiapepsin, or pepsin. Eventually, the x-ray crystallographic structures of renin (apo/complexes) were achieved to provide high-resolution molecular maps of the target enzyme [151]. As illustrated in Figure 22, substratebased inhibitors such as the highly potent peptidomimetic 95 [151a] show well-defined hydrophobic pockets for the P3-P1' side chains as well

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as hydrogen-bonding to the backbone of the inhibitor that exists in β-sheet type extended conformation. Structure-based design strategies of renin inhibitors have focused on systematic transformation of its substrate (angiotensinogen). Noteworthy examples which illustrate topographical designed include inhibitors 96 [152], 97 [153], 98 [154], and 99 [155], of which the latter macrocyclic inhibitor is quite novel in terms of having two D-aromatic amino acids and lacking a “transition state” bioisostere replacement at the P1-P1' site. In contrast to renin, the discovery of HIV protease inhibitors provides a high degree of synchronization of x-ray crystallography studies with iterative structure-based drug design efforts as well as the identification of nonpeptide ligands from mass screening (e.g., coumarins and pyrones) or 3D computational searching (e.g., haloperidol) to advance what has become a milestone achievement in rational drug design [113]; and for reviews on HIV protease see Reference 156 and the first chapter of this book). Representative of the scope of the many contributions to the discovery of both peptidomimetic and nonpeptide inhibitors of HIV protease (Figure 23) are the de novo designed C2symmetric

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Figure 22 Protease 3D structural models: renin-inhibitor complex and drug design

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Figure 23 Protease structure-based drug design: HIV protease-targeted peptidomimetics and nonpeptides

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inhibitors 100 [157] and 101 [158]; the nonsymmetric peptidomimetics 44 [50] and 102–106 [159–163, respectively]; and a series of nonpeptides derived originally from either 3-D computational searching 107 [164] or high-throughput sceening 108–110 [165–168], respectively. Of these compounds, FDA approval has been recently granted to 102 (Indinivar), 103 (Ritonavir), and 44 (Saquinavir). Currently, it is believed that there exists well over 150 x-ray crystallographic structures of HIV proteaseinhibitor complexes, not including mutated forms of the target enzyme that have also been determined to be important to develop inhibitors which will be effective in so-called HIV resistant strains. The initial series of x-ray crystallographic structures of HIV protease included the apoprotein [169] and enzymeinhibitor complexes derived from substrate-based analogues having P1-P1' substitutions by N1eΨ[CH2NH]N1e [170], LeuΨ[CH(OH)CH2]Val [171], PheΨ[CH(OH)CH2N]Pro [172], and LeuΨ[CH(OH)]Gly or statine [173]. As illustrated in Figure 24, the first reported HIV protease-inhibitor complex [170] with the pseudopeptide 111 provided a high-resolution map of the active site of the enzyme as formed in a C2-symmetric fashion by the homodimer, and the “flaps” of each monomeric subunit (i.e., residues 35–57) were shown to make intermolecular interactions with the backbone of the inhibitor by both direct hydrogen-bonding and through a structural water molecule (W301). Relative to the C2-symmetry of the target enzyme, the discovery of C2-symmetric inhibitors was successfully achieved by the design of PheΨ[CH(OH)]gPhe- and PheΨ[CH(OH)CH(OH)]gPhe-modified peptidomimetics (gPhe refers to gem-diamino-Phe in which the Cα-CO2H moiety is replaced by CαNH2) as exemplified by 101 [157] Among the plethora of other structure-based drug design strategies focused on HIV protease inhibitor discovery it is also noteworthy to highlight the nonpeptide leads 100 and 108–111 as they displace a key structural water (i.e., W301) and, as opposed to all previously discovered substrate-based inhibitors, are capable of direct hydrogen-bonding interactions to the HIV protease flap regions (Figure 24). These discoveries provide impetus for molecular design strategies to consider tightly bound water molecules as possible secondary “ligands” in either de novo design or iterative structure-based design of novel peptidomimetic or nonpeptide lead compounds. Previous studies [174] on biotinstrepavidin and an x-ray crystallographic structure of the complex showed that structural water molecule displacement (relative to the apoprotein) by key functional groups of the ligand (biotin) was possible. It is noted, however, that HIV protease is unique from other members of the aspartyl protease family with respect to the role of structural water W301 role in substrate/ inhibitor binding. The catalytic water, which is critical to the mechanism of substrate cleavage for all aspartyl proteases, has been a key feature in the design of a plethora of so-called “transition state” modified inhibitors that incorporate a tetrahedral

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Figure 24 Protease 3D structural models: HIV protease-inhibitor complex and drug design

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hydroxymethyl substituent within various types of nonhydrolyzable surrogates of the scissile amide bond (e.g., Ψ[CH(OH)], Ψ[CH(OH)CH2], and Ψ[CH(OH) CH2N]; see above). Nevertheless, there exist examples of highly potent inhibitors of renin e.g., the Ψ[CH2NH]-modified 41 [47] and macrocylic peptide 99 [155] and HIV protease e.g., the pyrone-based series 108–111 [165–168] respectively) that do not possess a tetrahedral CH(OH) moiety per se. Serinyl Proteases The serinyl proteases include trypsin, chymotrypsin-A, elastase, thrombin, kallikrein, cathepsins-A, G, and R, Factor VII, Factors IXa-XIIa, and tissue plasminogen activator. High-resolution x-ray crystallographic structures of this protease family have been determined for thrombin (for a review see [175]; also refer to Table II [176–182]), Factor Xa [183], trypsin [184], kallikrein-A [185], and elastase [186–190]. As illustrated in Figure 25, a substrate-based inhibitor of thrombin having a boronic acid, B(OH)2, substitution for the scissile amide was determine by x-ray crystallography to form a covalent bond to the active site Ser-195 residue [176]. The N-terminal Ac-D-Phe-Pro moiety of this inhibitor binds in a β-sheet type extended conformation that involves hydrogen-bonding contacts to the enzyme and well-defined hydrophobic and aromatic-aromatic (edge-to-face) stacking interactions. The inhibitor Arg side chain binds in an extended conformation and the guanidino moiety forms bidentate hydrogenbonding interactions with an Asp189 residue at the base of the S1 “specificity” pocket as well as additional hydrogen-bonds to the enzyme, one of which is mediated through a structural water. Relative to the substrate-based peptidomimetic inhibitors of thrombin having C-terminal electrophilic groups (e.g., aldehyde, ketone, and boronic acid), the discovery and structure-based design of nonpeptide inhibitors not having P1 electrophilic functionalization has also been extremely successful as represented by 52 [70], 60 [71], and 113 [182]. As shown in Figure 25, the design of a highly potent and selective amidinopiperidine-based thrombin inhibitor 113 was derived from analysis of the x-ray crystallographic structures of thrombin complexed with inhibitors 52 and 60. The latter two compounds showed different trajectories of their P1 side chains (i.e., guanidinoalkyl and amidinophenyl, respectively) into the S1 pocket to account for the observed opposite chirality preferences at the Cα-position of the P1 amino acid residues. Also, the C-terminal cycloalkyl moieties of both 52 and 60 were observed to bind to the socalled inhibitor “P-pocket” (i.e., the P2 substrate pocket), thus explaining that these compounds were not binding in a substrate-like conformation such as the peptidomimetic inhibitors Ac-D-Phe-Pro-boroArgOH as described above. Thus, the design of the novel amidinopiperidine-based inhibitor 113 illustrates a “transposition” of the P-pocket binding group to an N-substituted Gly-β-Asp scaffold.

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Figure 25 Protease 3D structural models: thrombin-inhibitor complex and drug design

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For other serinyl proteases, such as Factor Xa, kallikrein and elastase, the availability of x-ray crystallographic structure of the enzymes (apo/complexes) provides further examples in which structurebased drug design is being advanced (Table 2). In particular, elastase-targeted drug discovery is highlighted here as it illustrates substrate-based peptidomimetic inhibitor design strategies that have focused on key P2–P3 side chain and backbone hydrogen-bonding interactions with the enzyme (for reviews see Reference [191]). Several x-ray crystallographic structures have been determined for pancreatic elastase [187,188,189b,190,191] and leukocyte elastase [186,187,189a], including complexes with peptide substrate-based inhibitors having P1 electrophilic functionalities such as benzoxazole, [188] trifluoromethyl ketone [56,187,189], and α-,α-difluoro-β-ketoamide [190]. Recently, the design of peptidomimetic inhibitors incorporating nonpeptidyl P2-P3 replacements has resulted in the discovery of highly potent compounds [56,187]. Specifically, a lead series of highly potent trifluoromethylketonebased inhibitors of human leukocyte elastase which incorporate a N-carboxymethyl-3-amino-6-arylpyridone template (50, 116; see Figure 26) was developed and shown by x-ray crystallography to provide backbone hydrogen-bonding and a novel trajectory of a P2 group from the pyridone ring to the enzyme. Further modification of the pyridone ring to give the bicyclic pyridopyrimidine derivative 117 was predicted from molecular modeling studies to provide additional hydrogen-bonding to the enzyme as well as another site on the bicyclic heteroaromatic ring system for tethering various hydrophobic or hydrophilic groups. Finally, a series of novel dipeptide-based inhibitors (e.g., trifluoromethylacetyl-LeuPhe-p-isoproylanilide and a peptidomimetic derivative) are particularly intriguing because they bind “backwards” as based on analysis of the x-ray crystallography structures of their complexes with pancreatic elastase [191]. In this binding mode the trifluoromethylacetyl moiety is proximate to the active site Ser residue and the ligand backbone and side chain substructures make hydrogen bonding and hydrophobic contacts with the enzyme, respectively. Cysteinyl Proteases The cysteinyl proteases include papain; calpains I and II; cathepsins B, H, and L; proline endopeptidase; and interleukin-converting enzyme (ICE) and its homologs. The most well-studied cysteinyl protease is likely papain, and the first x-ray crystallographic structures of papain [193] and a peptide chloromethylketone inhibitor-papain complex [194] provided the first high resolution molecular maps of the active site. Pioneering studies in the discovery of papain substrate peptide-based inhibitors having P1 electrophilic moieties such as aldehydes [195], ketones (e.g., fluoromethylketone, which has been determined [196] to exhibit selectivity for cysteinyl proteases versus serinyl proteases), semicarbazones, and nitriles are noteworthy since 13C-NMR spectro-

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Figure 26 Protease 3D structural models: elastase-inhibitor complex and drug design

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scopic studies provided direct evidence that the active site Cys25 of papain forms a reversible covalent bond with such electrophiles (see review Reference 142). The x-ray crystallographic structures of cathepsin-B [197] and picornaviral 3C protease [198] have also been determined, but only in the apoprotein form. Recently, high-resolution x-ray crystallographic structures of ICE-inhibitor complexes have been determined [199,200]. This enzyme is structurally unique relative to other cysteinyl proteases (for a review see Reference 201) in that it has a heterodimeric architecture in which two subunits form the catalytically active enzyme site (actually, two p10/p20 heterodimers apparently create a tetrameric form of the competent protease). As shown in Figure 27, an x-ray crystallographic structure of ICE complexed with a substrate peptide-based chloromethylketone inhibitor (118 [200]) that is irreversibly bonded to the active site Cys285 shows the P1 Asp specificity pocket to be comprised of two Arg residues that lie at the base of the S1 binding pocket. Relative to other side chain binding pockets, a hydrophobic “channel” type S4 site exists for the P4 Tyr of the inhibitor, whereas the P3-P2 Val-Ala side chains are well exposed to solvent. With respect to the peptide backbone of the inhibitor, hydrogen bonding interactions between the P3 Val (both NH and CO) and the P1 Asp (NH) and the P10 monomer are predicted. The x-ray crystallographic structure of ICE complexed with the inhibitor Ac-Tyr-Val-Ala-Asp-aldehyde (119 [202]), supports structure-activity studies [202] that employed a systematic analysis of N-methylamino acid substitutions in which hydrogen bonding interactions between inhibitor and enzyme tolerated only N-Me-Ala replacement at the P2 site. Furthermore, C-terminal modification of P1 Asp by irreversible alkylating groups, such as the aryloxymethyl ketone analog 120 [203], have led to the first reported peptidomimetic inhibitor of ICE (121 [204]). Noteworthy in the structure of this peptidomimetic inhibitor is that a pyridone template provided an effective P2-P3 replacement, similar to that for the elastase inhibitor design (see above). It is likely that the backbone hydrogen bonding network between 121 and ICE is conserved as compared to the x-ray crystallographic structure of the substrate peptide-based inhibitor 118 complexed to the target enzyme. Finally, the x-ray crystallographic structure of the ICE homolog referred to as apopain, or CPP32, as a complex with a peptide-aldehyde inhibitor has been recently determined [205] and provides additional insight as the specificity of substrate recognition at both the S1 (P1 Asp) and S4 (P4 Asp) subsites. Thus, the opportunity for iterative structure-based drug design exists to advance novel peptidomimetic and nonpeptide inhibitors of ICE and/or its homologs. Metalloproteases The metalloproteases include both exopeptidases (e.g., angiotensin-converting enzyme, aminopeptidaseM, and carboxypeptidase-A) and endopeptidases (e.g.,

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Figure 27 Protease 3D structural models: ICE-inhibitor complex and drug design

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(e.g., thermolysin, endopeptidase 24.11 or NEP, collagenase, gelatinase and stromelysin). Historically, the most well-studied metalloprotease is thermolysin (for a review see [206]), and the first x-ray crystallographic structures of thermolysin [207] and several structurally distinct peptide inhibitors (see Table 2 [208–212]) provided the first high-resolution molecular maps of the active site and insight into the mechanistic roles of the metal ion for substrate hydrolysis. Specifically, the binding interactions of P1–P1' “transition state” amide bond isosteres (e.g., ψ[P=O(OH)NH]) as well as metal-chelating functionalities (e.g., hydroxamates, carboxylates, and sulfhydryls), introduced as N-substitutions on P1–'P2' peptide scaffolds [208–212] have been determined. Moreover, the structural and mechanistic information derived from studies on thermolysin have provided insight into the design of inhibitors of the therapeutically relevant target, angiotensin-converting enzyme or ACE (for reviews see Reference 142). This has been particularly significant since the 3D structure of ACE has not yet been determined. However, it is noted that x-ray crystallographic structures of carboxypeptidase, a related metalloprotease of the exopeptidase group, for both the apoprotein and a Glyψ[P=O(OH)NH]Phe-modified inhibitor complex have been reported [213]. As illustrated in Figure 28, the x-ray crystallographic structure of the Pheψ[P=O(OH)NH]Leu-modified peptidomimetic inhibitor 122 complexed with thermolysin show the Zn2+ coordination and P1'-P2' (or P1-P2' “collected product”) mode of binding. Relative to this structure the predicted molecular interactions between ACE and its inhibitors (e.g. Captopril [51], Enalaprilat [52], and Fosfinoprilat [214] as well as an emerging class of “dual specific” inhibitors of ACE and NEP (e.g., 123 [215]) may be envisaged. The ability to design specific inhibitors of several matrix metalloproteases (MMPs) is rapidly developing (for reviews see Reference 216). Such MMP targets include fibroblast collagenase (MMP-1), gelatinase-A (MMP-2), and stromelysin (MMP-3). Both x-ray crystallography and NMR spectroscopy have provided 3D structural information for several MMPs as well as MMP-inhibitor complexes (see Table 2; [217–226]). In the example of collagenase, “first generation” substrate-based inhibitor design strategies have focused on modifying the P1–P1' cleavage site (e.g., Gly-Phe, Ala-Tyr, and Ala-Phe) by N-terminal functionalities capable of Zn2+ coordination (e.g., sulfhydryl, carboxyl, phosphonoalkyl, and hydroxamate [227–229]). As shown in Figure 29, an x-ray crystallographic structure of the Nhydroxamate-modified peptide 124 complexed to fibroblast collagenase provides a molecular map of both its hydrogen-bonding interactions to the enzyme active site and binding to bound Zn2+ [219]. Potent MMP-1 inhibitors have been designed that tether the P2' side chain to the inhibitor's C-terminus as macrocylic rings [53,230]. In the example of gelatinase-A, potent inhibitors have been designed (e.g., 48, Figure 11 [54]) by N-terminal hydroxamate and P1' extended aromatic side chain modifications

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Figure 28 Protease 3D structural models: thermolysin-inhibitor complex ACE inhibitor drug design

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of a P1'–P3' peptide scaffold [54]. To date, no 3D structural information is available for MMP-2 with respect to either the apoprotein catalytic domain or inhibitor complexes thereof. Finally, in the example of stromelysin-1, potent inhibitors have been designed (e.g., 49, Figure 11 [55]) by N-terminal carboxyalkylamino functionalization that includes a P1 substituent. An x-ray crystallographic structure of a related MMP-3 inhibitor 125 shows (Figure 29) the hydrogen-bonding interactions at the active site and carboxylate coordination with the Zn2+ [223]. Finally, a recently determined x-ray crystallographic structure of an Pheψ[P=O(OH)CH2]Ala-modified peptide inhibitor 126 complexed with astacin (Figure 29) shows the extensive hydrogen-bonding network between inhibitor, enzyme, Zn2+, and a structural water [226]. It is expected that iterative structure-based design of inhibitors of the MMP family will enable the discovery of novel compounds with superior binding affinities and/or selectivities. C. Signal-Transduction Protein Targets Beyond proteases the opportunity for structure-based drug design is being realized in the rapidly developing area of signal-transduction research (e.g., intracellular protein and nucleic acid targets). Both x-ray crystallography and/or NMR spectroscopy have significantly contributed to a wide-scope database of 3D structural information for various catalytic and noncatalytic signal-transduction protein targets (see Table 3). These include tyrosine kinases (e.g., growth factor receptor kinases and Src family kinases; for reviews see Reference 231), serine/threonine and “dual specificity” kinases (e.g., mitogenactivated protein kinases and CDK2 and cAMP-dependent protein kinases; for reviews see Reference 232), phosphotyrosine phosphatases (e.g., PTP1B and Syp; for reviews see Reference 233), phosphoserine/phosphothreonine and “dual specificity” phosphatases (e.g., VH1 and CDC25; for reviews see Reference 234), noncatalytic “adapter” proteins (e.g., Crk, Grb2, Shc, and IRS-1; for reviews see Reference 85), transferases (e.g., Ras farnesyl transferase; for reviews see Reference 81), proline cis-trans isomerases (e.g., FKBP-12 and cyclophilin A; for reviews see Reference 235), and GTPbinding proteins (e.g., p21 Ras and α-β/γ heterotrimeric G-protein for GPCR superfamily; for reviews see References 236 and 237, respectively). The diversity of targets, mechanistic relationships (e.g., enzyme–substrate or regulatory protein–protein interaction), and potential therapeutic opportunities has created great impetus for focused research in the area of signal transduction (for reviews see References 265–267). Src Homology-2 and Homology-3 Domains The identification of noncatalytic regulatory domains referred to as Src homology (SH) domains has been rapidly advanced over recent years as a critical link

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Figure 29 Protease 3D structural models: matrix metallo protease-inhibitor complexes and drug design

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Page 613 Table 3 Some Known 3D Structures of Signal-Transduction Proteins (Apo/Complexes) Protein Target

Apo/Complex

Resolution

Reference

Src homology domains Abl SH2

—(Apo)

NMR

238

Src SH2

—(Apo)

2.5 Å

239

phosphopeptide

2.7 Å

239

phosphopeptide

NMR

240

PLC (C) SH2

phosphopeptide

NMR

241

Shc SH2

phosphopeptide

NMR

242

Lck SH2

phosphopeptide

2.2 Å

243a

p85 (N) SH2

phosphopeptide

2.0 Å

244

Syp SH2

phosphopeptide

2.0 Å

245

Grb2 SH2

phosphopeptide

2.1 Å

246

Syk SH2

phosphopeptide

NMR

247

Zap70 SH2-SH2

phosphopeptide (tandem)

1.9 Å

248

Src SH3

—(Apo)

NMR

249a

peptide

NMR

249b

Abl SH3

peptide

2.0 Å

250

Crk SH3

peptide

1.5 Å

251

Grb2 SH3

peptide

NMR

252

Grb2 SH3-SH2-SH3

—(Apo)

3.1 Å

253

Insulin receptor Tyr kinase

—(Apo)

2.1 Å

254

cAMP-dep. protein kinase

—(Apo)

3.9 Å

255

Tyr and Ser/Thr kinases

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(Ser/Thr)

peptide

2.9 Å

255

—(Apo)

2.4 Å

256a

protein (p27Kip1 inhibitory domain)

2.3 Å

256b

—(Apo)

2.3 Å

257

BH-PTP (Tyr)

—(Apo)

2.2 Å

258a

PTP1B (Tyr)

—(Apo)

2.8 Å

258b

peptide (C215S mutant enzyme)

2.6 Å

258c

VH1 (Tyr and Ser/Thr”)

—(Apo)

2.1 Å

259

PP-1 (Ser/Thr)

—(Apo)

2.1 Å

260

—(Apo)

1.95 Å

261

peptide

1.8 Å

261

peptide

NMR

262

Cyclophilin

peptide (cyclosporin-A)

2.8 Å

263

FKBP-12

FK-506 (macrolide antibiotic)

1.7 Å

264

Cell cycle-dep. protein kinase (CDK2) Mitogen-activated protein kinase (MAPK) Tyr and Ser/Thr phosphatases

pTyr binding domains IRS-1 PTB

IRS-1 PTB Pro cis-trans isomerases

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in deconvoluting both enzyme-substrate and regulatory protein-protein interactions for a number of signal-transduction pathways (for reviews see Reference 268). This emerging “superfamily” of proteins includes SH2 and SH3 domains, the so-called “choreographers of multiple signalling pathways,” and include very intriguing new therapeutic targets [85]. The SH2 domains have been determined to bind cognate phosphotyrosine (pTyr) containing proteins in a sequence-dependent manner relative to the amino acids contiguous to the C-terminal side of the pTyr residue (e.g., for Src SH2 a preferred sequence is ~pTyr-Glu-Glu-Ile~ versus ~pTyr-Tyr-Asn-Tyr for Grb2 [87,269]. The SH3 domains have been determined to specifically bind Pro-rich sequences of cognate proteins. Interestingly, as a result of the pseudosymmetrical nature of the SH3 domains there is the possibility of binding both N rarrow.gif C and C rarrow.gif N directions (e.g., for Src SH3 preferred sequences are ~Arg-Ala-Leu-Pro-Pro-LeuPro-Arg-Tyr and Ala-Phe-Ala-Pro-Pro-Leu-Pro-Arg-Arg, wherein Arg binds to a site 3 pocket [249b]). With respect to SH2 domain structure-based drug design, the first x-ray crystallographic structures of pTyr-containing peptide ligands complexed with Src SH2 domain [239] have been utilized to design the first peptidomimetic antagonists [89]. As illustrated in Figure 30, a molecular map of the tetrapeptide sequence ~pTyr-Glu-Glu-Ile~ complexed with Src SH2 [239] shows the pTyr binding pocket and a second binding site for the P+3 Ile residue. As previously described, a prototypic peptidomimetic AcpTyr-Glu-D-Hcy-NH2 (66) was first discovered by a peptide scaffold design strategy (see Figure 16; [77]) that took into account the x-ray crystallographic structure of Src SH2-phosphopeptide (Glu-ProGln-pTyr-Glu-Glu-Ile-Pro-Ile-Tyr-Leu, 127) complex. Further structure-based drug design modifications have led to the discovery a series of potent peptidomimetics having novel C-terminal functionalization (e.g., “transposed” side chain of the P+1 Glu or conformational constraint using a pyrrolidine ring; see Figure 30) as represented by 128–130 [89,270]. Studies focused on Src SH2 [88] have shown that the phosphate ester of pTyr is particularly critical for molecular recognition, and that significant loss in binding occurs by replacement with sulfate, carboxylate, nitrosyl, hydroxy, and amino. However, backbone modifications of pTyr which replace its acylated amino functionality with aromatic rings designed to form π-cation type interactions with the Arg-αA2 were effective substitutions [271]. Recently, high-resolution 3D structures have been described for the noncatalytic “adapter” protein Grb2 with respect to the apoprotein (SH3-SH2-SH3 [153]) as well as the individual SH2 and SH3 domains [246 and 252, respectively]. In the case of the SH2 domain of Grb2 an x-ray crystallographic structure of a phosphopeptide complex provided insight to the molecular basis of the specificity of Grb2 SH2 binding of ~pTyr-Xxx-Asn-Yyy~ sequences. As illustrated in Figure 31, the binding interactions of LysPro-Phe-pTyr-Val-Asn-

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Figure 30 SH2 domain 3D structural models: pp60Src SH2 domain antagonists

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Figure 31 SH2 domain 3D structural models: Grb2 SH2-SH3 domain antagonists

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Val (131) showed that the phophopeptide adopts a β-turn conformation about the P-P+3 residues and that the P+2 Asn side chain carboxamide moiety is extensively hydrogen bonded to the protein. In contrast to the well-defined binding pocket for the P+3 Ile of 127 to bind Src SH2, the P+3 Val of 131 engages in limited surface hydrophobic interactions because the Trp121 residue of Grb2 SH2 sterically blocks the phosphopeptide from attaining a similar binding mode. In the case of the SH3 domain, the binding of a cognate Pro-rich peptide sequence ~Pro-Pro-Pro-Val-Pro-Pro-Arg-Arg~ shows distinct pockets which recognize the Pro, Val, and Arg residues as illustrated in Figure 31. The peptide adopts a left-handed polyproline type-II helical conformation which projects the three aforementioned residues to their complementary binding pockets in the Grb2 SH3 domain. Tyrosine Phosphatases and Phosphotyrosine Binding Domains Two other types of signal-transduction proteins that recognize phosphotyrosine-containing sequences are tyrosine phosphatases (e.g., PTP1B, Syp, and CD45) and proteins that contain a noncatalytic motif referred to as a phosphotyrosine binding (PTB) domain. Although tyrosine phosphatases and PTB domains are structurally quite different from each other they both are similar with respect to the binding of pTyr-containing sequences with preference to the amino acids N-terminal to the pTyr residue. In other words, the tyrosine phosphatase PTP1 binds EGF receptor (pTyr992)-based phosphopeptide substrates Asp-Ala-Asp-Glu-pTyr-Leu-NH2 and Asp-Ala-Asp-Glu-pTyr-Leu-Ile-Pro-Gln-Gln-Gly equally [272], and the substitution of pTyr by nonhydrolyzable F2Pmp in the hexapeptide derivative has been reported [273] to give a highly potent inhibitor (see compound 69, Figure 14). In the case of PTB domains, ~Asn-Pro-Xxx-pTyr~ (where Xxx is variable) has been determined [274] as the cognate sequence for several PTB-domain-containing proteins such as Shc and the insulin receptor substrate-1 (IRS-1). The preference for amino acids N-terminal to the pTyr residue in binding to either tyrosine phosphatases or PTB domains is, therefore, opposite of that known for SH2 domains [275]. Among the tyrosine phosphatase superfamily, PTP1B was the first to be discovered and structurallydetermined by x-ray crystallography, as the apoprotein catalytic domain [258b]. The first x-ray crystallographic structure of PTP1B complexed with a phosphopeptide has also been very recently determined [258c] using a catalytically inactive Cys215 rarrow.gif Ser PTP1B mutant and the phosphopeptide Asp-Ala-Asp-Glu-pTyr-Leu-NH2 (133). As illustrated in Figure 32, the molecular interactions between the tyrosine phosphatase and the phosphopeptide are dominated by electrostatic (i.e., the pTyr, P-1 Glu, and P-2 Asp residues) and hydrogen bonding contacts to key amide functionalities of the backbone of 133. The P+1 Leu side chain forms hydrophobic contacts with

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Figure 32 PTP and PTB 3D structural models: PTP1B inhibitor and PTB antagonists

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several PTP1B residues at the surface proximate to the well-defined pTyr binding pocket. Such 3D structural information provides insight for the design of peptidomimetic inhibitors of PTP1B. In the case of the IRS-1 PTB domain, x-ray crystallographic studies of PTB complexed with a pTyrcontaining peptide (134) complex have shown (Figure 32) that the phosphopeptide forms a type-I β-turn within the Asn-Pro-Ala-pTyr sequence, and the peptide backbone is extensively hydrogen bonded to the PTB domain from the P+1-P-7 residues of 134 [261]. The pTyr binding pocket provides both electrostatic and multiple hydrogen bonding contacts to the phosphate ester moiety, and hydrophobic interactions exist for the P-1-P-3 side chains of the peptide. As in the case of PTP1B, such 3D structural information provides the opportunity for structure-based drug design to discover potent inhibitors which may be used for further exploration in cellular studies. V. Future Perspectives The impact of structure-based drug design on both peptidomimetic and nonpeptide drug discovery has been significant over the past few years. The integration of sophisticated computational chemical technologies, structural biology (x-ray crystallography and NMR spectroscopy), molecular diversity and high-through-put screening, and targeted biological testing are expected to provide invaluable guidance to drug discovery. These “technological tools” are contributing to an emerging “3D structure-activity database” that is the essence of rational drug design. Certainly, this intriguing area of peptidomimetic and nonpeptide drug discovery and design is providing tremendous insight to our understanding of molecular recognition and biochemical mechanisms. With particular regard to molecular diversity, the generation of new leads from combinatorial chemistry focused on synthetic peptide, peptidomimetic, or nonpeptide-type libraries will provide new opportunities (and challenges) for structure-based drug design strategies (for reviews see Reference 276). Novel scaffolds and templates will continue to be advanced and iteratively modified using “randomized” or “targeted” substructural replacements, and such work is exemplified in this review. Of particular significance to the field of peptidomimetic and nonpeptide drug discovery will be the rational use of D-amino acids, Nα-alkyl or Cα-alkyl amino acids, N-substituted Gly, XxxΨ[Z]Yyy (dipeptide isosteres), benzodiazepines, and amino-benzoic acids in structure-based drug design. Without question, structure-based drug design will be a decisive factor in drug discovery efforts ranging from de novo design to iterative structural optimization of peptidomimetic and nonpeptide lead compounds.

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231. (a) Sun H, Tongs NK. TIBS 1994; 19:480–485; (b) Fry MJ, Panayotou G, Booker GW, Waterfield MD, Protein Sci 1993; 2:1785–1797; (c) Pawson T. Curr Opinion Genet Develop 1992; 2:4–12; (d) Ulrich A, Schlessinger J. Cell 1990; 61:203–212. 232. (a) Taylor SS, Radzio-Andzelm Structure 1994; 2:345–355; (b) Taylor SS, Knighton DR, Zheng J, Sowadski JM, Gibbs CS, Zoller MJ. TIBS 1993; 18:84–89. 233. (a) Mauro LJ, Dixon JE. TIBS 1994; 19:152–155; (b) Feng G-S, Pawson T. Trends Genetics 1994; 10:54–58; (c) Fischer EH, Charbonneau H, Tonks NK. Science 1991; 253:401–406. 234. Shenolikar S. Ann Rev Cell Biol 1994; 10:55–86. 235. (a) O'Keefe SJ, O'Neill EA. Perspect Drug Discovery Des 1994; 2:57–84; (b) Fischer G. Angew Chem Int Ed Engl 1994; 33:1415–1436; (c) Armistead DM, Harding MW. Ann Rep Med Chem 1993; 28:207–215.

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236. (a) Marshall MS. TIBS 1993; 18:250–255; (b) Milburn MV, Tong L, DeVos AM, Brunger A, Yamaizumi Z, Nishimura S, Kim S-H. Science 1990; 247:939–945. 237. (a) Neer EJ. Protein Sci 1994; 3:3–14; (b) Simon MI, Strathmann MP, Gautam N. Science 1991; 252:802–808. 238. Overduin M, Rios CB, Mayer BJ, Baltimore D, Cowburn D. Cell 1992; 70:69–704. 239. Waksman G, Shoelson SE, Pant N, Cowburn D, Kuriyan J. Cell 1993 72:779–790. 240. Xu RX, Word JM, Davis DG, Rink MJ, Willard MJ Jr, Gampe RT Jr. Biochem 1995; 34:2107–2121. 241. Pascal SM, Singer AU, Gish G, Yamazaki T, Shoelson SE, Pawson T, Kay LE, Forman-Kay JD. Cell 1994 77:461–472. 242. Zhou M-M, Meadows RP, Logan TM, Yoon HS, Wade WS, Ravichandran KS, Burakoff SJ, Fesik SW. Proc Natl Acad Sci USA 1995; 92:7784–788. 243. (a) Eck MJ, Shoelson SE, Harrison SC. Nature 1993; 362:87–91; (b) Eck MJ, Atwell SK, Shoelson SE, Harrison SC. Nature 1994; 368:764–769; (c) Mikol V, Baumann G, Keller TH, Manning U, Zurini MGM. J Mol Biol 1995; 246:344–355; (d) Tong L, Warren TC, King J, Betageri R, Rose J, Jakes S. J Mol Biol 1996; 256:601–610. 244. Nolte RT, Eck MJ, Schlessinger J, Shoelson SE, Harrison SC. Nature (Struct Biol) 1996; 3:364–374. 245. Lee C-H, Kominos D, Jacques S, Margolis B, Schlessinger J, Shoelson SE, Kuriyan J. Structure 1994; 2:423–438. 246. Rahuel J, Gay B, Erdmann D, Strauss A, Garcia-Echeverria C, Furet P, Caravatti G, Fretz H, Schoepfer J, Grutter MG. Nature (Struct Biol) 1996; 3:586–589. 247. Narula SS, Yuan RW, Adams SE, Green OM, Green J, Philips TB, Zydowsky LD, Botfield MC, Hatada M, Laird ER, Zoller MJ, Karas JL, Dalgarno DC. Structure 1995; 3:1061–1073.

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248. Hatada MH, Lu X, Laird ER, Green J, Morgenstern JP, Lou M, Marr CS, Phillips TB, Ram MK, Theriault K, Zoller MJ, Karas JL. Nature 1995; 377:32–38. 249. (a) Yu H, Rosen MK, Shin TB, Seidel-Dugan C, Brugge JS, Schreiber SL. Science 1992; 258:1655–1668; (b) Feng S, Chen JK, Yu H, Simon JA, Schreiber SL. Science 1994; 266:1241–1247. 250. Musacchio A, Saraste M, Wilmann M. Nature (Struct Biol) 1994; 1:546–551. 251. Wu X, Knudsen B, Feller SM, Zheng J, Sali A, Cowburn D, Hanafusa H, Kuriyan J. Structure 1995; 3:215–226. 252. Goudreau N, Cornille F, Duchesne M, Parker F, Tocque B, Garbay C, Roques BP. Nature (Struct Biol) 1994; 1:898–907. 253. Maignan S, Guilloteau J-P, Fromage N, Arnoux B, Becquart J, Ducruix A. Science 1995; 268:291–293. 254. Hubbard SR, Wei L, Ellis L, Hendrickson WA. Nature 1994; 372:746–754. 255. (a) Zheng J, Knighton DR, Xuong H-H. Taylor SS, Sowadski JM, Ten Eyck LF. Protein Sci 1993; 2:1559–1573; (b) Karlsson R, Zheng J, Xuong N-H, Taylor SS, Sowadski JM. Acata Crystallogr 1993; D49:381–388. 256. (a) De Bondt HL, Rosenblatt J, Jancarik J, Jones HD, Morgan DO, Kim SH. Nature 1993; 363:595–02; (b) Russo AA, Jeffrey PD, Patten AK, Massague J, Pavletich NP. Nature 1996; 38:325–331.

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257. Zhang F, Strand A, Robbins D, Cobb MH, Goldsmith EJ. Nature 1994; 367:704–710.

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258. (a) Zhang M, Van Etten RL, Stauffacher CV. Biochem 1994; 33:11097–11105; (b) Barford D, Flint AJ, Tonks NK. Science 1995; 263:1397–1404; (c) Jia Z, Barford D, Flint AJ, Tonks NK. Science 1995; 268:1754–1758. 259. Yuvaniyama J, Denu JM, Dixon JE, Saper MA. Science 1996; 272:1328–1331. 260. Goldberg J, Huang H-B. Kwon Y-G, Greengard P, Nairn AC, Kuriyan J. Nature 1995; 376:745–753. 261. Eck MJ, Dhe-Paganon S, Trub T, Nolte RT, Shoelson SE. Cell 1996; 65:695–705. 262. Zhou M-M, Huang B, Olejniczak ET, Meadows RP, Shuker SB, Miyazaki M, Trub T, Shoelson SE, Fesik SW. Nature (Struct Biol) 1996; 3:388–393. 263. Pflugl G, Kallen J, Scuirmer T, Jansonius JN, Zurini MGM, Walkinshaw MD. Nature 1993; 361:91–94. 264. Van Duyne GD, Standaert RG, Karplus PM, Schreiber SL, Clardy J. Science 1991; 252:839–842. 265. Saltiel AR. Scientific American (Sci Med) 1995; 2:58–67. 266. Bridges AJ. Chemtracts (Org Chem) 1995; 8:73–107. 267. Huber HE, Koblan KS, Heimbrook DC. Curr Med Chem 1994; 1:13–34. 268. (a) Cohen GB, Ren R, Baltimore D. Cell 1995; 80:237–248; (b) Liu X, Pawson T. Recent Prog Hormone Res 49:149–160; (c) Fry MJ, Panayotou G, Booker GW, Waterfield MD. Protein Sci 1993; 2:1785–1797. 269. Cantley LC, Songyang Z. J Cell Sci 1994; 18 (Suppl):121–126. 270. Shahripour A, Para KS, Plummer MS, Lunney EA, Stankovic CJ, Holland DR, Rubin JR, Humblet C, Fergus JH, Marks JS, Saltiel AR, Sawyer TK. Bioorg Med Chem 1997; in press.. 271. Shahripour A, Plummer MS, Lunney EA, Para KS, Stankovic CJ, Rubin JR, Humblet C, Fergus JH, Marks JS, Herrera R, Hubbell SE, Saltiel AR, Sawyer TK. Bioorg Med Chem Lett 1996; 6:1209–1214. 272. Zhang Y-Z, Maclean D, McNamara DJ, Sawyer TK, Dixon JE, Biochemistry 1994; 33:2285–2290. 273. Burke TR Jr, Kole HK, Roller PP. Biochem Biophys Res Comm 1994; 204:129–134.

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274. (a) Kavanaugh WM, Williams LT. Science 1994; 266:1862–1865; (b) Kavanaugh WM, Turck CW, Williams LT. Science 1995; 268:1177–1179. 275. Eck MJ. Structure 1995; 3:421–424. 276. (a) Thompson LA, Ellman JA. Chem Rev 1996; 96:555–600; (b) Patel DV, Gordon EM. Drug Disc Today 1996; 1:134–144; (c) Chen JK, Schreiber SL. Angew Chem Int Ed Engl 1995; 34:953–969; (d) Andrews P, Cody WL, Leonard DM, Sawyer TK. In: Pennington MW, Dunn BM, Eds. Peptide Synthesis and Purification Protocols. Vol. II. Humana Press, 1994:305–328; (e) Zuckermann RN. Curr Opinion Struct Biol 1993; 3:580–584; (f) Pavia MR, Sawyer TK, Moos WH. Bioorg Med Chem Lett 1993; 3:387–396; (g) Jung G, Beck-Sickinger AG. Angew Chem Int Ed Engl 1992; 31:367–486.

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Index A Acquired immunodeficiency virus (AIDS), 41, 56, 62, 65, 70 Activate platelets, 247 Active site, 45, 48, 50, 54, 55, 56, 59, 61, 62, 63, 64, 65 Activity, 41, 48, 50, 55, 56, 64, 65, 67, 68, 69, 70 Acyclovir diphosphate, 154, 160, 164 Addison's disease, 195 Alcohol dehydrogenase (ADH) 202-203 Aldo-keto reductase superfamily, 240 Aldose reductase (ALR2) catalytic mechanism, 233-234 inhibitors, 231-232 mutations, 234 NADPH cofactor binding, 232-233 relation to diabetic complications, 229-231 structure active site, 233-235 NADPH-bound form, 231-233 ternary complex with zopolrestat, 235-239 α-ketoamide transition state, 282 http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_635.html (1 of 2) [4/9/2004 1:50:42 AM]

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ALR2 (see Aldose reductase) Amantidine, 462 Amide bond replacements, 563-565 1-amidinopiperidine, 256, 257 Amino acids C alpha, 128 N-methyl, 128 side chain constraints, 124-128 Aminoalcohols 326 Amphipathic alpha-helices (see Leucine zipper) Angiotensin, 321 converting enzyme, 120 Animal models thrombosis of, 267 Anthopleurins, 297, 300, 301, 302, 312, 313, 314 Antibody-neuraminidase complexes, 470 Anticoagulation, 247 Anticoagulant protein, 257

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Anti-influenza drugs, 462, 477-480 Antirhinoviral agents capsid-binding compounds, 497-500 clinical trials, 517-518 drug sensitivity groups, 503 resistance, 514-517 structure-activity relationships, 502-514 WIN compounds, 498-500 Antithrombin III, 248 Antitrypanosomal agents eflornithine, 365-367 melarsoprol, 365-366 pentamidine, 365 suramin, 365-366 Antiviral agent, 41, 67 Apparent mineralocorticoid excess (AME), 191 Argatroban, 251, 255 Arginine boronate esters, 250 guanidinium, 251 Arrhythmias, 296, 298 Aspartic proteinases http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_636.html (1 of 4) [4/9/2004 1:52:05 AM]

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inhibitor binding, 323, 332 inhibitors, 323 mechanism, 328 specificity, 333 structure, 322 transition state analogs, 323 Atherosclerosis 395, 398 Autoimmune disease, 395,398 atherosclerosis, 395, 398 insulin-dependant diabetes, 395, 398 myasthenia gravis, 398 rheumatoid arthritis, 395, 398 systemic lupus erythematosus, 398 disorders, 152 Available chemicals database (ACD), 381 B Bacteriorhodopsin, 131 BCX-34, 166, 167 Benzodiazepinone peptidomimetic, 571 β-barrels, 248 β-turn, 122, 124, 126, 127

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Bicyclic peptidomimetic, 251 Bidentate hydrogen bond, 252 Binding cleft, 45, 48, 61, 65 pocket, 56, 58, 60 Bivalent inhibitors, 257 Blood clot formation, 247 Boroarginine, 261 Boronate esters, 251 Boronic acid analog, 250 Bovine trypsin inhibitor mutants phage display, 287 positional requirements for Fxa inhibition, 286-288 random mutagenesis, 287 site specific mutagenesis, 285 structure of complexes, 285 C Calcium regulators, 296 cAMP generators, 296 Cancer, 395 Carbonyl reductase, 199 Cardiotonic activity, 301, 304, 305, 306, 314 Catalytic

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site, 52, 55, 61 triad, 247, 275 Catechol O-methyltransferase active site, 349-350, 355-356 catalysis, 345, 350-351 inhibition mechanism, 356-358 inhibitors, 351-353 kinetics, 346 physiological role, 344-345 structure AdoMet (see S-Adenosyl-L-methionine) crystal structure, 347-350

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[Catechol O-methyltransferase] drug complexes, 354-355 Mg++ ion, 349-350 S-adenosyl-L-methionine, 349 sequence, 345, 347 substrates, 345,346 Cation-p site, 277, 282 Ceriamide, 473 Charge calculation, 381 CHARMm, 133 Chemical library, 142 Chemi-informatics, 526, 535, 537 databases, 536 selection algorithm, 536 systems integration, 536 virtual library production, 536 Chemotherapy, 66 Chimeric receptor, 126 Chymotrypsin serine protease family, 248 Circular dichroism (CD), 438, 444 Coagulation cascade, 247, 265

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factors, 247 Combination therapy, 62, 68, 69 Combinatorial approach, 550-552 chemistry, 141, 525, 537 combinatorial explosion, 552 drug lead source, 528, 529 focusing, 530 parallel synthesis, 528, 532 refinement, 530 robotic instrumentation, 528, 532 scaffolds, 530, 532 structure-activity relationship (SAR), 529 pruning, 552 Compound selection, 531 chemi-informatics, 536 drug properties, 535 receptor fit, 536 similarity, 536 structure-activity relationship (SAR) models, 533, 536 virtual libraries, 533, 536 Computer programs BIOGRAF, 381

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CHARMM, 542-544 computational combinatorial ligand design (CCLD), 550-552 DELPHI, 383 DOCK, 379-383 MCSS, 542-544 Conformational change, 59, 60, 61, 70 degrees of freedom, 252 protein, 235-237 Congestive heart failure, 295, 296, 298, 301, 314 Cortiso, 193 Cortisone, 193 Cross linking, 137-138 Crystallography cocktail soak approach, 377-379 Crystal structure, 45, 54, 56, 59, 61, 64 aldose reductase, 231-239 other aldo-keto reductases, 240-241 glyceraldehyde-3-phosphate dehydrogenase, 374-375 phosphoglycerate kinase, 376-377, 388 triosephosphate isomerase, 371-373 Cutaneous T-cell lymphoma, 167 Cyclic template, 252

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Cyclotheonamide A (CtA), 254

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Cytokines, 395-398, 412, 420 alpha-beta, 396 atherosclerosis, 398 autoimmune disease and, 395, 98 beta sandwich 396 beta strands, 402 beta-trefoils, 396 4-helix bundle, 396 growth hormone, 398 immune system, 398 immunomodulation, 398 insulin-dependant diabetes, 398 myasthenia gravis, 398 network, 396 nuclear magnetic resonance (NMR) and, 396 production of, 395 prolactin, 398 receptors, 396 rheumatoid arthritis, 398 signalling by, 396 structures, 396 systemic lupus erythematosus, 398

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tumour necrosis factor, 398 x-ray crystallography and, 396 D ddI, 152 9-deazaguanine, 161, 163, 164, 165, 167 Diabetic complications, 229-231 Digoxin,295, 296, 297, 300 Dihydropteridine reductase, 197, 202 Diversity directed, 536 of candidate ligands, 542,555 dNTP binding, 56, 61 Drug, 43, 45, 53, 55, 56, 60, 62, 63, 65, 66, 67, 68, 69, 70 binding affinity prediction, 555 combinatorial, 550-552 design, 55, 66, 70, 402 docking, 241-242, 379-383, 542-544 fragment approach, 541 inhibition, 70 lead discovery, 377-384 optimization, 384-387 linked-fragment approach, 378

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resistance, 56, 66, 68, 70 scoring, 382-383 solvation effects, 544-547 Drug properties, 527, 535 absorption, 535 excretion, 535 metabolism, 535 refinement, 535, 536, 537 structure-based design, 527 DTic (tetrahydroisoquinnoline carboxylic acid), 124 DUP714, 250 E Energy minimization, 132-133 with CHARMM, 542-544 F Fab, 45, 49, 51, 56, 58, 59, 69 Factor Xa active site blocked (DEGR-Xa), 267, 269 active site substrate sequences, 271 modeled inhibitor complexes amidinoaryls, 279

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antistasin peptides, 281 cyclotheonamide, 284 dansyl-Glu-Glu-Arg-CMK, 280 DX-9065a, 276 SEL-2711, 283 natural inhibitors of AcAP's, 273-274 antistasin, 266, 268, 270, 271-272 ecotin, 268, 270, 274, 288 tick anticoagulant peptide (TAP), 266, 268, 270, 272-273

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[Factor Xa] TFPI, 270-271, 285, 287 structure and function, 267-269, 274 synthetic inhibitors of amidinoaryls, 277-279 antistasin peptides, 280-282 bisamidines, 277 BPTI mutants, 285-288 cyclotheonamide, 282 dansyl-Glu-Glu-Arg-CMK, 280 DX-9065a, 275-277 peptidyl argininals, 288 PPACK, 275 SEL-2711, 282 Factor XIII, 247 Factor XIIa, 119 Feline immunodeficiency virus (FIV), 441 Fibrin, 247 Fibrinogen, 247 Fibrinopeptide A, 250, 261 Flavonoid, 473 Fluoroketone analogs, 327 Fragment approach (see Computer programs, MCSS, CCLD)

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fragment-based programs, 541-542 Fourier maps, 154, 161, 166 G Geminal diol analogs, 327 Genomic data, 525, 527 GG167 (see Neu5Ac2en,4-guanidino) Glucopyranoside peptidomimetic, 571 Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) catalysis, 372, 374 crystal structure, 374-375 human, 374-375 inhibitors, 381-382, 384-387 Leishmania mexicana, 375-376, 385 sequence, 374 Glycol analogues 326 Glucocorticoid 193 G-protein coupled receptors (GPCR), 592-594 GRID maps, 475 H Heart attack, 247 Hemagglutinin, 459, 460, 462, 463, 464, 477 Hematophageous organisms

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Ancylostoma caninum, 273 Haementeria officinalis, 266 Ornithidorous moubata, 266 Hemopexin domain, 172-174 Hemostasis, 247 Heparin, 248 cofactor II, 248 High throughput screening, 530 Hirudin, 251 HOE 140, 127-128 Homology model building, 275-276,278 models, 527 scaffold development, 532 structure-based design, 527 sequence, 176 structure, 176-177 Hormone therapy, 206 Human immunodeficiency virus (HIV), 441 protease cleavage sites, 7 flexibility, 7-8 inhibitors, 586-587 AG1284, 22-27

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Page 640

[Human immunodeficiency virus] cyclic ureas, 21-22 hydroxycoumarins, 27-28 indinavir, 9, 15-17, 28-29 inversion of binding mode, 24 nelfinavir, 17-21, 29-30 nonpeptidic, 17-28 peptidic, 9-10 peptidomimetic, 10-19 ritonavir, 13-16, 28-29 saquinavir, 10-13, 28-29 symmetric binding, 13-15, 21-22 mutations, 28-32 resistance primary mutations, 29-30 secondary mutations, 30-32 sequential passage, 28 vitality factor, 32 structure active site, 5-7 three dimensional, 3-5 Wat 301, 6, 7, 10, 13, 16, 17, 21, 22, 24, 27

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integrase amino acid sequence, 90-91 amino terminal domain, 92, 102 biochemical properties, 85-88 biophysical properties, 88, 92 carboxyl terminal domain, 89, 92 structure of, 102-103 catalytic core domain, 88-89 biophysical properties of, 92-93 mutation of hydrophobic residues of, 102-103 structure of, 93-102 comparison to toher polynucleotidyl transferases, 96-100 conserved acidic residues in active site, 95-96 dimer, 100-102 domain structure, 88-92 inhibitors, 103-109 3'-azido-3'-deoxy-thymidine (AZT), 107-108 common pharmacophore of, 104-107 curcumin, 107, 111 design of, 110-112

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overview, 103-104, 108-109 rationale for, 85 Human prothrombin fragment F1, 261 Hydrophobic collapse, 252 Hydroxamate, 172, 182-184 Hydroxysteroid dehydrogenases, 191 7α, 197, 199 11β, 191-194, 203 17β, 193-199, 205-207 20β, 197, 199 Hypertension, 193 I Inflammation, 395 Influenza virus antigenic variation, 462, 468-470 classification, 459 drug resistance, 478, 479 inhibition, 478 replication cycle, 461 vaccines, 463 Inhibition, 50, 54, 60, 61, 65, 69, 70 mechanism, 65 Inhibitors, 41, 43, 45, 48, 50, 55, 56, 58, 59, 60, 61, 62, 63, 65, 66, 67, 68, 69, 70 http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_640.html (3 of 4) [4/9/2004 1:56:12 AM]

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design, 358-359

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[Inhibitors] 2-((3,4-dihydroxy-2-nitrophenyl)vinyl)phenyl-ketone, 352-353, 356 entacapone, 352-353, 360 first generation inhibitors, 351-352 tolcapone, 352-353, 360 Insulin-dependent diabetes 395,398 Interactions electrostatic, 544 hydrophobic, 545 van der Waals, 545 Interferon α, 435, 439, 440 activity, 442 cytotoxicity, 439, 443 receptor, 441 β, 435 structural studies, 443, 444 clinical uses, 436 definition, 435 γ, 435 activity, 448, 449 antibody studies, 445

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receptor, 446, 447, 450 intron A, 436 roferon, 436 side effects, 436 sub types, 435, 436 synthetic peptide studies, 448 signal transduction, 451 structural studies, 449-451 synthetic peptide studies, 446, 448 τ, 435, 439, 440, activity, 441-443 antibody studies, 442 expression, 442, 443 receptor, 441 signal transduction, 443 structural studies, 443, 44 synthetic peptide studies, 441, 442, 444 ω, 435, 440 Interleukin-1 accessory protein, 398, 401 affinity for receptors, 401 α, 398, 399, 401, 402, 406, 420, 421, 423, 427, 428 alternatively spliced, 399

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antagonistic activity, 416 auto-antibodies and, 401 autoimmune disease and, 401 barrel, 402, 404 β, 398, 399, 401, 402, 420, 421, 423, 427, 428 barrel, 406, 409 bulge, 415-416 converting enzyme (ICE), 399, 402, 412 hairpins, 405 strands and, 402 beta-trefoil fold, 402, 403, 404, 407 binding, 412-416 affinity, 421 epitope, 410 pocket, 412 catalytic activity of, 412 diad, 412 cysteine protease, 399, 412 enzyme mechanism, 412 epitopes of, 405, 413-416, 418 expression of, 399 fibroblast growth factors, 402 http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_641.html (3 of 4) [4/9/2004 1:56:34 AM]

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gene duplication, 398 glycosylation of, 399, 409 GTPase proteins and, 401

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[Interleukin-1] homology with CED-3 protein, 412 hydrophobic patch, 406 hydrophobic residues, 404 immunoglobulin superfamily and, 399 Kunitz family, 402 leukemia, 395 location of, 399 low molecular weight antagonists, 421-427 monoclonal antibodies and, 421 monocyte phagocytes by, 399 mutational studies of, 407, 409 N-terminal extension nuclear magnetic resonance (NMR), 402, 404 overexpression of, 419 peptide fragments, 416 peptomimetics, 412 precursor, 399 form, 399 receptor, 398, 402, 420 antagonist, 398, 399, 401, 407, 420, 421 binding, 404-405, 406, 407-410, 412-416

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recombinant, 421 sequence identity of, 398 signal transduction and, 401 site-directed mutagenesis, 412-416 structure of, 401-412, 419 synthesis of, 395, 419 substrate, 399 specificity, 412 subunits of, 412 systemic lupus erythematosus, 398 therapeutic strategies and, 412 tumor necrosis factor and, (TNF) 421 type I, 399, 401, 420 type 2, 399, 401, 420 x-ray crystallography, 401-412 Ion-channel modulators, 296, 297, 301 Ischemia, 296, 297 K Kampo drug, 473 Ketomethylene pseudo peptide bond, 260 Kinins, 119 turn propensity of, 123 bradykinin, 119 http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_642.html (2 of 4) [4/9/2004 1:57:29 AM]

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Kininogen, 119 high molecular weight, 119 low molecular weight, 119 Kinin receptors, 120 agonist binding site, 131-133 antagonists of, 124 antagonist site, 137-138 B1 subtype, 120 B2 subtype, 119, 120 chimeras and, 138, 139 mutagenesis of, 133-134 Kallidin, 119 molecular dynamics of, 123 pharmacology of, 121 solution conformation of, 121 Kunitz domains, 271, 273, 282, 285 Kyte-Doolittle, 131 L Lactam amide, 222-223 Leucine zipper propane functional map, 545-546 stereo view, 546

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yeast transcriptional activator protein GCN4, 545-546 Leydig cells, 193 Licorice, 193, 195-196 Ligand docking, 133 Lock-and-key, 159 M Malignant tissue, 214 transformation, 214

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Matrix-metalloproteinase (MMP), 171-186 fibroblast collagenase, 171-173, 176, 179, 183-184 gelatinase, 171-173, 184 matrilysin, 171-175, 182-184 neutrophil collagenase, 171-175, 183-184 stromelysin, 171-173, 184 Medicinal leech, 257 Metal requirement, 88, 89 Mineralocorticoid receptor, 193 Molecular fragments aliphatic, 542 aromatic, 542 charged, 542 polar, 542 modelling, 182-183 MuA transposase structural comparison to integrase, 97-100 Mutation, 62, 67, 68, 69 Myristic acid, 214 Myocardial infarction, 247 N

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NAD, 199, 201 NADP, 199, 201 NADPH, 199-201 NAPAP, 255, 261 Napthalenesulfonyl, 257 Neuraminidase active site, 470-472 carbohydrate structure, 467 dimensions, 465 enzyme function, 464, 473 inhibitors, 472-473, 476 molecular weight, 465 morphology, 464 topology, 466 Neu5Ac2en, 472, 473, 474, 477, 478, 480 4-guanidino, 475, 476, 477, 479 4-amino, 475, 476 Nonnecleoside ingibitor binding pocket (NNIBP), 56, 58, 59, 60, 61, 62, 63, 66, 67 Nonnucleoside reverse transcriptase inhibitors (NNRTI), 41, 45, 48, 49, 50, 56, 58, 59, 60, 61, 62, 63, 64, 65, 66, 67, 68, 69, 70 Nonnucleoside, 41, 45, 56 inhibitor, 45, 56 inhibitor binding pocket, 56

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RT inhibitor, 41, 56 Nonpeptide angiotensin agonist, 586-587 antagonist angiotensin, 586-587 cholecystokinin, 586-587 endothelin, 586-587 gastrin-releasing hormone, 586-587 glucagon, 586-587 neurokinin, 586-587 neuropeptide Y, 588-589 neurotensin, 586-587 oxytocin, 586-587 drug discovery, 559, 570, 586-587 vasopressin, 586-587 NPC 17731, 124 NPC 18325, 137 NPC 567, 124 Nuclear magnetic resonance (NMR), 402, 404, 407 Nucleic acid, 48, 51, 54, 64, 65, 69 Nucleophile, 221 Nucleoside, 41, 50, 52, 53, 56 reverse transcriptase (RT) inhibitor, 41, 50, 52, 53, 54, 55, 56, 65

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Nucleotide, 51, 53, 54, 55, 69 binding domain, 200 O Octahydroindole carboxylic acid (Oic), 124 Oxyanion hole, 248, 250 P Parkinson's disease disease, 359 therapy, 359-360 Peptides synthetic structure-function studies in, 437, 438, 440-442, 444, 446, 448 Peptidomimetic drug discovery, 591-598, 613 (see also Protease targets, Receptor targets, Signal transduction protein targets, Structure-based drug design) Peptidomimetics, 251 PGK (see Phosphoglycerate kinase) Pharmacophore, 225, 560 peptide and nonpeptide models, 593-594 Phenanthroline copper complexes as integrase inhibitors, 107 http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_644.html (1 of 4) [4/9/2004 1:59:33 AM]

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Phosphoglycerate kinase (PGK) catalysis, 376-377 crystal structure, 376-377, 388 human, 377 sequence, 377 Trypanosoma brucei, 376-377 Phosphorylysis, 151 Phosphostatine analogs, 327 Phosphotransfer, 222 Platelet aggregation, 247 Polymerase, 41, 45, 48, 50, 52, 54, 55, 56, 59, 61, 62, 63, 64, 65, 67, 69 active site, 45, 48, 50, 54, 55, 56, 59, 61, 62, 63, 64 catalytic site, 52, 55, 61 Polymerization, 41, 48, 50, 55, 61, 67, 70 mechanism, 56 Polynucleotidyl transferases, 96-100 Positive inotropes, 295, 296, 297, 298, 300, 309, 310, 314 inotropy, 295, 297, 298, 305, 314 PPACK, 250, 261 Primer grip, 48, 61 Proliferative diseases asthma, 214 http://legacy.netlibrary.com/nlreader/nlReader.dll?bookid=12640&filename=Page_644.html (2 of 4) [4/9/2004 1:59:33 AM]

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atherosclerosis, 214 fibrosis, 214 osteo arthritis, 214 psoriasis, 214 restenosis, 214 rheumatoid arthritis, 214 septic shock, 214 Protein C, 247 Protease targets angiotensin-converting enzyme (ACE) inhibitors, 576, 607, 610 aspartyl proteases, 595-596 classes, 567-568 cysteinyl proteases, 605 inhibitors elastase, 576, 597, 606 gelatinase, 576 human immunodeficiency virus (HIV) protease, 576-578, 595-596, 598, 600-602 interleukin-converting enzyme (ICE), 576, 597, 607-608 stromelysin, 576, 596, 609-612 thermolysin, 597, 610 thrombin, 576, 578-579, 596, 604 metalloproteases, 597-598, 607, 609-612

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[Protease targets] serinyl proteases, 603 x-ray structures apoprotein and complexes, 595-598 Protein data bank, 173 Protein kinase activation loop, 218, 219 calmodulin dependent, 219 caseine (CK-1), 218, 220, 222, 224 catalytic core, 214, 219-221 loop, 217, 221 cyclin dependent (CDK-2), 214, 215, 218-219 EGFR, 223 insulin receptor (IRK), 214-216, 218-220, 222 MAP, 214, 218-219 phosphorylase, 218-220, 222, 224 protein A (cAPK) 214-216, 218-220, 222-225 protein C, 224 phosphorylation, 217, 218

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