Caspase Allostery and Conformational Selection - American Chemical

Review pubs.acs.org/CR

Caspase Allostery and Conformational Selection A. Clay Clark* Department of Biology, University of Texas at Arlington, Arlington, Texas 76019, United States ABSTRACT: The role of caspase proteases in regulated processes such as apoptosis and inflammation has been studied for more than two decades, and the activation cascades are known in detail. Apoptotic caspases also are utilized in critical developmental processes, although it is not known how cells maintain the exquisite control over caspase activity in order to retain subthreshold levels required for a particular adaptive response while preventing entry into apoptosis. In addition to active site-directed inhibitors, caspase activity is modulated by post-translational modifications or metal binding to allosteric sites on the enzyme, which stabilize inactive states in the conformational ensemble. This review provides a comprehensive global view of the complex conformational landscape of caspases and mechanisms used to select states in the ensemble. The caspase structural database provides considerable detail on the active and inactive conformations in the ensemble, which provide the cell multiple opportunities to fine tune caspase activity. In contrast, the current database on caspase modifications is largely incomplete and thus provides only a low-resolution picture of global allosteric communications and their effects on the conformational landscape. In recent years, allosteric control has been utilized in the design of small drug compounds or other allosteric effectors to modulate caspase activity.

CONTENTS 1. Introduction 2. CaspasesAn Overview 2.1. Caspase MEROPs Designation and Enzyme Mechanism 2.2. Caspase Classifications and Domain Organization 2.3. Features in the Caspase−Hemoglobinase Fold 2.3.1. Pro-Domain LinkerEvolutionary Leftovers or Functional Evolution? 2.3.2. Turn 6 Provides Flexibility near the Active Site 2.3.4. Variations in Active Site Loops L1 and L4 2.3.5. Maturation May Result in Active Site Formation 3. Allostery 3.1. Introduction to Ligand Binding Models and Conformational Space 3.2. Conformational Space of Caspases 3.2.1. Global Free Energy Diagram of the Caspase Family 3.2.2. Caveats Concerning the Description of Conformational Selection in Caspase Ensembles Using the Caspase Structural Database 3.2.3. Monomers, Dimers, Heterodimers, and the Formation of an Active Enzyme 4. Complex Conformational Space and Allosteric Regulation 4.1. Introduction 4.2. Assembly Processes of the Zymogen Monomer © 2016 American Chemical Society

4.2.1. Short Pro-Domain Caspases Are Constitutive Dimers 4.2.2. Long Pro-Domain Caspases Form Homo- or Hetero-Oligomers 4.3. Conformational Selection in the Caspase Dimer 4.3.1. DARPins Inhibit Caspases but Not Always Allosterically 4.3.2. Pro-Domain Linker Affects Caspase Function 4.3.3. Allosterically Inhibiting Caspases through the Closed Conformation 4.3.4. Caspase-6 Expands Conformational Space and Provides More Ways To Allosterically Inhibit the Enzyme 4.3.5. Post-Translational Modifications and Potential Long Distance Communications 5. Conclusion Author Information Corresponding Author Notes Biography Acknowledgments References

6666 6667 6667 6668 6669 6669 6670 6672 6672 6675 6675 6680 6680

6683 6687 6689 6689 6690 6691

6692

6695 6699 6700 6700 6700 6700 6701 6701

6682 6682 6683 6683

Special Issue: Protein Ensembles and Allostery

6683

Received: September 14, 2015 Published: January 11, 2016 6666

DOI: 10.1021/acs.chemrev.5b00540 Chem. Rev. 2016, 116, 6666−6706

Chemical Reviews

Review

1. INTRODUCTION

described as below or above an as-yet-undefined threshold level required for cell death (Figure 1). Under low levels of activation, the “adaptive responses”, caspase activity from the apoptotic subclass is important for a variety of reactions, from cell migration and invasion, proliferation, differentiation, embryonic development, and innate immunity.11,16−21 At higher levels of activation, above the threshold, caspase activity is important for carrying out apoptosis. While the role of caspases in apoptosis is well known, their roles in the adaptive responses are less clear. Importantly, it is not known how cells maintain the exquisite control over caspase activity in order to retain the subthreshold levels of enzyme activity required for a particular adaptive response while preventing entry into apoptosis, although, broadly, two general mechanisms affect the levels of activated caspases. First, signaling pathways modulate the processing of the caspase zymogen, thereby increasing or decreasing the levels of mature, active caspase. Second, endogenous inhibitors bind to active caspases and either inhibit activity or target the enzyme for degradation by the proteasome. A third mechanism that involves post-translational modifications of caspases is becoming increasingly recognized as an effective method to fine tune caspase activity. Signaling pathways,15,22,23 endogenous inhibitors,24−27 and post-translational modifications28−31 have been described in detail, so this review will focus on those reactions that affect caspase conformational selection in the native ensemble. In parallel to the more than two decades of cell biological studies of caspase function in cell death and differentiation are biochemical and structural studies of caspases, with the first structures published in 1994.32,33 As a result, there is a large database consisting of several hundred structures of caspases, complexes of caspases with effector proteins or ligands, or isolated domains of caspases. The structures have been the subject of several excellent reviews,34−38 so selected structures will be highlighted here as examples of potential mechanisms for conformational selection and regulation of caspase ensembles in the cell. Finally, recent studies of several caspases39−43 have identified allosteric sites on the enzyme that bind small molecule effectors. In addition, caspases are modified post-translationally by phosphorylation28 or glutathionylation31,44 at regions away from the active site. Although it was known that zinc inhibits caspase activity,45 recent studies show that in addition to modifying the catalytic cysteine, zinc also functions allosterically in caspase-6 by stabilizing an inactive conformation of the enzyme.46 Overall, studies of caspase activation, post-translational modifications, and allosteric ligand binding show that the caspase native state is a complex ensemble of conformations in which the binding of ligands affects the population distribution of states within the ensemble. This review provides overviews of caspase structural organization and maturation (section 2) and allostery (section 3) and describes the ensemble of states, conformational selection by protein and ligand effectors, and potential mechanisms in which conformational selection may be utilized by cells to fine tune caspase activity (section 4).

It is estimated that 1010 cells are produced each day in a healthy adult human. In order to maintain homeostasis, the same number of cells is removed, and this occurs primarily by apoptosis or autophagy.1−3 Apoptosis is a program of regulated cell suicide that is carried out by caspases, a family of cysteinyl aspartate-specific proteases. More than two decades of cell biological, biochemical, and structural research have shown that all caspases are produced initially as inactive zymogens (procaspases) that must undergo processing, which usually involves dimerization and proteolytic cleavage, to yield the active protease. Caspase activation is tightly controlled, and the activation cascades are known in quite some detail,4−15 so they will be summarized briefly in this review in order to further describe conformational selection and allosteric regulation of the caspase native ensemble. One should note that while all caspases are described in this review, apoptotic caspases are a

Figure 1. Caspase activity is utilized in adaptive responses of cell development and differentiation. Above as-yet-undefined threshold levels, caspase activity is utilized in the cell death program. Levels of activity below the threshold required for apoptosis are not defined, so at present it is not clear where the cellular processes (green boxes) fall on the continuum of caspase activity.

primary focus due to the extensive structural database on this subclass of caspases. Studies in recent years have shown that, in addition to their roles in apoptosis, caspases are utilized by cells for a number of critical development and differentiation processes, and indeed, caspase function in cell development can be considered a continuum of activity in the cell, where activity is generally

2. CASPASESAN OVERVIEW 2.1. Caspase MEROPs Designation and Enzyme Mechanism

Caspases47 are classified in the MEROPs peptidase database48,49 as family C14A, Clan CD proteases, where “C” refers to cysteinyl proteases and “D” refers to families that contain 6667


Chemical Reviews

Review

some,61−66 or the PIDDosome,67 in so-called extrinsic or intrinsic activation pathways.68 In the extrinsic pathway, activated death receptors (bound by external ligand) form a scaffold that binds initiator procaspases through the DED (death effector domain) motifs in the prodomain.60,69−71 Binding to the scaffold results in dimerization and subsequent activation of the procaspase.7,72 In the intrinsic pathway, cellular response to various stimuli (cytotoxic drugs, DNA damage, reactive oxygen species (ROS)) results in activation of procaspase-9 through formation of the apoptosome, a multiprotein complex that is dependent on the binding of cytochrome c, upon its release from the mitochondria, to Apaf-1 and the CARD (caspase recruitment domain) of

Figure 2. Classical cysteine protease reaction mechanism.

similar fold or sequences to the caspases. The C14 family also contains metacaspases (C14B(M))50,51 and paracaspases (C14B(P)).50,52,53 Caspases utilize a histidine−cysteine catalytic dyad to hydrolyze peptide bonds after specific aspartate residues.54 The classic reaction mechanism (Figure 2) shows that the catalytic cysteine acts as the nucleophile following proton abstraction by the catalytic histidine (Figure 2, step 1). After formation of a covalent tetrahedral intermediate between the nucleophile and the peptide substrate, the histidine donates a proton to the amino moiety of the peptide leaving group (Figure 2, step 2). Following cleavage of the peptide bond, hydrolysis results in the release of the N-terminal part of the peptide and reprotonation of the catalytic histidine (Figure 2, step 3), and the covalent adduct is resolved by reprotonation of the catalytic cysteine (Figure 2, step 4). Formation of an oxyanion hole by backbone nitrogen atoms is critical for polarizing and stabilizing the scissile carbonyl group.55 In more recent analyses, molecular dynamics56 and computational55 studies of the enzyme mechanism have suggested a somewhat different reaction scheme for proton transfer in that a proton is first transferred internally from the backbone nitrogen of the P1 aspartate residue to the carboxylate of the P1 aspartate, followed by transfer to a water molecule and then to the catalytic histidine. The catalytic cysteine then donates a proton to the P1 aspartate backbone nitrogen. The overall result is the same as the classical view of the enzyme mechanism, that is, the catalytic histidine is protonated while the catalytic cysteine is deprotonated.

Figure 3. Domain organization and substrate sequence specificity of human caspases.

caspase-9.62−65,73 Also, the initiator procaspase-2 is activated upon formation of the PIDDosome, which is responsive to metabolic changes in the cell.29,36,67 The activated initiator caspases in turn activate the effector procaspase-3, which is then transformed into fully active, mature caspase-3, the executioner of apoptosis. All caspases recognize a tetrapeptide sequence, with a requirement for an aspartate residue at the P1 position (Figure 3).74 The exception is caspase-2, which demonstrates a preference for an additional residue at the P5 position.74,75 The S1 binding pocket forms a cavity in the surface of the binding groove that accommodates the P1 aspartate (Figure 4), and because the requirement for aspartate at P1 is conserved in caspases, enzyme specificity is determined primarily by the amino acid preference in the P4 position (Figure 3). Thus, based on substrate preference, caspases are classified as Group I (caspases-1, -4, -5, -14: preference of (W/L)EHD), Group II (caspases-2, -3, -7: preference of DEXD), or Group III (caspases-6, -8, -9, -10: preference of (L/V)EXD). In the sequence preference, “X” refers to multiple amino acids that are allowed at the position. Finally, caspases are classified based on the length of their pro-domains, either as long or as short pro-domain-containing caspases (Figure 3). In the long pro-domain caspases, the Nterminal regions vary between 119 and 219 amino acids and contain specialized sequences that interact with activation

2.2. Caspase Classifications and Domain Organization

The 11 human caspases are separated into two broad categories, depending on whether they function in inflammation (caspases-1, -4, and -5) or in apoptosis, although one should note that caspase-14 appears to be a specialized enzyme in that it is expressed primarily in cornifying epithelia and is involved in differentiation of corneocytes in the epidermis.57 The apoptotic caspases are further separated into two general subcategories, based on their entry into the apoptotic cascades, either as initiator (caspases-2, -8, -9, and -10) or as effector (caspases-3, -6, and -7) caspases. Initiator caspases are activated after formation of signaling complexes, such as the DISC (death-inducing signaling complex),7,36,58−60 the apopto6668


Chemical Reviews

Review

The caspase protomer is formed by the folding of the large and small subunits into a single domain, referred to as the caspase−hemoglobinase fold (Figure 5),89 consisting of a sixstranded β-sheet core (β1−β6) with five α-helices (H1−H5) on the protein surface. Helices H2 and H3 reside on the same face of the protein, while helices H1, H4, and H5 reside on the opposite face. The protomer also contains three short β-strands on the protein surface near the active site. Following the suggestion of McLuskey and Mottram,90 these three strands are referred to as β1, β2, and β3 because they are not fully formed in all caspases, and as described below, this region of the protein can undergo significant structural rearrangements, which does not occur for the six β-strands in the core. Five loops comprise the active site, and the loops are referred to as L1−L4 and L2′ (Figure 5). L1 connects β1 and helix 1, L2 and L2′ form the intersubunit linker (IL) and connect β4 in the large subunit to β5 in the small subunit, L3 connects β5 and helix 4, and L4 connects helix 5 and β6 (Figure 6). After cleavage of the IL during maturation, L2′ of one protomer interacts with L2 and L4 of the second protomer (Figure 5B) and stabilizes the active conformation,76,91 so a single protomer is not active because it lacks the cross-protomer interactions of the active site loops. A comparison of the amino acid sequences of the 11 human caspases is shown in Figure 6. The units of secondary structure corresponding to the six β-strands, five α-helices, the intersubunit linker (IL), and four active site loops, L1−L4, are indicated above the sequences. The loops are colored the same as those in Figure 5: L1, yellow; L2, red; L3, blue; L4, brown; and L2′, cyan. In addition, the processing sites in the pro-domain linker and in the IL are underlined. The alignment is based on the start of the first β-strand (β1), so gaps that may occur in the DED and CARD regions are not highlighted here. It is somewhat common in the caspase literature to refer to some loops or other regions of secondary structure by the amino acid numbers of the caspase in question.35,92,93 For example, the turn between β1 and β2 is sometimes referred to as the “124-loop” because it contains E124 in caspase-3.92 However, since the numbers differ for each caspase, the nomenclature is not consistent for all caspases, so the 124-loop in caspase-3 would be the “241-loop” in caspase-1, yet they refer to the same turn. It was also suggested that amino acid positions be referred to by the corresponding position in caspase-1;35 however, different loop sizes among the caspases make this problematic and require the use of further identifying terms, such as 381A-H, when the loop in question is longer than the corresponding loop in caspase-1. Because each unit of secondary structure is connected by a turn or loop, for consistency in this review, these units are indicated as T1−T13, recognizing that T1, T10, T11, and T13 correspond to active site loops L1, L2/L2′, L3, and L4, respectively (Figure 6). In addition, because the short surface β-sheet composed of β1−β3 is not fully formed in all caspases, the turns that connect the three short β-strands, T5−T8, could be represented by one loop connecting β3 in the core to helix H3 on the surface (Figure 5). However, turn 6, or T6, connecting β1 and β2 may be important in allosteric regulation and will be described in more detail below, so the three short β-strands are labeled here, as are their connecting loops in Figure 6. 2.3.1. Pro-Domain LinkerEvolutionary Leftovers or Functional Evolution? The alignment in Figure 6 shows five regions with significant differences in length among the caspases: the pro-domain linker, L1, T6, IL (or L2/L2′), and

Figure 4. Substrate binding to the caspase active site. (A) Identification of S5−S1′ binding sites relative to the P5−P1′ positions of the substrate. (B) The binding groove on the protein surface contains a deep pocket (S1) for binding the P1 aspartate. (C) Stick representation of substrate (DEVD) bound to caspase-3 demonstrating positioning of the catalytic histidine (H121) and cysteine (C163) near the peptide bond of the P1 aspartate. Relative positions of the S1−S4 binding sites are indicated in green (caspase-3 PDB ID 2J30).76

scaffolds (see sequence comparisons in Figure 6). Caspases-8 and -10 contain DEDs that interact with similar domains (DDs, death domains) on adaptor proteins, while caspases-1, -2, -4, -5, and -9 contain CARD domains that interact with comparable domains on adaptor proteins. The mechanisms of action for the DED or CARD interactions with adaptor proteins have been the subject of intense study and have been reviewed elsewhere.77−79 Interestingly, CARD-only proteins (COPs), or the similar PYRIN-only proteins (POPs), as well as adaptors in the death domain-containing superfamily are utilized in cells to regulate inflammatory or innate immune responses.80,81 Following the DED or CARD portions of the pro-domain is a region of up to ∼60 amino acids that links the activation domains to the protease domain (Figure 3, pro-domain linker). The short pro-domain caspases are constitutive dimers,66,82,83 so the DED or CARD activation domains are not required for dimerization and have been lost through evolution. The prodomain linker was retained, however, and rather than being evolutionary debris, the linker functions in dimer assembly,84,85 interactions with chaperones,86 or selection of substrates through use of an exosite for ligand binding,87 depending on the caspase, so this region of the pro-domain appears to have evolved to provide individualized functions to the short prodomain caspases. These features are described more fully in subsequent sections. 2.3. Features in the Caspase−Hemoglobinase Fold

An active caspase dimer is derived from the association and processing of two procaspase monomers. As shown in Figure 3, the procaspase monomer consists of an N-terminal pro-domain, described above, and a protease domain that contains large and small subunits connected by a linker, referred to as the intersubunit linker, IL. Mature caspases are homoheterodimers that contain two protomers, where a protomer consists of one large and one small subunit resulting from cleavage of the IL. 6669


Chemical Reviews

Review

Figure 5. Structure of representative caspase zymogen (A) (PDB ID 1GQF)88 and of mature enzyme (B) (PDB ID 2J30).76 In the zymogen, the intersubunit linker (L2/L2′ and IL) is uncleaved and the active site loops L1−L4 are disordered. Positions of strands β1−β6, β1−β3, and helices H1−H5 are indicated. Colors for active site loops L1 (yellow), L2 (red), L3 (blue), L4 (brown), and L2′ (cyan) correspond to regions of the sequences colored in Figure 6, and this coloring system is used throughout the review.

L4. Following cleavage of the pro-domain (underlined in Figure 6) the remaining flexible linker varies in length from ∼16 to ∼64 amino acids. While possible functions of the pro-domain linker have been described for caspases-3 and -7,84,86,87,108 similar functions for the comparable regions in the other caspases have not been determined. Interestingly, the linker forms a short α-helix in caspases-1 and -9, but it is unstructured in caspase-7 and other caspases (Figure 7). Although no role has been assigned for the helix in terms of dimer stability or allosteric regulation, it is interesting to note that this region is phosphorylated at two sites in caspase-9, S144 and Y153,109 and at one site in caspase-7,110 Y58 (Figure 6, red, and Figure 8). The side chain of S144 is exposed to solvent, so it is difficult to speculate on how phosphorylation of this site affects protein stability or function. Tyrosine 153, however, forms hydrogen bonds with the side chain of D340, which resides on L2′, and the backbone amide of R408. The side chain of R408 forms hydrogen bonds across the dimer interface with N389′ of helix

5 of the second protomer (helix 5′ in Figure 8A). Phosphorylation of Y153 may affect the open-to-closed transition of L2′ (described below), thereby inactivating the enzyme, or it may decrease dimer stability by inhibiting the interdimer interactions of R408. As shown in Figure 8B, Y153 is conserved in several caspases while D340 is conserved in all caspases, but N389 and R408 are not conserved. Interestingly, however, helix 5 and the C-terminal regions are modified in other caspases (Figure 8B), suggesting that, although the specific interactions observed in caspase-9 may not be conserved in other caspases, modifications of the pro-domain linker, helix 5, or the C-terminal region of the small subunit may have similar effects in allosteric regulation of caspase activity, through changes in either protein stability or conformation, thus resulting in conformational selection in the native ensemble. A more global view of post-translational modifications is discussed in section 4. 2.3.2. Turn 6 Provides Flexibility near the Active Site. A second region of the protein that shows large differences 6670


Chemical Reviews

Review

Figure 6. Sequence alignment of 11 human caspases. All sequences are from the Universal Protein Resource Knowledgebase (UniProtKB)94 and are available at http://www.uniprot.org. The following UniProt accession codes were used in the alignment: caspase-1, P29466 (ref 95); caspase-2, P42575 (ref 96), caspase-3, P42574 (refs 97,98); caspase-4, P49662 (ref 99); caspase-5, P51878 (ref 100); caspase-6, P55212 (ref 101); caspase-7, P55210 (refs 102,103); caspase-8, Q14790 (ref 69); caspase-9, P55211 (ref 104); caspase-10, Q92851 (refs 105,106); caspase-14, P31944 (ref 107). For each caspase, processing sites are underlined, and secondary structural elements of the protease domain are indicated above the sequence, beginning with β-strand 1, β1. Active site loops L1−L4 and L2′ are also labeled with the corresponding turn identification, and the colors are the same as those in Figure 5 for each loop. The catalytic histidine and cysteine residues are in bold, and sites of phosphorylation or glutathionylation are shown in red.

Figure 7. Structure of caspase-1 (PDB ID 1ICE),33 caspase-9 (PDB ID 1JQX),111 and caspase-7 (PDB ID 1F1J).112 The pro-domain linker (Figure 3) forms a short α-helix in caspases-1 and -9 but is unstructured in caspase-7. Helices 1, 4, and 5 are indicated as H1, H4, and H5.

6671


Chemical Reviews

Review

below in regard to conformational selection in the native ensemble, but the region appears to allow for the evolution of individualized function within a structure that is well conserved in the caspase family. 2.3.4. Variations in Active Site Loops L1 and L4. Two active site loops, L1 and L4, show significant variations in length among the caspases (Figure 6). In the case of L1, caspase-8 contains the longest loop, with an insert of seven amino acids compared to caspase-3, while caspase-14 exhibits the shortest L1, with only five amino acids. Likewise, L4 can be categorized as short (caspases-1, -4, -5, and -9), medium (caspases-8 and -14), or long (caspases-2, -3, -6, -7, and -10) based on its length (Figure 6). The L1 loops in caspases-1 and -9 are shorter by three amino acids compared to L1 of caspase-3 (Figure 11), although the loops are structurally very similar. The additional residues in caspase-3 allow a short helix to form. In caspase-8, the insertion in L1 is accommodated by extending the short helix as well as the apex of the loop such that contacts at the base of the loop are maintained. Currently, there is no structural information on caspase-14, so it is not clear how the caspase maintains critical contacts in this loop that stabilize the active conformation. For comparison, the length of L3 is highly conserved, and in the active conformation the loop forms the substrate-binding groove at the base of the active site (Figure 11). In contrast, L4 is long in caspase-3, medium length in caspase-8, and short in caspases-1 and -9 (Figure 11). Contacts in L1 are critical to the active conformation because amino acids in the loop align the catalytic H121 for catalysis, and the loop contributes to the S1 binding pocket by hydrogen bonding to the P1 aspartate (Figure 12). In caspase-3, T62 in L1 hydrogen bonds to H121, and hydrophobic contacts between amino acids F55 and M61 in L1 and F128 in the surface β-sheet (β1-β3) stabilize the loop.114 Arginine 64, at the base of L1, hydrogen bonds to the P1 aspartate, thus stabilizing substrate binding. Loop 1 is very mobile in the active conformation,92,114 so mutations or ligand binding that affect loop dynamics inactivate the enzyme. In the case of caspase-8, it is not clear that the additional residues affect the activity of the mature enzyme; however, the additional residues may be important for maintaining the zymogen in the inactive state, as described below. Amino acids in L4 form part of the S2 binding pocket (Figure 13). In caspase-3, F256 (L4), F204 (L3), and W206 (L3) form a hydrophobic pocket for binding small hydrophobic residues of the substrate (Figure 13A). In caspase-9, the shorter L4 loop allows for the binding of larger hydrophobic residues that can contribute to the hydrophobic interactions (Figure 13B). The larger L4 in caspase-3 would sterically clash with larger hydrophobic groups. Thus, contacts in L4 are important for determining substrate specificity in the caspases. 2.3.5. Maturation May Result in Active Site Formation. The fifth region of caspases with large variations in length is the intersubunit linker, or IL (Figure 6). Caspases-3, -7, and -14 have short ILs (∼16 amino acids), while the remaining caspases have intermediate to long ILs. In the zymogen, cleavage of the IL separates the loop into two regions, referred to as L2 and L2′ because the loops form part of the active site, although in most cases L2′ is cleaved again, so the length of L2′ will depend on the position of the second cleavage site. The presence of the two cleavage sites imparts great flexibility in regulating activity because the sequences in these sites provide control over zymogen maturation, typically

Figure 8. Phosphorylation sites at the N-terminus of caspase-9 (PDB ID 1JXQ111). (A) The pro-domain linker is phosphorylated at two sites, S144 and Y153, in caspase-9. The cyan-colored region refers to L2′, as in Figures 3 and 6. (B) Sequence comparison for interactions observed in caspase-9. Amino acids that are conserved are shown in bold, and sites of post-translational modifications are shown in red. The numbers above the sequences refer to position in caspase-9 (see Figure 6).

among the caspases is turn 6. As shown in Figure 9A, the sequence includes the three surface β-strands, β1−β3, and turns 6−8 (T6−T8). In most caspases, T6 is a short turn that connects β1 and β2 near the active site (Figure 9B), and the short T6 results in an open central cavity in the dimer interface (Figure 9C). In caspase-9, however, an insertion of several amino acids (Figure 9A) results in a longer loop in turn 6 (Figure 9D) that extends into the dimer interface and occludes the central cavity (Figure 9E). The sequence alignment also suggests that caspases-1, -4, and -5 contain a similar insert to that of caspase-9 (Figure 9A). An alignment of the structures of caspases-1 and -3, however, shows that T6 is actually very similar in the two proteins. Instead, the three surface β-strands, β1−β3, are not well formed in caspase-1, so the insertion is present in T7, which connects β2 and β3 (Figure 10). Thus, for caspase-1, and presumably caspases-4 and -5, the central cavity is open, as observed for caspase-3. The lack of sequence conservation in this region of the structure shows that structural alignments are more reliable in determining the placement of the inserts. The structural fluidity in β1−β3 and the adjoining loops leads to important conformational variations in the native ensemble, as described below. Caspase-9 forms weak dimers in solution (KD > 1 μM)111 compared to caspase-3, which has a dimer dissociation constant in the low nanomolar range.82 Functionally, the longer turn 6 in caspase-9 occludes a well-described allosteric site in the interface cavity.40,41 Thus, the insertion may, on the one hand, result in an increase in stability of the dimer due to new interactions between the longer turn 6 loops in each protomer, but on the other hand, the allosteric site may no longer be accessible for ligand binding. The flexibility in the β1−β3 strands and their adjoining loops is described more fully 6672


Chemical Reviews

Review

Figure 9. Inserts in the short β-sheet and turns on caspase surfaces. (A) Sequences of the turn 6 region, as in Figure 6. (B and C) Cartoon (B) and space-filling model (C) of caspase-3 (PDB ID 2J30).76 (D) The central cavity is shown in red. The longer turn 6 in caspase-9 (D, PDB ID 1JXQ)111 occludes the central cavity (E).

cleavage of the IL and formation of the substrate-binding pocket.117−119 Models of the caspase-3 zymogen suggested that the short IL was sufficiently long to allow active site formation if the IL were released from binding to the dimer interface,120 based on the observation that a mutation in the dimer interface resulted in constitutive activation of procaspase-3,121 most likely due to expulsion of the IL from the interface (Figure 14C).122 Lengthening the IL results in constitutive activation of procaspase-3, but the results are confounded by alternate cleavage of the variants at sites other than the IL processing site. 120 Thus, there remains some discussion on the “zymogenicity” of caspase-3, that is, the ratio of the activity of the mature caspase to that of the zymogen.117 While it is clear that the zymogen of caspase-3 does not possess sufficient enzymatic activity to carry out apoptosis, the issue of zymogen activation and selection of an active conformation in the zymogen is important in the design of allosteric activators for

by combining sequences for self-cleavage with sequences that link maturation to activation of other caspases.115,116 Cleavage of the IL results in displacement of the IL from binding in the dimer interface (Figure 14A). The loop, now called L2′, moves across the body of the protease domain and forms new interactions with L2 and L4 of the second protomer. Regarding active site formation, L3 moves from a solventexposed position toward the dimer interface and forms the substrate-binding groove (Figure 14B). The presence of the IL in the interface prevents the insertion of L3 due to steric clashes, but removal of the IL provides sufficient room for the “elbow loop” of L3 to insert in the dimer interface such that P201 on L3 forms one side of a triplet with R164 on L2 and Y197 on β6 (caspase-3 numbering). The interactions are important for stabilizing L2 in the active conformation, and importantly, the catalytic cysteine is positioned for catalysis. While the three residues in Figure 14 are not conserved in all caspases (Figure 6), similar movements occur following 6673


Chemical Reviews

Review

zymogen, turn 6 moves into the position observed in the mature caspase-3. For the inactive zymogen, the “up” position of turn 6 is stabilized by hydrogen bonds formed between R164 (on L2), Y197 (on β-strand 6), and backbone atoms of turn 6 (Figure 17C). Insertion of the elbow loop from L3 into the dimer interface (see Figure 14B) results in the flipping of Y197 and E124 into the “down” position, where the three residues form hydrogen-bonding interactions that stabilize L2 in the active conformation as well as turn 6 in the down position (Figure 17D and 17E). The importance of the interactions is shown by the nearly 100-fold decrease in activity of mature caspase-3 upon substitution of E124 and Y197.92 In addition, the binding of small molecule allosteric inhibitors to the dimer interface, as described in more detail below, prevents Y197 from flipping to the down position, thus preventing insertion of the elbow loop of L3 and formation of the substrate-binding pocket.40,41 In addition to structural information on the maturation of the short IL caspase zymogens of caspase-3 and caspase-7, there is extensive information on the maturation of caspase-8, with an intermediate-length IL, and of caspase-6, with a long IL. In the case of caspase-8, active site loop 1 (L1) is folded into the active site and interacts with the IL to stabilize an occluded active site conformation (Figure 18).113,124 Thus, the inset in L1, described above, may provide flexibility so that the loop can fold into the active site. The region of the IL that becomes L2 after cleavage (red in Figure 18) occupies the substrate-binding pocket and prevents L3 from inserting into the active site (blue in Figure 18, top). As a result of the position of the IL, L3 rotates toward helix 5, so L3, H5, and L4 are disordered, requiring the ordering of a significant portion of the protomer upon activation. On the other side of the protein, because the IL intercalates between L3 and turn 6, the short surface β-sheet, β1−β3, and helix 3 must rotate away from the dimer interface to avoid steric clashes. Consequently, the catalytic H317 is rotated nearly 180° away from the active site (Figure 18, bottom middle). Thus, the intermediate-length IL of procaspase-8 affects every active site loop, the short surface β-sheet, helices-3 and -5, and the catalytic residues. In contrast to procaspase-8, the zymogen of caspase-6 demonstrates fewer structural rearrangements upon activation (Figure 19).119 The long IL intercalates between L3 and turn 6, as with procaspase-8; however, L3 is able to insert the elbow loop in the dimer interface and form the substrate-binding pocket. The resulting rotation of the short β-sheet, β1−β3, and helix 3 provide sufficient room in the interface for both IL and elbow-loop binding. Although L4 is disordered in procaspase-6, L1, the short surface β-sheet, and L3 are formed, and the catalytic H121 and C163 require minor movements upon activation. Strikingly, the second cleavage site in the IL (see Figure 6), TEVD, is positioned in the active site for selfactivation, so intramolecular cleavage of the site by procaspase6 results in removal of the IL from the active site, structural rearrangements in L2 that stabilize L4, and movement of helix 3 and the short surface β-sheet into the active position. The structural data agree with previous biochemical data that show procaspase-6 self-activates both in vitro and in cellulo.125 The structural variations in the intersubunit linkers reveal three general themes regarding mechanisms to maintain the zymogen in the quiescent state. First, the IL prevents the elbow loop of L3 from inserting into the dimer interface, thereby preventing formation of the substrate-binding pocket. As a

Figure 10. Comparison of the β1−β3 region in caspase-1 (PDB ID 1ICE)33 (colored) and caspase-3 protomer (gray). In both caspases, turn 6 is very similar but the insertion in caspase-1 shown in Figure 9A is accommodated in turn 7. As a result, the central cavity is open, as shown in Figure 9C for caspase-3.

the treatment of cancers since the constitutively active procaspase-3 zymogen is not inhibited efficiency by XIAP, the endogenous inhibitor of caspase-3.122 The binding of a small molecule activator with micromolar affinity123 or synthetic antibodies with nanomolar affinity117 shows that either there is a large activation barrier between the active and the inactive states or there is a large change in conformational free energy such that picomolar affinities may be required for efficient activation. Nevertheless, cleavage of the IL and movements of L2′ result in formation of the “loop bundle”, which is characterized by extensive hydrogen bonding between loops L2′, L2, and L4 of the second protomer (Figure 15). The interactions have been shown to be critical for stabilizing the active conformation.76,91 In the zymogen, because the IL is bound in the interface, the amino acids that comprise the loop bundle are not positioned correctly to stabilize the active configuration. In principle, cleavage of the IL is not required to form most of the interactions in the loop bundle (Figure 14C) should the IL binding to the interface be abolished. While this is observed in the long-IL zymogens during maturation, as described below, no effectors have been identified thus far that facilitate the inactive-to-active conformational transition in the caspase-3 or -7 zymogens, aside from cleavage of the IL. Recent structures of the caspase-3 zymogen117 show that while the majority of the protease domain is similar to that of the mature caspase, the binding of the IL in the dimer interface results in disordered active site loops (Figure 16A). Remarkably, the “active-proenzyme” conformation was observed structurally following the binding of inhibitor, AcDEVD-CMK, and the data show that the active zymogen conformation is similar to that of the mature caspase in that L2′ moves out of the interface, L1 and L4 become ordered, and the elbow loop of L3 moves from a position in the active site to the dimer interface (Figure 16B). The movements result in positioning the catalytic cysteine and histidine in their active conformations (Figure 17A and 17B). Because the IL is not cleaved to generate the L2 and L2′ loops, however, the loop bundle (Figure 15) is not fully formed, so the catalytic activity is limited. In the inactive zymogen, turn 6 is found in a solvent-exposed position such that the short β-sheet, β1−β3, is not well formed (Figure 17A and 17B). Upon maturation, or upon forming the active 6674


Chemical Reviews

Review

Figure 11. Comparison of active site loops L1, L3, and L4. Caspase-8 contains the longest L1, while caspase-3 contains the longest L4. The length of L3 is conserved in the caspases. Caspase-3 (PDB ID 2J30)76 is shown in yellow; caspase-1 (PDB ID 1ICE)33 is shown in green; caspase-8 (PDB ID 1QTN)113 is shown in orange; and caspase-9 (1JXQ)111 is shown in peach.

result, L3 is generally disordered. Second, the IL disrupts turn 6

3. ALLOSTERY

due to steric clashes caused by IL intercalation between L3 and

3.1. Introduction to Ligand Binding Models and Conformational Space

turn 6, resulting in destabilizing the catalytic residues. Third,

The word “allostery” was originally coined in 1961 by Jacques Monod to describe the mechanism of feedback inhibition of metabolic enzymes by regulatory ligands126 and is taken from the Greek “allo”, meaning “other” and “stereo”, to described the stereospecificity of two binding sites for substrate and regulatory ligands.127 The mechanism describes the binding of an inhibitor and substrate in two distinct sites rather than in two overlapping sites. Static structures of proteins, such as those of caspases described in the previous section, provide a wealth of information on various ground states in the conformational space for the protein, but the static structures provide little information on the transitions between states, activation barriers between the ground states, or on high-energy states

the IL stabilizes a conformation with an occluded or distorted active site such that significant rearrangements are required to form the active conformation. The extensive structural database of caspases shows that activation is not a binary function where the protease is either in an inactive conformation or in an active conformation. As described below, these general themes for changing active site features are not unique to the zymogen and will reappear in the conformational landscape of the mature caspase. 6675


Chemical Reviews

Review

Figure 12. Interactions in L1 stabilize the active conformation. F55, M61, and F128 form a hydrophobic cluster that stabilizes L1 and the surface β-sheet (β1−β3) in the active conformation. T62 hydrogen bonds with the catalytic H121, and R64 interacts with the P1 aspartate of the substrate. Caspase-3 (PDB ID 2J30)76 is used in the example here. Figure 14. Movements of the IL (L2′) (A) and L3 (B) after cleavage of the IL to generate L2 and L2′ active site loops. (C) Model for activation of the zymogen in the absence of IL cleavage.

Figure 15. Formation of the loop bundle upon zymogen activation. In caspase-3, cleavage of the IL to form L2 and L2′ results in rearrangement of L2′ from one protomer and L2 of the second protomer to form an extensive hydrogen-bonding network across the dimer interface. The prime (′) indicates amino acids from the second protomer (caspase-3 PDB ID 2J30).76

Figure 13. Interactions in L4 are important for determining substrate specificity. In caspase-3 (A), hydrophobic interactions contributed by residues in L3 and L4 form the S2 binding pocket. In caspase-9 (B) the shorter L4 allows larger hydrophobic residues in the P4 position to interact with the hydrophobic groups. The larger L4 in caspase-3 prevents large hydrophobic residues in P4 due to steric clashes. Caspase-3 (PDB ID 2J30)76 and caspase-9 (PDB ID 1JXQ)111 are used in the examples here.

binding (free energy landscape), and shifts in populations due to ligand binding (population shift).129 Biological macromolecules exist in ensembles of states with fluctuations around an average structure governed by Boltzmann’s principle,130 that is, the ensemble is a collection of closely related conformations. The ability of proteins to form substates due to their dynamic nature is critical to their function, as proteins display a wide range of molecular motions, from side-chain rotamer selection or rotations that facilitate ligand binding, to secondary structural rearrangements, to larger folding or unfolding events.131 Because of the ensemble

that may exist. In addition, the paradigms derived solely from structural data generally fail to quantitatively describe the changes in a system, particularly when functional changes cannot be reconciled with a lack of structural changes, such as may be observed with some site-specific mutants.128 A unifying formalism was described recently as a way to quantify allostery from perspectives of preferred ligand binding states (structural), changes in free energy due to preference in ligand 6676


Chemical Reviews

Review

Figure 17. Conformational changes in turn 6, catalytic C163 and H121, and the dimer interface during maturation of procaspase-3. Amino acid side chains are shown for the inactive zymogen (PDB ID 4JQY, cyan),117 active zymogen bound to Ac-DEVD-CMK inhibitor (PDB ID 4JR0, yellow),117 and mature caspase-3 (PDB ID 2J30).76

Figure 16. Conformational changes in procaspase-3 during maturation. (A) Overlay of procaspase-3 (PDB ID 4JQY, colored)117 and mature caspase-3 (PDB ID 2J30, gray).76 (B) Comparison of active site loops in mature caspase-3 (PDB ID 2J30, gray),76 procaspase-3 (PDB ID 4JQY, cyan),117 and active zymogen conformation of procaspase-3, with Ac-DEVD-CMK bound (PDB ID 4JR0, yellow).117

In this example shown in Figure 20, two of the substates are enzymatically inactive while one substate is enzymatically active. Post-translational modification (PTM) of the enzyme results in selection of one inactive substate (Figure 20, top right). Because the three substates are in equilibrium, selection of one substate by modification of the enzyme leads to redistribution of the population of substates such that the conformation that represents the modified (and inactive) substate is favored. Likewise, an allosteric inhibitor binds to the second inactive substate, which in this example is different from that state that is stabilized by the PTM, and stabilizes the second inactive substate (Figure 20, right lower). As with the state stabilized by the PTM, selection of a substate that is enzymatically inactive by the allosteric ligand results in redistribution of substates such that the ligand-bound state is favored. Finally, one substate in the ensemble is enzymatically active and selected by the binding of substrate (Figure 20, right middle). In this case, the substrate selects the active state, again resulting in redistribution of substates in the ensemble such that the active state is favored. In contrast to the simple example shown here, the cellular milieu is much more complex, and the presence of multiple substrates, post-translational modifications, and allosteric ligands compete for substates within the native ensemble. Thus, the activity of the enzyme in the cell reflects the relative distribution of these states, and the activity will fluctuate based on the availability of substrates, the concentration and affinity of allosteric ligands, and levels of post-translational modifica-

nature of protein conformational states, perturbations in the ensemble result in a redistribution of structures in the ensemble. As such, allostery refers to the stabilization of cooperative units due to ligand binding, resulting in selection of states within the native ensemble.128,132,133 The sites for allosteric ligand binding are spatially distinct from the substrate-binding site, so allostery has been described as “regulation at a distance”.134 The binding of ligand at one site perturbs the dynamic equilibrium of the ensemble and causes a change at another site, usually described by changes in conformation or dynamics, and is correlated with a change in function. An example of the phenomenon is shown in Figure 20. In this case, the average conformation can be described by three closely related substates in the ensemble, only one of which is active. One should note that while this review focuses on enzyme activity, particularly of caspase proteases, allostery has been shown to be critical to the regulation of a variety of cellular signaling reactions, from the Lac operon,130 nuclear receptors,135 the ubiquitin−proteasome system,136 cytochrome P450,137 hemoglobin;132,138−140 fatty acid binding protein 4 (FABP4),141,142 nitrogen regulatory protein c (NtrC),143−145 response regulator RegA,146 lysine-, arginine-, ornithine-binding (LAO) protein,147 and molecular chaperones.148 6677


Chemical Reviews

Review

Figure 18. Procaspase-8 undergoes large conformational rearrangements upon activation. The intermediate-length IL stabilizes L1 in a position in which it is folded into the active site (bottom left panel: yellow, procaspase-8; gray, mature caspase-8). The position of the IL also causes rotations of β1−β3 (Turn 6) and of helix 3 away from the dimer interface (bottom middle panel: colored, procaspase-8; gray, mature caspase-8), resulting in nearly 180° rotation of the catalytic H317. The IL also prevents L3 from forming the substrate-binding pocket (top panel) and must move out of the active site for the active conformation to form (bottom right panel: colored, IL of procaspase-8; gray, L2 of mature caspase-8. The green color indicates region of the IL that is removed upon processing. Procaspase-8 PDB ID 2K7Z124 and caspase-8 PDB ID 1QTN.113

The induced-fit model of Koshland,150 also called the “reaction front” model,142 postulates that the original conformation undergoes a structural rearrangement into an optimal conformation and that the rearrangement is induced by the binding of ligand.151 This model presupposes that a protein has two states, open and closed, and that the conformational change optimizes the bound complex. The conformational selection model152 postulates that there are a large number of pre-existing states of the macromolecule and that the state which is most complementary to the ligand will bind. The binding event is then followed by a shift in the population of states toward the most complementary state. The two models (Figure 21) are considered to be competing and mutually exclusive models, but the conformational selection model is often invoked to describe the relaxation kinetics of ligand binding.151,153 The models are distinguished by the ordering of the binding step and protein conformational change along the reaction pathway. As shown in Figure 21, one assumes that two states exist in the macromolecule, P1 and P2. While P1 binds substrate weakly and is inactive, P2 binds substrate tightly and is active. In the induced-fit model, the protein is observed in the P1 state in the absence of ligand.

tions. The activity will also reflect the ability of the protein to sample all substates in a biologically relevant time scale, as some substates may have high activation barriers and thus may have very slow transitions.142 The selection of states within the ensemble provides the cell with a mechanism to exquisitely fine tune activity and links the enzyme activity to other cellular signaling pathways. In short, conformational selection through allosteric ligand binding provides the cell with a means to sense changes in conditions and to respond rapidly. An underlying premise of allosteric regulation is that of molecular recognition.149 As described in the previous section, for example, caspases recognize their substrates through a specific tetrapeptide sequence. Thus, enzyme active sites must be complementary to their substrates. Likewise, allosteric sites on enzymes must recognize specific ligands complementary to that site, and while the precise mechanisms are not known, protein conformational dynamics contribute to protein−ligand (or substrate) interactions. Two models have been proposed to describe protein−ligand interactions, the induced-fit model, and the conformational selection model. Both models have a common feature in that they both posit the ability of a protein to undergo a conformational change in the presence of ligand. 6678


Chemical Reviews

Review

conformational rearrangement to a more optimal binding surface. In the conformational selection model, P1 and P2 exist in a pre-established equilibrium, and the population of each state is governed by the equilibrium constant, K3, although the relative population of the inactive P1 is higher than that of P2.

Figure 22. Conformational selection and induced-fit models for ligand binding to an allosterically regulated protein. In this model, the protein binds ligand (cyan triangle) in the active site. The protein also contains an allosteric site (brown) that binds allosteric effector (I). Binding of I shifts the population to the inactive P1, while binding ligand shifts the population to the active P2.

Figure 19. Structural rearrangements upon maturation of procaspase6. The IL of procaspase-6 (cyan and green; PDB ID 3NR2)119 intercalates between Turn 6 and L3, positioning the second IL cleavage sequence, TEVD in the active site for intramolecular cleavage. Minor structural rearrangements are necessary following IL cleavage because the active site is very similar to that of the mature caspase-6 (gray, PDB ID 3P4U).

In the presence of ligand, P2 is selected due to its complementary binding surface, even though it is present at a lower relative population than P1, and the formation of P2:L results in a population shift of the remaining P1 and P2 to reestablish the equilibrium governed by K3. With sufficient concentration of L, all of P1 shifts to the active P2 state. The example in Figure 21 focuses on a single ligand and a single binding site. When the binding site is considered the enzyme active site and the ligand is considered the enzyme substrate then a more realistic, albeit more complex, scenario is shown in Figure 22 for an allosteric enzyme. In this model, the protein exists in two states, P1 and P2, as in the simple model of Figure 21, but in addition to binding ligand, L, in the active site, the protein also binds an inhibitor, I, allosterically. As in the previous model, P1 represents an inactive state that binds ligand (substrate) weakly, while P2 represents an active state that binds ligand tightly, and P1 has a higher population due to the relative thermodynamic stabilities of the two states. In the absence of allosteric effector (I), the model reduces to that shown in Figure 21, that is, the front face of the model in Figure 22 shows the conformational selection or induced-fit conformational changes in the presence of substrate (L). While the substrate binds tightly to P2 and weakly to P1, the opposite is true for the allosteric effector (I). The effector binds more strongly to P1 than it does to P2 and results in a population shift to the P1I state. In this model, the relative affinity of I for P1 over P2 will determine the efficiency by which the allosteric effector inhibits the enzyme. Importantly, because the binding of the allosteric effector is reversible, removal of the effector results in re-establishing the intrinsic population distribution of P1 relative to P2 based on their thermodynamic stabilities. The conformational selection model also allows for the presence of higher energy states relative to the more stable ground state (Figure 23). Although the higher energy states are present at relatively low concentrations, the complementary

Figure 20. Example of selection within an ensemble of states of a biological macromolecule resulting in redistribution of states in the ensemble.

Figure 21. Reaction pathways for conformational selection (left) and induced-fit (right) ligand binding models.

When ligand is present, it binds to P1 to form the weak P1:L complex because the binding site is not optimal for the ligand. The binding event induces a conformational change to the tight-binding P2 state (P2:L), so the binding of ligand induces a 6679


Chemical Reviews

Review

Figure 23. Energy landscape of two species, A and B, in which a highenergy conformation, β, allows conversion of A to B through conformational selection.

ligand-binding surfaces allow the selection of the minor states and subsequent shift in population toward that state. When accounting for higher energy states in the conformational landscape, the simple model in Figure 20 becomes more biologically significant, as shown in Figure 23. In this case, two conformations of a protein, A and B, exist in an energy landscape, where A also contains a higher energy substate, β, and a lower energy ground state, A. In this scenario, the relative population of species favors A. If the binding site of β is complementary to the ligand, and thus higher affinity than A, then the binding of ligand shifts β to the more stable ground state, B. Following the binding of ligand to β and the subsequent conformational change to B, species A reequilibrates to A and β based on the relative free energies of the two substates. In keeping with the simple models shown in Figures 21 and 22, one state is enzymatically active, A, for example, and one state is enzymatically inactive, B, for example. The binding of allosteric ligand inactivates the protein by selecting the highenergy substate β from the substates in the ensemble of active protein, A, and shifts the conformation to the inactive state, B. One should note that the opposite scenario is also true. If B represents the enzymatically active state and A represents an inactive state then the allosteric effector selects the higher energy substate, β, from the ensemble of substates in A and shifts the conformation to the active state B. In this scenario, the allosteric effector functions as an activator rather than an inhibitor. There is considerable interest in defining residue-to-residue interactions responsible for allosteric regulation.129,154−156 In these studies, interactions among discrete sets of residues, called interaction networks, propagate the allosteric communication between an allosteric site and an active (or functional) site. Similar to the induced-fit mechanism described above, amino acids in the allosteric site propagate strain energy, or “frustration”, that results from binding allosteric ligand to the active site through the residue-to-residue interaction network (RRIN). Interaction networks have been described for some allosteric proteins, such as CheY154,155,157 and PDZ family proteins.158 Interestingly, a RRIN has been described for caspase-1 between an allosteric site in the dimer interface and the active site.39,156 In this case, four residues were identified as critical to the propagation of the allosteric signal from the allosteric site to the active site in the protomer (Figure 24), and several other residues were identified through computational studies as important for interprotomer communication between the two active sites.156 The dimer interface has been identified as an allosteric site in several caspases,40−42 and the allosteric mechanism facilitates a transition that stabilizes an inactive

Figure 24. Residue-to-residue interaction network (RRIN) in caspase1.39,156 (Top) Numbers represent positions in caspase-1 sequence. Four residues in the RRIN are shown in bold in the sequence (top), and their positions are shown on the structure of the caspase-1 protomer (bottom) (PDB ID 1ICE).33

conformation with disordered active site loops. The transition appears to reverse the activation mechanisms described in the previous section, where the elbow loop of L3 is expelled from the dimer interface, resulting in disruption of the substratebinding groove and disordering of the active site loops. The two states, that is, active, with ordered active site loops, and inactive, with disordered active site loops, are two primary states observed thus far in caspases. In the caspase-1 RRIN, however, E390 is present only in caspases-1, -4, and -5 (Figure 24, top). While at present there are insufficient mutational or computational studies on other caspases to speculate on a common communication pathway in the caspases, it is interesting to note that caspase-7 is phosphorylated at two tyrosine residues near the RRIN of caspase-1, as described in more detail below. 3.2. Conformational Space of Caspases

3.2.1. Global Free Energy Diagram of the Caspase Family. Building on the simple model shown in Figure 23, which posits two states, A and B, that interconvert through a high-energy substate, β, the caspase conformational space can be represented by a complex group of states that represent the protomer, two dimeric states, one of which is enzymatically active, and the heterodimer, as shown by the general model in Figure 25. 6680


Chemical Reviews

Review

substate, α, also is present in the ensemble of C and represents the enzymatically active zymogen dimer. The zymogen dimer is processed to form the mature caspase dimer (A), in an irreversible reaction, or the active zymogen dimer, α, is stabilized to a lower energy state that resembles state A through interactions with effectors. The mature caspase dimer (A) also contains a higher energy substate, β, that is inactive and represents the dimer with disordered active sites, described above, or other inactive substates in the ensemble, which are described in more detail below. Binding of allosteric effectors stabilizes the inactive substate to yield B. Examples of caspases in each of these states are described in section 4. This model represents the global conformational space for the caspase family of proteases, recognizing that differences in the relative free energies will vary for each caspase. For example, in caspases that are constitutive dimers, such as caspases-3, -6, and -7, the activation barrier between C, the homodimer, and δ, the monomer, is very high, so access to the

Figure 25. Free energy landscape describing the conformational space of caspases. In this model, D represents the heterodimer, C represents the procaspase dimer, A represents the active mature dimer, and B represents the inactive mature dimer.

In this model, the caspase zymogen is observed in two states represented as the homodimer (C) and the heterodimer (D). A high-energy substate (δ) is present in the ensemble of C and allows for heterodimer formation (D). A second high-energy

Figure 26. Model for the global conformational space of caspases. Inactive states are represented by yellow interfaces, while active states are represented by red interfaces. The pro-domain is shown by the blue line, and active site loop L2 is shown by the red line. DED refers to death effector domain, and DD refers to death domain. One should note that the DED-DD domain is replaced by the CARD domain in some caspases and activation complexes. 6681


Chemical Reviews

Review

associate with ligand, while many other protein molecules would require a transition to the binding-competent state prior to ligand association. Due to the limitations of the structural database and the lack of kinetic descriptions of ligand binding, the ensuing discussion of caspases is a comparison of the structures of different liganded states rather than a discussion of substates in the conformational ensembles. 3.2.3. Monomers, Dimers, Heterodimers, and the Formation of an Active Enzyme. Putting the complex free energy landscape model shown in Figure 25 into a structural model provides the basis for understanding the global landscape available to the caspase proteases in the cell, resulting in the model shown in Figure 26. Starting with the zymogen monomer (P1), several reactions are available for the monomer to form the zymogen dimer through homodimerization (P2/ P3), homodimerization through interactions with an activation complex (P13/P14), heterodimerization to form an inactive (P12) or active (P15/P16) complex, or aggregation. Each zymogen dimer is assumed to be an ensemble of at least two states, inactive (P2, P13, P15) and active (P3, P14, P16), where the relative population of the two states is determined by their relative free energies, as expressed in the equilibrium constants K2, K10, and K12. In this model, interactions with an activation complex shifts the equilibrium (K10 and K12) toward the active conformation. Following dimerization, maturation results from cleavage of the intersubunit linker (IL), as described above and represented as Cleavage 1 in Figure 26, resulting in a mature caspase dimer that retains the pro-domain but with the IL cleaved to yield the L2 and L2′ active site loops (P4/P5/P6). After Cleavage 1, the mature caspase dimer is represented by an ensemble that includes the active enzyme (P4), a low-energy inactive state (P5), described above as the state with the disordered active site, and a high-energy inactive state (P6). The equilibrium constants, K3 and K4, describe the relative distribution of the active and inactive states, recognizing that P5 (and P8 below) may be the lowest free energy state for some caspases. One recognizes that many caspases have two cleavage sites in the IL (see Figure 6) and that the order of cleavage of the two sites may be important for determining enzyme activity.116 While the two cleavage sites represent an additional mechanism to fine tune caspase activity, possibly by providing access to additional conformational space, for purposes of the model presented here, the IL is considered either intact or cleaved. Cleavage 2 results in removal of the pro-domain and provides an ensemble centered on the fully mature dimer, P7, P8, and P11. In this model, there are two primary differences in the dimer ensembles. First, as described below, the pro-domain may stabilize the dimer so that when it is removed the dimer dissociates and is in equilibrium with the protomer, P9/P10. Since there is no evidence for an active protomer, the equilibrium constant, K6, most likely favors P10. Second, the pro-domain is bound by chaperones, which may sequester the active enzyme in the cell.108,159 Binding of a chaperone to the prodomain is not a priori allosteric in that the binding event may not cause a structural change in the active site, that is, a population shift as described above; however, the interactions provide a reversible mechanism to decrease the population of active P4, as do allosteric effectors that shift the population toward P5 or P6, so it is considered here with other effectors as a way to regulate caspase activity. The interactions may be used by the cell to fine tune activity under nonapoptotic conditions, and the second cleavage removes the regulatory component.

free energy well of the heterodimer, D, is limited and thus irrelevant on a biological time scale. In contrast, by reversing the relative free energies of δ and C, the initiator caspases reside primarily in a monomeric state where either the heterodimer or the homodimer is accessible through the high-energy substates δ and α, respectively. Finally, in some caspases the relative free energies of the active and inactive states of the mature caspase (A and β) may be close, so that β represents a significant, or even major, state of the mature caspase ensemble. In those cases, allosteric effectors may stabilize β to a lower free energy ground state, B. While evidence for the various conformations is discussed in the section 4, the model shows a complex interplay between states that provides a means to fine tune caspase activity through a combination of evolutionary changes in the free energy landscape and the development of allosteric effectors. Limiting access to various regions of the free energy landscape results in specific mechanisms for regulating caspase activity, as described below. 3.2.2. Caveats Concerning the Description of Conformational Selection in Caspase Ensembles Using the Caspase Structural Database. Although the structural database for caspases is quite extensive, there are a number of intrinsic deficiencies in the database that are problematic for describing conformational selection in caspase allostery. As described above (Figure 21), the conformational selection model refers to a mechanism by which a protein interconverts between a ligand-free state and a ligand-bound state, so the model describes a binding event between the protein and the ligand. In this case, “binding” includes both association and dissociation reactions, but pre-steady-state kinetic measurements are not performed routinely for caspases, so there are no data describing the kinetics of ligand association and dissociation. In addition, the caspase structural database does not adequately address substates in the ensemble. As shown in Figures 21 and 22, conformational selection is an alternative model to induced fit, and ligand binding reactions can incorporate conformational selection or induced-fit mechanisms. The description of allostery in this review is consistent with considerations of ensemble shifts, but at present there is insufficient kinetic data on ligand association and dissociation to make a strong case for conformational selection or to exclude induced fit. For example, to make a case for conformational selection, one assumes that a fraction of the total protein molecules that are present in the unliganded ensemble has a structure that is competent to bind ligand (called the binding-competent state) and that the ensemble also contains unliganded protein molecules that are not in the correct binding-competent state. The small fraction of the total unliganded molecules in the binding-competent state has an intrinsic rate for ligand association, while unliganded protein molecules that are not in the correct binding-competent state transition to that state. In order to explain the ensemble shift, studies on ligand association would also need to describe the rate of interconversion from the state that is not competent to bind ligand to the binding-competent state, such as the rate of the transition from P1 to P2 in Figure 21. Finally, a complete description of the ensemble assumes the presence of many substates in the ensemble and that each of those substates may have a different rate of interconversion to the bindingcompetent state. Overall, a subpopulation of the protein molecules in the binding-competent state would rapidly 6682


Chemical Reviews

Review

Since there is no evidence that the pro-domain affects the intrinsic free energies of P4, P5, or P6, at present these states are considered equivalent to P7, P8, and P11, respectively, except as noted below. Likewise, the equilibrium constants K3 and K4 are considered equivalent to K7 and K5, respectively. Overall, the conformational landscape of caspases shows a complex milieu that can be described as a rough energy funnel with multiple interconverting states,142,152,160,161 where caspase activity can be modulated by interactions with itself or with other proteins and the native ensembles equilibrate with higher energy states that can be selected by allosteric regulators. Evidence for these states and their potential regulation in signaling is discussed in the next section.

4. COMPLEX CONFORMATIONAL SPACE AND ALLOSTERIC REGULATION 4.1. Introduction

As shown in Figure 26, the monomer of the caspase zymogen (P1) can form homodimers, with or without the assistance of an activation complex, or two different heterodimers, depending on the zymogen. The conformational landscape available to the monomer is described first (section 4.2), focusing on the ability of caspases to form multiple complexes. The conformational landscape available to the caspase dimer is then described (section 4.3). Potential mechanisms of conformational selection used by the cell to fine tune caspase activity include shifting monomer assembly to select specific dimeric complexes and post-translational modifications to select enzymatically inactive conformations. Allosteric modulators are described, as are the conformational transitions that result in enzymatic inactivation.

Figure 27. Assembly of the caspase-3 zymogen to the homodimer. (A) Assembly model as shown in Figure 26. (B) Fraction of four species observed during assembly at several protein conformations (dashed lines) and as a function of urea concentration. The four species are placed at their approximate peak in the fraction of total species as the homodimer unfolds in urea, and the free energy for each transition (at 25 °C) is shown above the assembly model. Adapted with permission from ref 162. Copyright American Chemical Society 2001.

4.2. Assembly Processes of the Zymogen Monomer

4.2.1. Short Pro-Domain Caspases Are Constitutive Dimers. The caspase-3 zymogen is the prototypical shortdomain caspase, and the general conclusions are assumed to hold for other short pro-domain zymogens that also are constitutive dimers. The assembly of the homodimer has been studied in detail,84,85,162−165 and the equilibrium constant for dimer assembly (Figure 26, K1) strongly favors the homodimer, with an upper limit for dissociation estimated at ∼50 nM.82 Equilibrium folding studies of the procaspase-3 dimer show that the protein folds through an unusual four-state mechanism (Figure 27). The procaspase-3 monomer (U) folds into a monomeric structure (I) that appears to be partially folded. The monomeric intermediate assembles into a partially folded dimer (I2) that undergoes a transition to the native dimer (N2). On the basis of the level of hydrophobic surface area buried in dimerization (I ⇆ I2) compared to the surface area of the dimer interface in the mature caspase-3,166 the amino acid residues that form the dimer interface appear to be largely intact in the monomer, I. Following dimerization, I2 undergoes a transition that, based on changes in tryptophan fluorescence emission, appears to involve active site rearrangements that results in the native dimer.82,162 One should note that although there is evidence from folding experiments to show two dimeric structures in equilibrium (N2 and I2) and there is evidence from synthetic antibody selection that establishes the presence of two (at least) dimeric structures in the procaspase-3 dimer,117 there is no evidence to show that the structures are equivalent, that is, I2 (Figure 27B) is not P3 (Figures 26 and 27A). Indeed, while the current data support the model of multiple states in the native ensemble for procaspase-3, it is likely that the data report on

different conformations. For example, in the protein folding and assembly experiments, the lowest energy state, N2, also demonstrates enzymatic activity while the dimeric intermediate, I2, a higher energy state, is inactive. In contrast, synthetic antibodies select a high-energy and active state within the native ensemble,117 where the lowest energy state is inactive. The difference most likely is due to the ensemble average approach to the protein folding studies, which examines global spectroscopic properties of the protein under a variety of conditions that perturb the equilibrium,167 such that species populated at relatively low concentrations may not be observed. In contrast, synthetic antibodies bind very tightly to specific epitopes on the proteins and thus can select poorly populated states within the ensemble.168 Although equilibrium studies showed that a large portion of the conformational free energy occurs upon dimerization (Figure 27B, ∼19 kcal/mol of ∼26 kcal/mol), kinetic studies showed that dimerization occurs very slowly, on the order of ∼70 M−1 s−1.85 As shown in Figure 28, following the formation of several intermediates during refolding, the monomer forms a dimerization-competent species (Figure 28A, I8) that assembles to the native dimer, N2. The slow second-order rate constant for association shows that the initial encounter complex for dimerization is very inefficient, that is, the conformation that is most compatible with forming the N2 dimer is scarcely populated in the I8 ensemble. The data suggest that the very low equilibrium dissociation constant for the dimer, ∼50 nM, is due to a very slow rate constant for dissociation of the dimer or the dimer undergoes a transition to a high-energy state from which it dissociates. 6683


Chemical Reviews

Review

Figure 28. Two systems used to examine slow assembly of the caspase monomer. (A) Procaspase-3 dimer interface was mutated to decrease dimerization. Adapted with permission from ref 165. Copyright American Chemical Society 2013. (B) Caspase-8 dimer interface was mutated to improve dimerization. Adapted with permission from ref 85. Copyright Wiley & Sons 2009.

then equilibrated with the native dimer such that the highenergy state was significantly populated relative to the native dimer, N2. Overall, the results suggest that the ensemble of states of the wild-type procaspase-3 contains high-energy states that have are not well populated without perturbations, such as mutations, to stabilize the higher energy states. The second system utilized to examine caspase dimerization is that of caspase-8. The caspase-8 dimer interface contains negative-design elements171 that require more precise surface complementarity for dimerization (Figure 28B). In the case of caspase-8, for example, the aromatic ring of F468 from one protomer interacts across the dimer interface with the flat surface afforded by P468 in the second protomer. These features are in contrast to the constitutive dimer of caspase-3, which presents a relatively flat surface in the protomer (Figure 28B). Although the negative design elements may decrease dimerization efficiency, they also may be important to lower aggregation of proteins with an exposed β-sheet, as might occur in the caspase protomer, due to the requirement for more precise complementarity. Mutations of several negative design elements in caspase-8 resulted in improved dimerization, although the variants did not form constitutive dimers.170 Together with the experiments of procaspase-3, however, the data show that caspases have inefficient dimerization encounter complexes, and mutations that further decrease the efficiency of dimerization result in effectively trapping the protein in the

Practically, the very slow association rate suggests that dimerization can be inhibited by a relatively minor decrease in the association rate constant, possibly resulting in a stable monomer, as one observes in the long pro-domain caspase zymogens (described below). This hypothesis was tested in two ways. First, a procaspase-3 variant was examined for changes in dimerization, where a mutation was introduced into the dimer interface.121,165 Second, the dimer interface of caspase-8, which forms weak dimers in solution,7,15,72,169 was mutated to improve dimer formation.170 The caspase-3 protomer contains a valine residue as the center of β-strand 6 that forms hydrophobic contacts in the dimer interface with the comparable valine of the second protomer (Figure 28). A mutation of V266 to histidine abolished activity in the mature caspase-3121 and dramatically affected association of the dimer.165 In refolding kinetic studies it was observed that the monomer of the V266H variant formed kinetic traps late in the folding reaction due to an effect on the second-order rate of dimerization, that is, the presence of H266 had little effect on folding of the monomer, but H266 made an already inefficient dimerization reaction even more inefficient. With a lowered ability to form dimers, the V266H monomers formed alternative species (Figure 28A, I7*) that established a kinetic competition between aggregation of the monomer or the eventual assembly to a high-energy dimer (N2*). The N2* state 6684


Chemical Reviews

Review

Figure 29. Examples of regulation of the extrinsic (caspase-8) and intrinsic (caspase-9) pathways leading to caspase-3 activation.

monomeric state. As noted below (section 4.2.2), this feature of the caspase zymogen is used by the cell to form a variety of functional complexes. The short pro-domain caspases-3, -6, and -7 evolved as a cluster, called cluster II,172 separate from initiator caspases-8, -9, and -10 (cluster III) and the inflammatory caspases-1, -2, -4, -5, and -14 (cluster I).172−174 Although comparable equilibrium and kinetic data on the folding of initiator caspases are not available, the evolutionary branch point of short pro-domain caspases presumably included species with improved dimerization interaction complexes, subsequent loss of the CARD or DED dimerization domains, and selection of stable, enzymatically inactive ground states in the dimer ensemble. The formation of constitutive dimers not only expanded the conformational landscape for caspases but also provided the need to expand regulatory mechanisms to activate and control activity of the dimers. A few of these features are shown in Figure 29 for the activation of caspase-3 by the extrinsic (caspase-8) or intrinsic (caspase-9) pathway. The procaspase-8 homodimer is activated on the DISC and can be negatively regulated by several reactions that affect activation, such as high levels of c-FLIP,175 phosphorylation,176,177 epigenetic silencing,178−181 or mutations.182,183 Likewise, procaspase-9 is activated on the apoptosome, which is regulated by a variety of processes, including phosphorylation, inhibition of caspase-9, or prevention of Apaf-1 oligomerization.184 Because caspase activation mechanisms are intimately connected to cell signaling pathways, the downregulation of caspases is frequently observed as a way to escape apoptosis in cancers.185,186 Thus, it is of great interest to reactivate the apoptosis cascades in order to induce apoptosis in cancer cells,12 and efforts generally focus on inhibiting proteins that downregulate caspase-3 activation, increasing DISC

Figure 30. Strategy for activating procaspase-3 for inducing apoptosis in cancer cells by selecting for the high-energy, enzymatically active, state in the native ensemble.

formation by the agonist TRAIL (TNF-related apoptosisinducing ligand), or increasing flux through the intrinsic pathway (Figure 29, boxes).187,188 Studies of the constitutive dimers of the short pro-domain caspases may provide new avenues for reactivating apoptosis in cancer cells through the development of allosteric activators (Figure 30). On the basis of the presence of high-energy, 6685


Chemical Reviews

Review

Figure 31. Oligomerization reactions for long pro-domain caspases described in this section. (A) Interaction of the procaspase-9 monomer with XIAP results in formation of an inactive heterodimer. Activation complexes facilitate heterodimer formation between procaspase-8 and c-FLIPL (B) or homodimer formation (C).

catalytically active states in the procaspase-3 native ensemble, allosteric activators that select the active states should prove effective in inducing apoptosis by increasing the relative population of the active state. This strategy was shown in principle through the development of a synthetic antibody that activates procaspase-3117 and through mutation of the zymogen.121,122 In the former studies, the synthetic antibody bound with nanomolar affinity, which is insufficient to adequately populate the active state (Figure 30, P3), and the authors suggest that picomolar affinities may be required to induce significant levels of activity in the zymogen due to the very high energy of the active state compared to the enzymatically inactive ground state. In the latter studies, a mutation in the dimer interface constitutively activated procaspase-3, most likely due to expulsion of the intersubunit linker from its binding site in the interface, allowing the loop bundle to form.121 Importantly, the results showed that the active procaspase-3 was not inhibited efficiently by XIAP, and thus, the constitutively active procaspase-3 effectively kills cells.122

It is worth noting that small molecule activators of procaspase-3 have been described.123,189−192 Small molecule in silico screens identified several potential allosteric activators of procaspase-3, but low affinities resulted in a modest increase in enzyme activity,123 and in agreement with the later studies of Wells and co-workers on synthetic antibodies,117 the data suggest that the binding free energy of the small molecule is several orders of magnitude too low for effectively stabilizing the high energy active state. Finally, small molecules that form fibrils serve as activating platforms for procaspase-3 through induced proximity, but rather than inducing an allosteric transition, the fibrils appear to facilitate locally high concentrations of the active state, resulting in zymogen activation through self-processing.190−192 Interestingly, the zymogen is also activated by Aβ fibrils,192 which may have important clinical implications in understanding caspasedependent neurotoxicity in Alzheimer’s disease. Together, the current data on caspase-3 zymogen activators, which act either through allosteric or direct cleavage mechanisms, show that directly targeting procaspase-3 is an effective strategy for 6686


Chemical Reviews

Review

Figure 32. Caspase-9 is inhibited by XIAP through formation of a heterodimer. (A and C) Caspase-9 dimer (PDB ID 1JXQ).111 (B and D) Caspase9 heterodimer with BIR3 domain of XIAP (PDB ID 1NW9).196

site 2, and highlighted in Figure 32C). The endogenous caspase inhibitor, XIAP (X-linked inhibitor of apoptosis protein), inhibits both the initiation phase and the effector phase of apoptosis by inhibiting caspases-9, -3, and -7.195 While XIAP inhibits caspase-3 through direct interaction of the BIR2 (baculovirus IAP repeat 2) domain with the caspase-3 active site,27 XIAP inhibits caspase-9 by forming a heterodimer.195,196 The BIR3 domain of XIAP binds to the caspase-9 protomer through interactions between β-strand 6, helix 5, and the long turn 6 of caspase-9 (see Figure 9D) and three helices of XIAP-BIR3 (Figure 32B and 32D). In addition, L2′ is maintained in the open conformation due to interactions between the BIR3 domain and the L2′ loop (Figure 32B). While the open and “closed” conformations of L2′ are described in more detail below, the presence of the long turn 6 and the binding of BIR3 to β-strand 6 prevents rotation of L2′ to the closed conformation. The heterodimer with XIAP inhibits caspase-9 by preventing the insertion of active site loop L3 toward β-strand 6 and formation of the substrate-binding groove. Similar to the caspase-9 homodimer, several residues in XIAP-BIR3 clash with the elbow loop of L3 when the substrate-binding groove forms, so L3 remains disordered and caspase-9 is unable to bind substrate. Thus, XIAP selects a low-energy, inactive, ground state in the energy landscape and stabilizes the inactive state through heterodimerization. In contrast to caspase-9, the zymogen of caspase-8 forms a heterodimer that is enzymatically active (Figure 31B). Caspase8 interacts with the cellular FLICE-like inhibitory protein long (c-FLIPL), also through interactions with β-strand 6. In contrast to the BIR3 domain of XIAP, however, c-FLIPL contains a

inducing apoptosis. The direct activators may eventually prove useful as probes for understanding changes in caspase activation mechanisms in various disease states. 4.2.2. Long Pro-Domain Caspases Form Homo- or Hetero-Oligomers. The long pro-domain caspase zymogens are monomers in solution until interactions with activation complexes facilitate dimerization (Figure 26).111,193 Although the precise conformational rearrangements that result in activation of the zymogens on the scaffolds are not known, the presence of stable zymogen monomers provides an opportunity to form multiple oligomeric species. Two of the three oligomeric states shown in the conformational landscape model of Figure 26 are described in this section, as shown in Figure 31. Interactions in the so-called death-fold superfamily, including CARD, DED, and DD domains, have been described in detail,194 and while the “activation complex” in Figures 26 and 31 show DED-DD adaptors, similar interactions occur between CARD domains. Thus, while recognizing that the activation complexes are substantially different and recognizing that assembly and regulation is both highly complex and integral to signaling,36,58,64,67,79 the activation complexes are considered here as general scaffolds for facilitated oligomerization, while specific oligomeric states of the caspases are described below. It should be noted again that this discussion focuses primarily on the apoptotic rather than the inflammatory caspases. Caspase-9 forms a homodimer on the apoptosome, and only one of the two active sites is functional (Figure 32A). Steric clashes in the dimer interface, due to the presence of bulky aromatic residues on β-strand 6 (see Figure 6), propagate to one active site and prevent insertion of active site loop L3 and formation of the substrate-binding groove (Figure 32A, active 6687


Chemical Reviews

Review

The caspase-8 protomer in the heterodimer with c-FLIPL is catalytically active, with a fully formed substrate-binding groove (Figure 33A). Interestingly, the L2′ loop is maintained in the closed conformation, that is, rotated into the dimer interface, even though the caspase-8 protomer is active. Positioning of L2′ is important for forming the “loop bundle” stabilizing interactions between L2′, L2, and L4 (see Figure 15), and as described in more detail below, maintaining the L2′ loop in the closed conformation is a common allosteric regulatory mechanism of caspase activity. In the case of the caspase-8:cFLIPL heterodimer, c-FLIPL provides several amino acids that form a partial loop bundle network with the caspase-8 protomer, and a network of amino acids span the dimer interface and form a hydrogen-bonding network that stabilizes the catalytic cysteine (Figure 33B).197 Two of the amino acids in the network, R391 and Y392, are located at the N-terminal portion of L2′, so the open conformation of L2′ would not maintain the network of interprotomer hydrogen bonds. As shown below, the homodimer of caspase-8 forms the loop bundle by rotation of L2′ into the open conformation. Interestingly, the hydrogen-bonding network in the caspase8:c-FLIPL heterodimer is in a similar position as the allosteric RRIN described previously for caspase-1 (Figure 24).39,156 Functionally, the formation of the caspase-8 homodimer versus the caspase-8:c-FLIPL heterodimer on the DISC is important to the survival or death signaling reactions in the cell (Figure 34). The caspase-8:c-FLIPL heterodimer inhibits necroptosis by inhibiting the formation of the RIPK1:RIPK3 (receptor interacting protein kinase 1 and 3) complex.198 The primary function of caspase-8:c-FLIPL appears to be inhibiting necroptosis by cleavage of RIPK3.199 Adding further complexity the decision of life versus death in caspase-8 signaling, the level of activity of caspase-8 in the heterodimer depends on the processing that occurs in caspase-8

Figure 33. Heterodimer of caspase-8 and c-FLIPL. (A) Formation of the heterodimer with c-FLIPL results in a catalytically active caspase-8 protomer with L2′ in the closed conformation (PDB ID 3H11).197 (B) Amino acids in c-FLIPL and caspase-8 form a hydrogen-bonding network that stabilizes the catalytic cysteine, C360.

caspase-like fold, with a six-stranded β-sheet core and five external α-helices but without an active site (Figure 33A).197

Figure 34. Dimerization pathways of the caspase-8 zymogen result in formation of the caspase-8 homodimer or the caspase-8:c-FLIPL heterodimer with different functional consequences. 6688


Chemical Reviews

Review

as well as in c-FLIPL, such that the lower levels of activity affect substrate selection.169,200 The complex assembly and processing events of the caspase-8 zymogen result in a fine tuning of caspase activity that is used in nonapoptotic events such as cell development and immune cell proliferation.72,199,201,202

Figure 36. Binding of DARPin inhibitors to caspase-2 (top, PDB ID 2P2C),205 caspase-3 (bottom left, PDB ID 2XZD),204 and caspase-7 (PDB ID 4LSZ).203

Figure 35. Conformational landscape of the caspase dimer discussed in this section.

At present, it is not clear how the states in the conformational ensemble of caspase-8 are selected on the activating complex to fine tune enzyme activity, but the varying levels of activity that depend on the processing of caspase-8 or c-FLIPL suggest an allosteric mechanism that may include the selection and stabilization of active states within the ensemble. 4.3. Conformational Selection in the Caspase Dimer

The focus of this section is the dimeric caspase following Cleavage 1, that is, processing of the intersubunit linker (Figure 35). Conformational equilibria and selection are discussed in terms of protein−protein interactions, protein−ligand interactions, and post-translational modifications that affect partitioning of the conformational landscape. 4.3.1. DARPins Inhibit Caspases but Not Always Allosterically. Grutter and co-workers used ribosome display techniques to develop designed ankyrin repeat proteins (DARPin) that bind to caspases-2, -3, and -7 (Figure 36).203−205 Each of the three DARPins binds to its respective caspase utilizing a different binding mode, and only in the case of caspase-2 does the DARPin inhibit the caspase allosterically. DARPin 7.18 binds to caspase-7 with an equilibrium dissociation constant of 144 nM, but there was no detectable change in activity upon complex formation.203 Structural studies (Figure 36, bottom right) showed that the binding epitope resides laterally along β-strand 2 and does not affect the active site. Rather, the DARPin also binds to the caspase-7 zymogen and prevents processing of the intersubunit linker.

Figure 37. Binding of DARPin AR_F8 to caspase 2. The DARPin (green and peach) interacts with L3 and L4 of the caspase, resulting in movement of the catalytic cysteine, C303 (PDB ID 2P2C).205 Caspase-2 inhibited with Ac-LDESD-cho peptide is shown in gray (PDB ID 1PYO).206

DARPin AR_F8 binds to caspase-2 with an equilibrium dissociation constant of 4.1 nM and exhibits mixed-type inhibition with an uncompetitive component, suggesting an allosteric inhibition mechanism.205 Structural studies show that the DARPin binds on the H1, H4, and H5 face of the caspase and interacts with active site loops L3 and L4 (Figure 37). Binding of DARPin AR_F8 to caspase-2 opens the active site cleft and displaces the active cysteine, C303, by ∼1 Å (Figure 37). The data show a new mode of inhibition that does not utilize the closed complex of L2′ (described below) and demonstrates the presence of high-energy states in the caspase 6689


Chemical Reviews

Review

Figure 38. Comparison of wild-type caspase-3 (gray, PDB ID 2J30)76 and V266H variant (color, PDB ID 4EHA).92 The X-ray structure of V266H was used in molecular dynamics simulations to observe rotations in helix 3, the short surface β-sheet, β1−β3, and the catalytic residues, C163 and H121.

ensemble (P6/P11 in Figures 26 and 35). In the case of caspase2, the binding affinity of 4.1 nM is sufficient to select the high energy, and enzymatically inactive, state in the ensemble, thus shifting the population to the inactive state. A similar high-energy, catalytically inactive, state was also observed in a variant of caspase-3. A mutation of V266 to histidine in the dimer interface abolished activity,121 and subsequent structural and molecular dynamics studies showed that the mutation resulted in a transient rotation of helix 3 toward the dimer interface (Figure 38).92 The rotation of helix 3 propagated to turn 6 and caused a lateral shift in the short surface β-sheet, β1−β3, resulting in displacement of the catalytic H121 and C163 by ∼2 Å. The displacement of β1 and H121 causes steric clashes with substrate, so the conformation is not compatible with catalysis. Further mutational studies showed that activity could be restored in the V266H variant by disrupting communications between H266 in the dimer interface and residues on helix 3,114 supporting the conclusion that a high-energy, catalytically inactive, state is present in the ensemble of mature caspase. Selecting the inactive state, as observed with the binding of DARPin AR_F8 to caspase-2, may provide an effective strategy for designing allosteric inhibitors with high specificity, as the epitope recognized by DARPin AR_F8 on caspase-2 is not present on caspase-3. Likewise, there is no evidence, at present, that helix 3 in caspase-2 undergoes similar transitions as observed in caspase-3. A DARPin that binds to caspase-3 has also been described, but the inhibitor binds to the active site and acts as a competitive inhibitor, with an affinity of 9.6 nM.204 Structural studies show that DARPin 3.4 interacts with L4 of caspase-3, and the variable loops of the DARPin occupy the substratebinding groove (Figure 39). Interestingly, the interaction traps Y204 in the unliganded conformation (Figure 39A). In the unliganded caspase, Y204, on L3, occupies the S2 binding site and must rotate toward L2 for substrate to bind. Molecular dynamics simulations show that Y204 is very mobile and rotates between the unliganded and the active positions (Figure

Figure 39. DARPin 3.4 binds to the active site of caspase-3 (A) and traps Y204 in the unliganded conformation (PDB ID 2XZD).204 (B) Molecular dynamics simulations of wild-type caspase-3 show rotations in Y204 from the unliganded conformation, which clashes with substrate binding, to the active conformation. (B) Two hundred frames (at 250 ps intervals) of a 50 ns simulation.

39B).92 Thus, while inhibition by DARPin 3.4 is not allosteric, the structures demonstrate an interaction that could be manipulated by allosteric effectors, that is, selecting inactive states by trapping Y204 in the unliganded state. Overall, the three DARPins show inhibition by three binding modes: preventing maturation of the zymogen, presumably by limiting access to the intersubunit linker cleavage site (caspase7); selecting a high-energy inactive state from the native ensemble, where the state demonstrates displaced catalytic residues (caspase-2); and binding in the substrate-binding groove as a competitive inhibitor (caspase-3). The data further establish the presence of high-energy states in the ensemble and provide strategies for selecting the inactive states. It is interesting to note that DARPin 3.4 selected the low-energy ground state rather than the higher energy inactive state of caspase-3, as for DARPin AR_F8 and caspase-2, suggesting that the high-energy state of caspase-3 may have a higher energetic barrier than that of caspase-2. Using the more recently developed allosteric mutants of caspase-3 may provide access to DARPin selection by increasing the population of the inactive species in the ensemble.92,114 4.3.2. Pro-Domain Linker Affects Caspase Function. It is worth noting several functions of the pro-domain linker (see Figure 3) in a review of caspase allostery and conformational selection. While interactions with the pro-domain linker may not be allosteric, per se, the linker does affect caspase activity in the cell. As shown in Figure 6, the pro-domain linker is one of 6690


Chemical Reviews

Review

Figure 40. Pro-domain linker regions of human caspases. Underlined amino acids represent cleavage sites. Amino acids in red show sites of phosphorylation.

the least conserved regions of the caspases. When considered to include amino acids C-terminal to the Cleavage 2 site (Figure 26) to the beginning of β-strand 1, the linker varies in size from 17 to 64 amino acids (Figure 40). In addition to the extra N-terminal helix described above for some caspases (Figure 7), the pro-domain linker is phosphorylated in several caspases (Figure 40, amino acids in red), although the functions of the phosphorylation events are not known at present. In addition, the pro-domain of caspase-9 is important for docking of ERK to the CARD domain and subsequent phosphorylation of T125.207 In caspase-7, the linker functions as an exosite for substrate selection87 and protects a nuclear localization signal.208 In caspase-3, the region is ubiquitinated by cIAP1,209 leading to degradation by the proteasome, it functions as a molecular chaperone to facilitate assembly of the dimer,84,85 it is bound by the chaperone Hsp27,86 which sequesters the protease from its substrates, and it silences the activity of caspase-6.210 The mechanisms for the pro-domain interactions and how they affect caspase activity are not known, but there are three general themes for regulating caspase activity: first, the interaction can inhibit (or enhance) processing of the intersubunit linker; second, the interaction could occur in the caspase active site, as observed for competitive inhibitors; third, the interaction could act allosterically by disrupting the positions of the catalytic groups. The pro-domain of caspase-

3, for example, is known to bind weakly to the protease domain,82 but thus far the mechanisms appear not to be allosteric. Here, the conformational landscape shows separate states before (P4/P5/P6) and after (P7/P8/P11) Cleavage 2 (Figure 26). In terms of allosteric regulation, it is not yet clear whether the presence of the pro-domain linker affects allosteric interactions, so it is not clear whether P4 differs from P7, for example. In terms of cellular function, however, the presence of the pro-domain linker does affect caspase activity in the cell. An example of the importance of the interaction between Hsp27 and caspase-3 is shown in Figure 41. Prior to Cleavage 2, which removes the pro-domain, the small heat shock protein, Hsp27, binds to the pro-domain of caspase-3 and sequesters the caspase from its substrates.86 Removal of the pro-domain in caspase-7 was shown to dramatically increase apoptosis, likely due to the removal of interactions that sequester the protease in the cytosol.108 Following Cleavage 1, in the intersubunit linker, both the caspase-3 zymogen and the mature caspase-3 can interact with Hsp27. For the zymogen, the interaction inhibits its activation by the initiator caspases-8 and -9 (Figure 41), while the enzymatically active mature caspase is sequestered from its substrates. The interactions are intimately linked to kinase signaling, particularly in regard to phosphorylation of caspase-3 (described below) and of Hsp27.211 Under normal cellular conditions, the interactions with Hsp27 may be used to fine tune caspase activity in the cell. Under apoptotic conditions, the pro-domain of caspase-3 is removed, resulting in removal of the Hsp27 interaction site and release of the active protease in the cytosol. Likewise, in some cancers, the increased expression of Hsp27 effectively prevents apoptosis, most likely due to sequestering caspase-3.212,213 For these reasons, the mature enzyme is separated on the conformational landscape in regard to the presence or absence of the pro-domain. Although the broad outlines are known regarding the importance of the interactions in the cell for regulating caspase activity and the low conservation of this region among caspases, further experimentation is required to understand how the linker is individualized for each caspase and how the cellular interactions affect partitioning of the ensemble. 4.3.3. Allosterically Inhibiting Caspases through the Closed Conformation. As described above, allosteric inhibition of caspases ultimately works, in principle, by disrupting the positions of the catalytic histidine−cysteine dyad. The specific details of how the disruption is achieved in each caspase leads to the possibility of designing allosteric

Figure 41. Model for the interaction of Hsp27 with the pro-domain of caspase-3. 6691


Chemical Reviews

Review

Figure 42. Comparison of caspase-7 closed structures in the zymogen (PDB ID 1GQF)88 and unliganded,118 mature (PDB ID 1K86), conformations with the active conformation (PDB ID 1F1J).112

In contrast to caspase-7, the active site loops of caspase-1 lie closer to those of the zymogen than to the active conformation, particularly for the substrate-binding groove, L3 (Figure 43).214,215 The closed conformation of procaspase-1 requires an expansion of the definition of closed because the region of L2′ (in the intersubunit linker) does not occupy the central cavity of the dimer interface, as observed in caspase-7 and the heterodimer of caspase-8:c-FLIPL. In procaspase-1, the Nterminal region of the intersubunit linker, which becomes L2 after cleavage of the IL, is folded into the interface and occupies the central cavity. Upon cleavage of the IL, L2 rotates out of the interface and toward L4 but L3 remains solvent exposed, as in the zymogen, and the loop bundle between L2, L2′, and L4 is not fully engaged. As a result, both catalytic residues are not positioned for catalysis, and the catalytic cysteine must move ∼3 Å from its position in the zymogen to its active position. 4.3.4. Caspase-6 Expands Conformational Space and Provides More Ways To Allosterically Inhibit the Enzyme. Caspase-6 is unique among the caspases in that three distinct allosterically inhibited conformations have been described. In the first conformation, L2′ forms the closed conformation by rotating into the central cavity of the dimer interface (Figure 44). As a result, the surface β-sheet, β1−β3, shifts away from the active site and moves the catalytic histidine ∼1.5 Å away from its active position (Figure 44B). The binding of L2′ in the interface of caspase-6 results in steric clashes with the elbow loop of L3, preventing its insertion

inhibitors that are highly selective. A general conformation used to affect caspase activity is the closed loop structure of L2′. In this conformation, the loop folds into the dimer interface, as one observes in the zymogen (see Figure 5A), and prevents loop bundle formation (see Figure 15). Following maturation (Cleavage 1) of the intersubunit linker, L2′ remains inserted into the dimer interface in many caspases, so the changes in active site loop structure can be assessed at several stages of maturation (zymogen-unliganded-active), and the data show a range of active site structures for the closed loop L2′ conformations, from close to the active structure (P7 in Figure 26) to close to the zymogen structure, with disordered active site loops (P8 in Figure 26). In the case of caspase-7 (Figure 42), upon cleavage of the IL, L2′ remains in the closed conformation but L3 is able to insert the “elbow” loop in the dimer interface, and there is little disruption in the surface β-sheet, β1−β3, and little movement in the catalytic histidine. In contrast to the closed conformation of procaspase-8 in the caspase-8:c-FLIPL heterodimer, described above (Figure 33), the closed conformation of caspase-7 is stabilized mostly by van der Waals interactions, so the protein lacks the hydrogen-bonding network observed in the caspase8:c-FLIPL heterodimer, which stabilizes the catalytic cysteine in the active conformation. Consequently, in the closed conformation of caspase-7, the catalytic cysteine is ∼3 Å from its active position and is similar to the position in the zymogen. 6692


Chemical Reviews

Review

Figure 44. Closed conformation of caspase-6 (A) results in disordered active site loops L2 and L3. (B) Comparison of turn 6 in the active and closed conformations. (C) Phosphorylation site on helix 5, S257. (D) D257 clashes with P201 of L2′, preventing rotation of the loop. Active caspase-6 PDB ID 3OD5,119 S257D caspase-6 PDB ID 3S8E.216 Figure 43. Comparison of caspase-1 zymogen (PDB ID 3E4C)214 and unliganded, mature (PDB ID 1SC1),215 conformations with the active conformation (PDB ID 1ICE).33

by about two turns (Figure 45C). Turn 5 extends into the substrate-binding pocket, and helix 3 moves away from the dimer interface, causing movement of helix 2 to avoid steric clashes. The substrate-binding groove is intact, but L2′ is in the closed conformation, which while preventing the loop bundle from forming between L2, L2′, and L4, may be important for stabilizing the extended helix since Y198 on L2′ hydrogen bonds with Q137 on helix 3 (Figure 45C). In addition, the extended helix is stabilized by several hydrogen-bonding interactions between active site loop L1 and helices 1, 2, and 3 (Figure 45C). The result of the transition is to trap the catalytic histidine, H121, in the inactive state, facing away from the catalytic C163. As shown in Figure 46, the residues that form the hydrogen-bonding network are not conserved in the other caspases. In particular, E53, which interacts directly with H121, is found only in caspases-6 and -1. Likewise, amino acids in the turn 6 region (β1−β3) are not conserved among the caspases (Figure 47). All caspases, with the exception of caspase-6, contain helix breaking glycine and proline residues near β3, which presumably restrict the conformational space of those caspases by preventing the extended helix formation. The introduction of helix breaking amino acids in caspase-6 resulted in a reduction in helicity,93 suggesting that removing the barriers to forming the extended helix could expand the conformational space of the other caspases as well. Similarly to the closed form of caspase-6 that is trapped by phosphorylation (Figure 44), the extended helix form of caspase-6 is trapped by the binding of zinc to an allosteric site near helix 5 (Figure 48).46 The zinc is coordinated by three residues, E244 on helix 5, K36 on the pro-domain linker, and H287 at the C-terminus. As shown in Figure 49, the amino

into the dimer interface, so L3 is disordered. Since L2 also is disordered, the loop bundle does not form between L2′, L2, and L4. Interestingly, L3 is similar to the solvent-exposed L3 loop of the caspase-1 zymogen rather than procaspase-6. As described above (Figure 19), the IL binds in the active site of the zymogen, so L3 is inserted to form the substrate-binding groove. The closed conformation of caspase-6 is trapped by phosphorylation of S257 on helix 5 (Figure 44C and 44D).216,217 In this case, steric clashes between the phosphate, which is viewed in the S257D phosphomimetic (Figure 44D), prevents rotation of L2′ out of the dimer interface, thus maintaining L3 in the disordered state. In addition to trapping the closed conformation by phosphorylation, the zymogen can be trapped in the inactive state by forming a tetramer.218 In this case, an allosteric peptide facilitated tetramerization by binding near helix 2 and β3, which maintains the inactive zymogen. It is not yet clear why the zymogen does not autoactivate in the tetramer, since the cleavage site of the intersubunit linker remains bound in the active site, but tetramerization may affect active site dynamics in a way to inhibit cleavage of the IL. Finally, the unliganded caspase-6 forms an inactive conformation in which the short β-strand, β1−β3, undergoes a coil-to-helix transition that results in extending helix 3 by nearly four turns and disrupts the active site catalytic residues (Figure 45).46,93,116 The extended helix results in steric clashes with L1 of the active conformation, so L1 shifts such that helix 1 is extended 6693


Chemical Reviews

Review

Figure 45. Extended helix conformation of caspase-6. (A) The extended helix 3 is shown in orange. (B) Transition of the β-strand, β1−β3, in the unliganded-to-active conformations (top) compared to the zymogen-to-active conformations (bottom). Specific amino acid interactions that stabilize the extended helix conformation. (D) Catalytic dyad maintained ∼9 Å from their active positions. Caspase-6 zymogen PDB ID 3NR2,119 unliganded caspase-6 PDB ID 2WDP,219 active caspase-6 PDB ID 3OD5.119

Figure 46. Comparison of amino acid residues that form a hydrogenbonding network with the catalytic H121 in the extended helix conformation of caspase-6.

Figure 47. Comparison of amino acid residues in the short surface βsheet, β1−β3, that undergoes a coil-to-helix transition in caspase-6. The catalytic histidine (H121 in caspase-6) is shown in bold. Amino acids in red form stabilizing hydrogen bonds. Helix breaking proline or glycine residues are shown in blue or green, respectively.

acids are not conserved in other caspases. As described above, the pro-domain linker is the least conserved region of the protein, so K36 is found only in caspase-6. Glutamate 244 is conserved only in the effector caspases-3, -6, and -7, while H287 is found only in caspase-6. Interestingly, the zinc-binding site in caspase-6 was shown to bind a peptide in caspase-7 (Figure 50).220 While the peptide most likely originated from E. coli during expression and

purification of the recombinant protein, the data show the potential for a common allosteric site in caspases that evolved to bind zinc, in the case of caspase-6, or peptides, in the case of 6694


Chemical Reviews

Review

landscape, as observed in the caspases, can provide a range of energetically distinctive states in the ensemble. Caspases contain a common allosteric site in the dimer interface (Figure 53A), and there are several examples of small molecules that bind to the allosteric site, two of which stabilize the zymogen-like disordered state in the ensemble. Although citrate is not an allosteric activator (Figure 53D), its binding in the interface demonstrates the possibility of developing small molecule activators by optimizing hydrogenbonding interactions in the native conformation. It appears that the population is sampled at low frequency, however, since malonate stabilizes the active state by binding in the S1 pocket of the active site (Figure 53E), suggesting that fluctuations in the active site may be more easily trapped than fluctuations in the dimer interface. Two compounds that bind in the allosteric site of the dimer interface, FICA in caspase-7 (Figure 54) and compound 34 in caspase-1 (Figure 55), inhibit the enzyme allosterically by stabilizing the zymogen-like conformation, with disordered active site loops. Together, the data suggest that the unliganded conformations lie closer to the zymogen than to the active caspase on the energy landscape. The overall results are similar for the two allosteric inhibitors, although the specific mechanisms are somewhat different. In caspase-7, the binding of FICA to the allosteric site prevents Y223 from transitioning toward the interface, as observed in the active conformation, due to steric clashes with FICA. This results in the substratebinding groove remaining solvent exposed. In caspase-1, steric clashes between compound 34 and R286, on active site loop L2, prevent R286 from transitioning to its active conformation in the dimer interface, resulting in a shift of the catalytic cysteine to the inactive conformation. Overall, the data show that the closed conformation, with disordered active sites (P5/P8 in Figure 26), is a relatively lowenergy state in the ensemble and can be exploited to inhibit the enzyme by utilizing a common allosteric site in the dimer interface. It is worth noting that in some cases, such as caspase1, the mature dimer is not stable and dissociates to the individual protomers (P9 and P10 in Figure 26).221 The potential coupling between dimerization and substrate binding adds an additional layer of complexity to the conformational landscape. 4.3.5. Post-Translational Modifications and Potential Long Distance Communications. It has been well established that caspases are modified by phosphorylation28,29,110,176,216,222−227 and glutationylation,31,44,228 but how the modification controls caspase activity under conditions of the adaptive response is not known. One should note that the catalytic cysteine thiol is also modified directly by glutathione229 or nitrosylation,230 and while the reactions are important links between caspases and the metabolic conditions of the cell, only potential allosteric mechanisms are described here. The sites of caspases that are modified by phosphorylation or glutathionylation are shown in red in Figure 6. It is quite likely that the current data regarding post-translational modifications of caspases is largely incomplete, but with available data, one can begin to see patterns emerge of potential allosteric sites on the enzyme. In some cases, phosphorylation may simply prevent zymogen maturation by inhibiting Cleavage 1 of the intersubunit linker, and several phosphorylation sites are observed in the IL near the processing sites (Figure 6). Thus, while technically those sites decrease the activity of the caspase, the mechanism is relatively straightforward since the zymogen

Figure 48. Zinc binding (red spheres) to an allosteric site in caspase-6 traps the extended helix conformation (PDB ID 4FXO).46

Figure 49. Comparison of amino acid residues that coordinate zinc in caspase-6 (red amino acids). For caspases-1, -7, and -9, amino acids in red are sites of post-translational modifications.

caspase-7, although it should be stressed that so far no function has been ascribed to peptide binding at this site in caspase-7. Mutation of six residues near the zinc-binding site of caspase6 converts the putative common allosteric site into a peptidebinding site in caspase-7 (Figure 50B). Importantly, Q260 on helix 5 and Q260′ on helix 5′, across the dimer interface, provide a cavity for binding histidine on the peptide. In caspase3, H234 and H234′, in the same position on helix 5, fill the cavity by forming interdimer charge−charge interactions with the neighboring E231, suggesting that the site may have evolved in caspase-3 for increased dimer stability. Overall, the closed complexes described here provide a large ensemble of partially ordered structures that lies between the conformations of the active enzyme, with ordered active site loops, and the zymogen, with disordered active site loops. In terms of the energy landscape model described above (Figure 23), where two low-energy ground states, A and B, are separated by a higher energy state, β, the energy of the closed states of the caspases appear to vary, depending on the level of ordering of the active site loops (Figure 51). The relative position on the energy landscape is important when considering the design of allosteric effectors to the closed conformation (Figure 52). While allosteric inhibitors may require lower binding free energy if the closed conformation exhibits more disorder in the active site loops, conversely, allosteric activators may require higher binding free energy to stabilize the active state. In this way, a common conformational 6695


Chemical Reviews

Review

Figure 50. Comparison of a putative common allosteric site in caspases. The site evolved to bind zinc in caspase-6 (A) (PDB ID 4FXO)46 or peptides in caspase-7 (B) (PDB ID 2QL9).220 In caspase-3 (C) (PDB ID 2J30),76 no ligand binding has been observed.

cleavage rather than selection of a more active state in the ensemble. After accounting for modifications in the IL, however, one observes that many other sites are modified. Rather than considering each individual modification and how it may affect the conformational selection of a specific caspase, it is beneficial to approach the modifications in a more global sense, as suggested recently by Hardy.30 The responses of caspases to individual modification events are largely unknown, with the noted exception of the phosphorylation of caspase-6, which as described above traps the protein in the closed conformation (Figure 44). At present it is not known how an individual modification affects a specific caspase, and a more complete database will require further biochemical and structural studies of the modification sites. Here, the modifications shown in Figure 6 were first mapped onto each caspase individually, as shown in Figure 56. One should note that the analysis includes six caspases and that caspase-2 is missing from the structural representation because the currently identified phosphorylation sites are either in the

Figure 51. Modification of the energy landscape of two species, A and B, in which a high-energy state, β, allows conversion of A to B through conformational selection. Differences in the closed conformations of caspases provide a range of structures with variations in the energy of β.

has little to no activity compared to the mature enzyme. Likewise, if a modification in this region increases caspase activity then most likely it is due to an increase in zymogen 6696


Chemical Reviews

Review

Figure 52. Schematic of the range of closed conformations, where the more disordered closed structures lie on the left of the continuum and the more ordered closed structures lie on the right of the continuum. Caspase-6 is the only caspase that forms the extended helix conformation.

Figure 53. Common allosteric site in the caspases. (A) The dimer interface contains an allosteric site that is common in the caspases (PDB ID 1F1J).112 (B) Binding of compound 34 to the allosteric site in caspase-1 (PDB ID 2FQQ).41 (C) Binding of FICA to the allosteric site in caspase-7 (PDB ID 1SHL).40 (D) Binding of citrate to the allosteric site in caspase-7 (PDB ID 2QL9).220 (E) Binding of malonate to the active site of caspase-1 (PDB ID 1SC3).215

Figure 54. Inhibition of caspase-7 by binding of FICA to the dimer interface (PDB ID 1SHL).40 (Top) Steric overlap of the substratebinding loop, L3, in the active conformation and L2′ in the closed complex. (Bottom) Allosteric inhibitor, FICA, binds to the allosteric site in the dimer interface and prevents L3 insertion.

Figure 6, the amino acid is conserved in all caspases (S or T) except for caspases-10 and -14. Structurally, the site is in a loop at the C-terminus of helix 3, as shown in the lower left circled region in caspases-3, -7, and -8 (Figure 56). On the basis of the individual data, it is not clear how modification of turn 9 affects conformational selection and thus enzyme activity because it is over 30 Å from the active site. The site is near helices 2 and 3, which as described above undergo

pro-domain or in the intersubunit linker, so the sites are not represented in the structure. An examination of the individual caspases shows that some sites are modified on several caspases, while other sites have been identified only on a single caspase. For example, in turn 9, S150 of caspase-3, T173 in caspase-7, and S347 in caspase-8 are phosphorylated, and, as shown in 6697


Chemical Reviews

Review

Figure 55. Inhibition of caspase-1 by binding of compound 34 to the dimer interface (PDB ID 2FQQ).41 (Top) Steric overlap of R286, on active site L2, prevents movement to the active conformation with compound 34 bound in the interface. (Bottom) The catalytic C285 remains in the zymogen-like conformation.

transitions in caspases-3 and -6 during the selection of inactive states in the ensemble. Due to the proximity to these dynamic helices, one may speculate that phosphorylation of turn 9 affects enzyme activity by influencing the transitions of helices 2 and 3. When examined globally, however, turn 9 also is part of a zinc-binding site in caspase-9,231 so the site may be functionally linked to other regions in the caspase structure. In addition, several modifications occur near the active site, but the modifications differ for each caspase (active sites positioned in the top left and bottom right in the left panel or bottom left and top right in the right panel, Figure 56). When all of the sites of modifications are mapped onto a single caspase structure, including the two known zinc-binding sites, as shown in Figure 57, the global analysis begins to provide a low-resolution picture of potential allosteric hot spots on the protein as well as possible modes of communication between the sites. As shown in Figure 57 (top), two general regions of the protein may serve as hot spots for allosteric regulation (Figure 57, top). Not surprisingly, the active site is identified as one “hot spot”, most likely because communication pathways that affect activity ultimately should connect to the active site (Figure 57, bottom). Each of the active site loops are represented in the network, including amino acids such as Y204 (caspase-3 numbering), which as described above (Figure 39) undergoes dynamic fluctuations that can be utilized to trap the amino acid in an

Figure 56. Sites of post-translational modifications and zinc-binding sites for six caspases. Modified amino acids are shown as spheres, and for clarity the sites are circled.

inactive state, and amino acids that participate in loop bundle formation between L2, L2′, and L4. The hot spot also includes the short surface β-sheet, β1−β3, that not only undergoes structural rearrangements in caspase-6 to form the extended helix conformation but also moves laterally in the caspase zymogen, due to the binding of the IL in the interface, resulting in positioning the catalytic histidine away from the active position. The second hot spot is on the opposite side of the active site, at the “bottom” of the protein as represented in Figure 57. This area includes the C-terminal regions of helices 2 and 3, on one side of the central β-sheet, the C-terminal regions of helices 1 6698


Chemical Reviews

Review

Figure 57. Sites of caspase post-translational modifications mapped onto a single structure (top). Phosphorylation and glutathionylation sites are orange, while zinc-binding sites are shown in green. (Bottom) Hot spot near the caspase active site.

and 4, and a zinc-binding site, which was also described above as a peptide binding site (Figure 58). The short N-terminal helix found on some caspases also resides in this area (Figures 7 and 8), suggesting that the pro-domain linker is part of the communication network. When viewed in this way, the single modification of S150 in turn 9 of caspase-3, for example, may be propagated to other regions of the protein through the overlapping interaction networks of the “hot spots”. Currently, there are no known protein effectors of caspases that bind to the helix-1, -4, -5 or turn 9 regions of the protein, so the modifications most likely affect activity through conformational rearrangements rather than affecting protein− protein binding interactions. It is also worth noting that the resolution of the global map will be greater with a larger modification database, particularly when linked to biochemical and structural data for the hot spots. With greater resolution, the broad features of the single hot spot may separate into wellresolved communication networks, where perturbations of amino acids in the network are propagated long distances to other networks on the protein. It is interesting to note that the analysis does not identify the dimer interface as a hot spot for allosteric regulation, because the allosteric site of the dimer interface is not modified in the cell. While the site can be utilized in selecting the inactive state

by binding small drug compounds, it is not used by the cell to modulate activity, except as noted above as a docking site for L2′ and stabilizing the closed conformation.

5. CONCLUSION While the broad outline of caspase activation is known, the molecular details of allosteric regulation are not. The active and inactive states of caspases are more accurately described as an ensemble of states, where small changes in a few amino acids or larger changes in secondary structure result in inactivation by stabilizing inactive states in the ensemble. The main goal of this review is to provide a global view of the caspase conformational landscape and mechanisms used to select states in the ensemble. The complex conformational landscape provides the cell with multiple opportunities to select various conformational states as a way to fine tune caspase activity. Admittedly, however, our understanding of how cells fine tune caspase activity in order to carry out reactions required for differentiation or development, while simultaneously preventing apoptosis, is still in the very early stages. Negative design elements in the dimer interfaces of some caspases effectively trap the monomer, leading to selective mechanisms for activation by homo- or heterodimerization and thus control over the assembly of the activating complexes. 6699


Chemical Reviews

Review

Figure 58. Sites of caspase post-translational modifications mapped onto a single structure. (Top) Modification sites at the C-termini of helices 2 and 3 (left) or the termini of helices 1, 4, and 5 on the other side of the central β-sheet. (Bottom) Hot spot near the caspase active site. Modified amino acids are shown as surface representation, and zinc-binding sites are shown in green.

Similarly, the constitutively dimeric caspases exhibit several conformations that vary in the degree of loop disorder as well as higher energy active conformations that can be selected among the population of states. Years of structural and cell biological studies have led to the design of small molecules to chemically select various states in the ensemble as well as an improved understanding of the effects of manipulating the ensemble in the cell. Continued development of these methodologies will provide a greatly improved picture of the ensemble. In the cell, the allosteric ensemble is controlled by posttranslational modifications and metal (and perhaps, ion) binding, although aside from a few well-studied examples, the effects of the modifications on conformational selection are largely unknown. With improved databases on modifications of the caspases, which will include combinations of biochemical, structural, and cell biological studies, as well as a complete understanding of metal and ion binding, the low-resolution picture of global allosteric communications and their effects on the conformational landscape will progress into an improved understanding of communication networks that affect selection in the ensemble. The current low-resolution global picture of allosteric sites afforded by the presently incomplete database will resolve into an understanding of how the sites evolved to provide specific functionality in the caspases, which in turn should provide strategies for the design of highly selective allosteric modulators. The post-translational modifications are

important mechanisms in the proliferation of several cancers, so understanding how the modifications affect conformational selection will provide potential targets for new therapies in cancer treatment.

AUTHOR INFORMATION Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest. Biography Clay Clark earned his Master’s degree (1989) in Biology from the University of San Francisco under the supervision of Dr. John Cobley and his Ph.D. degree (1994) in Biochemistry from Texas A&M University under the supervision of Dr. Thomas O. Baldwin. He conducted postdoctoral research in the Department of Biochemistry and Molecular Biophysics at Washington University with Dr. Carl Frieden. In 1999, he obtained his first independent position as Assistant Professor of Biochemistry at North Carolina State University. In 2015, he moved his laboratory to the University of Texas at Arlington. He is Professor and Chair of Biology at UT Arlington. He is on the editorial board of several journals and serves as Deputy Chair of the Biochemical Journal. More information can be found at http:// www.uta.edu/biology/clark/index.php. 6700


Chemical Reviews

Review

(25) Scott, F. L.; Denault, J.-B.; Riedl, S. J.; Shin, H.; Renatus, M.; Salvesen, G. S. XIAP Inhibits Caspase-3 and -7 Using Two Binding Sites: Evolutionarily Conserved Mechanism of IAPs. EMBO J. 2005, 24, 645−655. (26) Huang, Y.; Park, Y. C.; Rich, R. L.; Segal, D.; Myszka, D. G.; Wu, H. Structural Basis of Caspase Inhibition by XIAP: Differential Roles of the Linker versus the BIR Domain. Cell 2001, 104, 781−790. (27) Riedl, S. J.; Renatus, M.; Schwarzenbacher, R.; Zhou, Q.; Sun, C.; Fesik, S. W.; Liddington, R. C.; Salvesen, G. S. Structural Basis for the Inhibition of Caspase-3 by XIAP. Cell 2001, 104, 791−800. (28) Parrish, A.; Freel, C. D.; Kornbluth, S. Cellular Mechanisms Controlling Caspase Activation and Function. Cold Spring Harbor Perspect. Biol. 2013, 5, 1−24. (29) Kurokawa, M.; Kornbluth, S. Caspases and Kinases in a Death Grip. Cell 2009, 138, 838−854. (30) Dagbay, K.; Eron, S. J.; Serrano, B. P.; Velázquez-Delgado, E. M.; Zhao, Y.; Lin, D.; Vaidya, S.; Hardy, J. a. A Multipronged Approach for Compiling a Global Map of Allosteric Regulation in the Apoptotic Caspases. Methods Enzymol. 2014, 544, 215−249. (31) Huang, Z.; Pinto, J. T.; Deng, H.; Richie, J. P. Inhibition of Caspase-3 Activity and Activation by Protein Glutathionylation. Biochem. Pharmacol. 2008, 75, 2234−2244. (32) Walker, N. P. C.; Talanian, R. V.; Brady, K. D.; Dang, L. C.; Bump, N. J.; Ferenza, C. R.; Franklin, S.; Ghayur, T.; Hackett, M. C.; Hammill, L. D.; et al. Crystal Structure of the Cysteine Protease Interleukin-1b-Converting Enzyme: A (p20/p10)2 Homodimer. Cell 1994, 78, 343−352. (33) Wilson, K. P.; Black, J.-A. F.; Thomson, J. A.; Kim, E. E.; Griffith, J. P.; Navia, M. A.; Murcko, M. A.; Chambers, S. P.; Aldape, R. A.; Raybuck, S. A.; et al. Structure and Mechanism of Interleukin-1 B Converting Enzyme. Nature 1994, 370, 270−275. (34) Cohen, G. Caspases: The Executioners of Apoptosis. Biochem. J. 1997, 326, 1−16. (35) Fuentes-Prior, P.; Salvesen, G. S. The Protein Structures That Shape Caspase Activity, Specificity Activation and Inhibition. Biochem. J. 2004, 384, 201−232. (36) Park, H. H. Structural Features of Caspase-Activating Complexes. Int. J. Mol. Sci. 2012, 13, 4807−4818. (37) Shi, Y. Caspase Activation, Inhibition, and Reactivation: A Mechanistic View. Protein Sci. 2004, 13, 1979−1987. (38) Yan, N.; Shi, Y. Mechanisms of Apoptosis through Structural Biology. Annu. Rev. Cell Dev. Biol. 2005, 21, 35−56. (39) Datta, D.; Scheer, J. M.; Romanowski, M. J.; Wells, J. A. An Allosteric Circuit in Caspase-1. J. Mol. Biol. 2008, 381, 1157−1167. (40) Hardy, J. A.; Lam, J.; Nguyen, J. T.; O’Brien, T.; Wells, J. A. Discovery of an Allosteric Site in the Caspases. Proc. Natl. Acad. Sci. U. S. A. 2004, 101, 12461−12466. (41) Scheer, J. M.; Romanowski, M. J.; Wells, J. A. A Common Allosteric Site and Mechanism in Caspases. Proc. Natl. Acad. Sci. U. S. A. 2006, 103, 7595−7600. (42) Hardy, J. A.; Wells, J. A. Dissecting an Allosteric Switch in Caspase-7 Using Chemical and Mutational Probes. J. Biol. Chem. 2009, 284, 26063−26069. (43) Velázquez-Delgado, E. M.; Hardy, J. A. Phosphorylation Regulates Assembly of the Caspase-6 Substrate-Binding Groove. Structure 2012, 20, 742−751. (44) Meissner, F.; Molawi, K.; Zychlinsky, A. Superoxide Dismutase 1 Regulates Caspase-1 and Endotoxic Shock. Nat. Immunol. 2008, 9, 866−872. (45) Perry, D. K.; Smyth, M. J.; Stennicke, H. R.; Salvesen, G. S.; Duriez, P.; Poirier, G. G.; Hannun, Y. A. Zinc Is a Potent Inhibitor of the Apoptotic Protease, Caspase-3. A Novel Target for Zinc in the Inhibition of Apoptosis. J. Biol. Chem. 1997, 272, 18530−18533. (46) Velazquez-Delgado, E. M.; Hardy, J. A. Zinc-Mediated Allosteric Inhibition of Caspase-6. J. Biol. Chem. 2012, 287, 36000−36011. (47) Alnemri, E. S.; Livingston, D. J.; Nicholson, D. W.; Salvesen, G.; Thornberry, N. A.; Wong, W. W.; Yuan, J. Human ICE/CED-3 Protease Nomenclature. Cell 1996, 87, 171.

ACKNOWLEDGMENTS This work was supported by NIH grant GM065970. REFERENCES (1) Renehan, A. G.; Booth, C.; Potten, C. S. What Is Apoptosis, and Why Is It Important? Br. Med. J. 2001, 322, 1536−1538. (2) Edinger, A. L.; Thompson, C. B. Death by Design: Apoptosis, Necrosis and Autophagy. Curr. Opin. Cell Biol. 2004, 16, 663−669. (3) Maiuri, M. C.; Zalckvar, E.; Kimchi, A.; Kroemer, G. Self-Eating and Self-Killing: Crosstalk between Autophagy and Apoptosis. Nat. Rev. Mol. Cell Biol. 2007, 8, 741−752. (4) Martin, S. J.; Green, D. R. Protease Activation during Apoptosis: Death by a Thousand Cuts? Cell 1995, 82, 349−352. (5) Enari, M.; Sakahira, H.; Yokoyama, H.; Okawa, K.; Iwamatsu, A.; Nagata, S. A Caspase-Activated DNase That Degrades DNA during Apoptosis, and Its Inhibitor ICAD. Nature 1998, 391, 43−50. (6) Nguyen, J. T.; Wells, J. A. Direct Activation of the Apoptosis Machinery as a Mechanism to Target Cancer Cells. Proc. Natl. Acad. Sci. U. S. A. 2003, 100, 7533−7538. (7) Muzio, M.; Stockwell, B. R.; Stennicke, H. R.; Salvesen, G. S.; Dixit, V. M. An Induced Proximity Model for Caspase-8 Activation. J. Biol. Chem. 1998, 273, 2926−2930. (8) Budihardjo, I.; Oliver, H.; Lutter, M.; Luo, X.; Wang, X. Biochemical Pathways of Caspase Activation during Apoptosis. Annu. Rev. Cell Dev. Biol. 1999, 15, 269−290. (9) Shi, Y. Mechanisms of Caspase Activation and Inhibition during Apoptosis. Mol. Cell 2002, 9, 459−470. (10) Kumar, S. Mechanisms Mediating Caspase Activation in Cell Death. Cell Death Differ. 1999, 6, 1060−1066. (11) Creagh, E. M.; Conroy, H.; Martin, S. J. Caspase-Activation Pathways in Apoptosis and Immunity. Immunol. Rev. 2003, 193, 10− 21. (12) MacKenzie, S. H.; Clark, A. C. Death by Caspase Dimerization. Adv. Exp. Med. Biol. 2012, 747, 55−73. (13) Degterev, A.; Boyce, M.; Yuan, J. A Decade of Caspases. Oncogene 2003, 22, 8543−8567. (14) Denault, J.-B.; Salvesen, G. S. Caspases: Keys in the Ignition of Cell Death. Chem. Rev. 2002, 102, 4489−4499. (15) Boatright, K. M.; Salvesen, G. S. Mechanisms of Caspase Activation. Curr. Opin. Cell Biol. 2003, 15, 725−731. (16) Connolly, P. F.; Jäger, R.; Fearnhead, H. O. New Roles for Old Enzymes: Killer Caspases as the Engine of Cell Behavior Changes. Front. Physiol. 2014, 5, 1−10. (17) Hyman, B. T.; Yuan, J. Apoptotic and Non-Apoptotic Roles of Caspases in Neuronal Physiology and Pathophysiology. Nat. Rev. Neurosci. 2012, 13, 395−406. (18) Li, Z.; Sheng, M. Caspases in Synaptic Plasticity. Mol. Brain 2012, 5, 15. (19) Blander, J. M. A Long-Awaited Merger of the Pathways Mediating Host Defence and Programmed Cell Death. Nat. Rev. Immunol. 2014, 14, 601−618. (20) Matalova, E.; Lesot, H.; Svandova, E.; Vanden Berghe, T.; Sharpe, P. T.; Healy, C.; Vandenabeele, P.; Tucker, A. S. Caspase-7 Participates in Differentiation of Cells Forming Dental Hard Tissues. Dev. Growth Differ. 2013, 55, 615−621. (21) Creagh, E. M. Caspase Crosstalk: Integration of Apoptotic and Innate Immune Signalling Pathways. Trends Immunol. 2014, 35, 631− 640. (22) Tschopp, J.; Schroder, K. NLRP3 Inflammasome Activation: The Convergence of Multiple Signalling Pathways on ROS Production? Nat. Rev. Immunol. 2010, 10, 210−215. (23) Ouyang, L.; Shi, Z.; Zhao, S.; Wang, F.-T.; Zhou, T.-T.; Liu, B.; Bao, J.-K. Programmed Cell Death Pathways in Cancer: A Review of Apoptosis, Autophagy and Programmed Necrosis. Cell Proliferation 2012, 45, 487−498. (24) Shiozaki, E. N.; Shi, Y. Caspases, IAPs and Smac/DIABLO: Mechanims from Structural Biology. Trends Biochem. Sci. 2004, 29, 486−494. 6701


Chemical Reviews

Review

(48) Rawlings, N. D. MEROPS: The Peptidase Database. Nucleic Acids Res. 2006, 34 (90001), D270−D272. (49) Rawlings, N. D.; Waller, M.; Barrett, A. J.; Bateman, A. MEROPS: The Database of Proteolytic Enzymes, Their Substrates and Inhibitors. Nucleic Acids Res. 2014, 42, D503−D509. (50) Uren, A. G.; O’Rourke, K.; Aravind, L. A.; Pisabarro, M. T.; Seshagiri, S.; Koonin, E. V.; Dixit, V. M. Identification of Paracaspases and Metacaspases: Two Ancient Families of Caspase-like Proteins, One of Which Plays a Key Role in MALT Lymphoma. Mol. Cell 2000, 6, 961−967. (51) Vercammen, D.; Declercq, W.; Vandenabeele, P.; Van Breusegem, F. Are Metacaspases Caspases? J. Cell Biol. 2007, 179, 375−380. (52) Asplund-Samuelsson, J.; Bergman, B.; Larsson, J. Prokaryotic Caspase Homologs: Phylogenetic Patterns and Functional Characteristics Reveal Considerable Diversity. PLoS One 2012, 7, 1−14. (53) Choi, C. J.; Berges, J. A. New Types of Metacaspases in Phytoplankton Reveal Diverse Origins of Cell Death Proteases. Cell Death Dis. 2013, 4, e490. (54) Stennicke, H. R.; Salvesen, G. S. Catalytic Properties of the Caspases. Cell Death Differ. 1999, 6, 1054−1059. (55) Miscione, G. P.; Calvaresi, M.; Bottoni, A. Computational Evidence for the Catalytic Mechanism of Caspase-7. A DFT Investigation. J. Phys. Chem. B 2010, 114, 4637−4645. (56) Sulpizi, M.; Rothlisberger, U.; Carloni, P. Molecular Dynamics Studies of Caspase-3. Biophys. J. 2003, 84, 2207−2215. (57) Denecker, G.; Ovaere, P.; Vandenabeele, P.; Declercq, W. Caspase-14 Reveals Its Secrets. J. Cell Biol. 2008, 180, 451−458. (58) Peter, M. E.; Krammer, P. H. The CD95(APO-1/Fas) DISC and Beyond. Cell Death Differ. 2003, 10, 26. (59) Weber, C. H.; Vincenz, C. A Docking Model of Key Components of the DISC Complex: Death Domain Superfamily Interactions Redefined. FEBS Lett. 2001, 492, 171−176. (60) Hughes, M. A.; Harper, N.; Butterworth, M.; Cain, K.; Cohen, G. M.; MacFarlane, M. Reconstitution of the Death-Inducing Signaling Complex Reveals a Substrate Switch That Determines CD95Mediated Death or Survival. Mol. Cell 2009, 35, 265−279. (61) Cain, K.; Bratton, S. B.; Langlais, C.; Walker, G.; Brown, D. G.; Sun, X.-M.; Cohen, G. M. Apaf-1 Oligomerizes into Biologically Active ∼ 700-kDa and Inactive ∼ 1.4-MDa Apoptosome Complexes. J. Biol. Chem. 2000, 275, 6067−6070. (62) Adams, J. M.; Cory, S. Apoptosomes: Engines for Caspase Activation. Curr. Opin. Cell Biol. 2002, 14, 715−720. (63) Riedl, S. J.; Salvesen, G. S. The Apoptosome: Signalling Platform of Cell Death. Nat. Rev. Mol. Cell Biol. 2007, 8, 405−413. (64) Bao, Q.; Shi, Y. Apoptosome: A Platform for the Activation of Initiator Caspases. Cell Death Differ. 2007, 14, 56−65. (65) Pop, C.; Timmer, J.; Sperandio, S.; Salvesen, G. S. The Apoptosome Activates Caspase-9 by Dimerization. Mol. Cell 2006, 22, 269−275. (66) Bratton, S. B.; Walker, G.; Srinivasula, S. M.; Sun, X.-M.; Butterworth, M.; Alnemri, E. S.; Cohen, G. M. Recruitment, Activation and Retention of Caspases-9 and -3 by Apaf-1 Apoptosome and Associated XIAP Complexes. EMBO J. 2001, 20, 998−1009. (67) Tinel, A.; Tschopp, J. The PIDDosome, a Protein Complex Implicated in Activation of Caspase-2 in Response to Genotoxic Stress. Science 2004, 304, 843−846. (68) MacKenzie, S.; Clark, A. C. Death by Caspase Dimerization. In Protein Dimerization and Oligomerization in Biology; Matthews, J. M., Ed.; Landes Bioscience: Autsin, TX, 2012; pp 55−73. (69) Muzio, M.; Chinnaiyan, A. M.; Kischkel, F. C.; O’Rourke, K.; Shevchenko, A.; Ni, J.; Scaffidi, C.; Bretz, J. D.; Zhang, M.; Gentz, R.; et al. FLICE, a Novel FADD-Homologous ICE/CED-3-like Protease, Is Recruited to the CD95(Fas/APO-1) Death-Inducing Signaling Complex. Cell 1996, 85, 817−827. (70) Medema, J. P.; Scaffidi, C.; Kischkel, F. C.; Shevchenko, A.; Mann, M.; Krammer, P. H.; Peter, M. E. FLICE Is Activated by Association with the CD95 Death-Inducing Signaling Complex (DISC). EMBO J. 1997, 16, 2794−2804.

(71) Ashkenazi, A.; Dixit, V. M. Death Receptors: Signaling and Modulation. Science 1998, 281, 1305−1308. (72) Oberst, A.; Pop, C.; Tremblay, A. G.; Blais, V.; Denault, J.-B.; Salvesen, G. S.; Green, D. R. Inducible Dimerization and Inducible Cleavage Reveal a Requirement for Both Processes in Caspase-8 Activation. J. Biol. Chem. 2010, 285, 16632−16642. (73) Chu, Z.-L.; Pio, F.; Xie, Z.; Welsh, K.; Krajewska, M.; Krajewski, S.; Godzik, A.; Reed, J. C. A Novel Enhancer of the Apaf1 Apoptosome Involved in Cytochrome c-Dependent Caspase Activation and Apoptosis. J. Biol. Chem. 2001, 276, 9239−9245. (74) Talanian, R. V.; Quinlan, C.; Trautz, S.; Hackett, M. C.; Mankovich, J. A.; Banach, D.; Ghayur, T.; Brady, K. D.; Wong, W. W. Substrate Specificites of Caspase Family Proteases. J. Biol. Chem. 1997, 272, 9677−9682. (75) Kitevska, T.; Roberts, S. J.; Pantaki-Eimany, D.; Boyd, S. E.; Scott, F. L.; Hawkins, C. J. Analysis of the Minimal Specificity of Caspase-2 and Identification of Ac-VDTTD-AFC as a Caspase-2Selective Peptide Substrate. Biosci. Rep. 2014, 34, 127−137. (76) Feeney, B.; Pop, C.; Swartz, P.; Mattos, C.; Clark, A. C. Role of Loop Bundle Hydrogen Bonds in the Maturation and Activity of (pro)caspase-3. Biochemistry 2006, 45, 13249−13263. (77) Earnshaw, W. C.; Martins, L. M.; Kaufmann, S. H. Mammalian Caspases: Structure, Activation, Substrates, and Functions during Apoptosis. Annu. Rev. Biochem. 1999, 68, 383−424. (78) Strasser, A.; O’Connor, L.; Dixit, V. M. Apoptosis Signaling. Annu. Rev. Biochem. 2000, 69, 217−245. (79) Mariathasan, S.; Monack, D. M. Inflammasome Adaptors and Sensors: Intracellular Regulators of Infection and Inflammation. Nat. Rev. Immunol. 2007, 7, 31−40. (80) Matusiak, M. CARD- and Pyrin-Only Proteins Regulating Inflammasome Activation and Immunity. Immunol. Rev. 2015, 265, 217−230. (81) Park, H. H.; Lo, Y.-C.; Lin, S.-C.; Wang, L.; Yang, J. K.; Wu, H. The Death Domain Superfamily in Intracellular Signaling of Apoptosis and Inflammation. Annu. Rev. Immunol. 2007, 25, 561−586. (82) Pop, C.; Chen, Y.-R.; Smith, B.; Bose, K.; Bobay, B.; Tripathy, A.; Franzen, S.; Clark, A. C. Removal of the pro-Domain Does Not Affect the Conformation of the Procaspase-3 Dimer. Biochemistry 2001, 40, 14224−14235. (83) Cain, K.; Brown, D. G.; Langlais, C.; Cohen, G. M. Caspase Activation Involves the Formation of the Aposome, a Large (∼700 kDa) Caspase-Activating Complex. J. Biol. Chem. 1999, 274, 22686− 22692. (84) Feeney, B.; Clark, A. C. Reassembly of Active Caspase-3 Is Facilitated by the Propeptide. J. Biol. Chem. 2005, 280, 39772−39785. (85) Milam, S. L.; Clark, A. C. Folding and Assembly Kinetics of Procaspase-3. Protein Sci. 2009, 18, 2500−2517. (86) Voss, O.; Batra, S.; Kolattukudy, S. J.; Gonzalez-Mejia, M. E.; Smith, J. B.; Doseff, A. I. Binding of Caspase-3 Prodomain to Heat Shock Protein 27 Regulates Monocyte Apoptosis by Inhibiting Caspase-3 Proteolytic Activation. J. Biol. Chem. 2007, 282, 25088− 25099. (87) Boucher, D.; Blais, V.; Denault, J.-B. Caspase-7 Uses an Exosite to Promote poly(ADP Ribose) Polymerase 1 Proteolysis. Proc. Natl. Acad. Sci. U. S. A. 2012, 109, 5669−5674. (88) Riedl, S. J.; Fuentes-Prior, P.; Renatus, M.; Kairies, N.; Krapp, S.; Huber, R.; Salvesen, G. S.; Bode, W. Structural Basis for the Activation of Human Procaspase-7. Proc. Natl. Acad. Sci. U. S. A. 2001, 98, 14790−14795. (89) Aravind, L.; Koonin, E. V. Classification of the CaspaseHemoglobinase Fold: Detection of New Families and Implications for the Origin of the Eukaryotic Separins. Proteins: Struct., Funct., Genet. 2002, 46, 355−367. (90) McLuskey, K.; Mottram, J. C. Comparative Structural Analysis of the Caspase Family with Other Clan CD Cysteine Peptidases. Biochem. J. 2015, 466, 219−232. (91) Witkowski, W. A.; Hardy, J. A. L2′ Loop Is Critical for Caspase7 Active Site Formation. Protein Sci. 2009, 18, 1459−1468. 6702


Chemical Reviews

Review

(92) Walters, J.; Schipper, J. L.; Swartz, P.; Mattos, C.; Clark, A. C. Allosteric Modulation of Caspase-3 through Mutagenesis. Biosci. Rep. 2012, 32, 401−411. (93) Vaidya, S.; Hardy, J. A. Caspase-6 Latent State Stability Relies on Helical Propensity. Biochemistry 2011, 50, 3282−3287. (94) Consortium, U. The Universal Protein Resource. Nucleic Acids Res. 2008, 36, D190−D195. (95) Thornberry, N. A.; Bull, H. G.; Calaycay, J. R.; Chapman, K. T.; Howard, A. D.; Kostura, M. J.; Miller, D. K.; Molineaux, S. M.; Weidner, J. R.; Aunins, J.; et al. A Novel Heterodimeric Cysteine Protease Is Required for Interleukin-1b Processing in Monocytes. Nature 1992, 356, 768−774. (96) Wang, L.; Miura, M.; Bergeron, L.; Zhu, H.; Yuan, J. Ich-1, an Ice/ced-3-Related Gene, Encodes Both Positive and Negative Regulators of Programmed Cell Death. Cell 1994, 78, 739−750. (97) Fernandes-Alnemri, T.; Litwack, G.; Alnemri, E. S. CPP32, a Novel Human Apoptotic Protein with Homology to Caenorhabditis Elegans Cell Death Protein Ced-3 and Mammalian Interleukin-1 BetaConverting Enzyme. J. Biol. Chem. 1994, 269, 30761−30764. (98) Tewari, M.; Quan, L. T.; O’Rourke, K.; Desnoyers, S.; Zeng, Z.; Beidler, D. R.; Poirier, G. G.; Salvesen, G. S.; Dixit, V. M. Yama/ CPP32 Beta, a Mammalian Homolog of CED-3, Is a CrmA-Inhibitable Protease That Cleaves the Death Substrate poly(ADP-Ribose) Polymerase. Cell 1995, 81, 801−809. (99) Faucheu, C.; Diu, A.; Chan, A. W. E.; Blanchet, A.-M.; Miossec, C.; Herve, F.; Collard-Dutilleul, V.; Gu, Y.; Aldape, R. A.; Lippke, J. A.; et al. A Novel Human Protease Similar to the Interleukin-1b Converting Enzyme Induces Apoptosis in Transfected Cells. EMBO J. 1995, 14, 1914−1922. (100) Munday, N. a.; Vaillancourt, J. P.; Ali, A.; Casano, F. J.; Miller, D. K.; Molineaux, S. M.; Yamin, T. T.; Yu, V. L.; Nicholson, D. W. Molecular Cloning and pro-Apoptotic Activity of ICErelII and ICErelIII, Members of the ICE/CED-3 Family of Cysteine Proteases. J. Biol. Chem. 1995, 270, 15870−15876. (101) Fernandes-Alnemri, T.; Litwack, G.; Alnemri, E. S. Mch2, a New Member of the Apoptotic Ced-3/Ice Cysteine Protease Gene Family. Cancer Res. 1995, 55, 2737−2742. (102) Fernandes-Alnemri, T.; Takahashi, A.; Armstrong, R.; Krebs, J.; Fritz, L.; Tomaselli, K. J.; Wang, L.; Yu, Z.; Croce, C. M.; Salveson, G.; et al. Mch3, a Novel Human Apoptotic Cysteine Protease Highly Related to CPP32 Advances in Brief Mch3, a Novel Human Apoptotic Cysteine Protease Highly Related to CPP321. Cancer Res. 1995, 55, 6045−6052. (103) Duan, H.; Chinnaiyan, A. M.; Hudson, P. L.; Wing, J. P.; He, W.-W.; Dixit, V. M. ICE-LAP3, a Novel Mammalian Homologue of the Caenorhabditis Elegans Cell Death Protein Ced-3 Is Activated during Fas- and Tumor Necrosis Factor-Induced Apoptosis. J. Biol. Chem. 1996, 271, 1621−1625. (104) Duan, H.; Orth, K.; Chinnaiyan, A. M.; Poirier, G. G.; Froelich, C. J.; He, W.-W.; Dixit, V. M. ICE-LAP6, a Novel Member of the ICE/ Ced-3 Gene Family, Is Activated by the Cytotoxic T Cell Protease Granzyme B. J. Biol. Chem. 1996, 271, 16720−16724. (105) Fernandes-Alnemri, T.; Armstrong, R. C.; Krebs, J.; Srinivasula, S. M.; Wang, L.; Bullrich, F.; Fritz, L. C.; Trapani, J. A.; Tomaselli, K. J.; Litwack, G.; et al. In Vitro Activation of CPP32 and Mch3 by Mch4, a Novel Human Apoptotic Cysteine Protease Containing Two FADDlike Domains. Proc. Natl. Acad. Sci. U. S. A. 1996, 93, 7464−7469. (106) Dixit, V. M. Fas-Associated Death Domain Protein Interleukin1beta -Converting Enzyme 2(FLICE2), an ICE/Ced-3 Homologue, Is Proximally Involved in CD95- and p55-Mediated Death Signaling. J. Biol. Chem. 1997, 272, 6578−6583. (107) Eckhart, L.; Ban, J.; Fischer, H.; Tschachler, E. Caspase-14: Analysis of Gene Structure and mRNA Expression during Keratinocyte Differentiation. Biochem. Biophys. Res. Commun. 2000, 277, 655−659. (108) Denault, J.-B.; Salvesen, G. S. Human Caspase-7 Activity and Regulation by Its N-Terminal Peptide. J. Biol. Chem. 2003, 278, 34042−34050. (109) Allan, L.; Clarke, P. R. Apoptosis and Autophagy: Regulation of Caspase-9 by Phosphorylation. FEBS J. 2009, 276, 6063−6073.

(110) Fu, Y.; Yang, G.; Zhu, F.; Peng, C.; Li, W.; Li, H.; Kim, H.-G.; Bode, A. M.; Dong, Z. Antioxidants Decrease the Apoptotic Effect of 5-Fu in Colon Cancer by Regulating Src-Dependent Caspase-7 Phosphorylation. Cell Death Dis. 2014, 5, e983. (111) Renatus, M.; Stennicke, H. R.; Scott, F. L.; Liddington, R. C.; Salvesen, G. S. Dimer Formation Drives the Activation of the Cell Death Protease Caspase 9. Proc. Natl. Acad. Sci. U. S. A. 2001, 98, 14250−14255. (112) Wei, Y.; Fox, T.; Chambers, S. P.; Sintchak, J.; Coll, J. T.; Golec, J. M. C.; Swenson, L.; Wilson, K. P.; Charifson, P. S. The Structures of Caspases-1, -3, -7 and -8 Reveal a Basis for Substrate and Inhibitor Selectivity. Chem. Biol. 2000, 7, 423−432. (113) Watt, W.; Koeplinger, K. A.; Mildner, A. M.; Heinrikson, R. L.; Tomasselli, A. G.; Watenpaugh, K. D. The Atomic-Resolution Structure of Human Capase-8, a Key Activator of Apoptosis. Structure 1999, 7, 1135−1143. (114) Cade, C.; Swartz, P.; MacKenzie, S. H.; Clark, A. C. Modifying Caspase-3 Activity by Altering Allosteric Networks. Biochemistry 2014, 53, 7582−7595. (115) Roy, S.; Bayly, C. I.; Gareau, Y.; Houtzager, V. M.; Kargman, S.; Keen, S. L.; Rowland, K.; Seiden, I. M.; Thornberry, N. A.; Nicholson, D. W. Maintenance of Caspase-3 Proenzyme Dormancy by an Intrinsic “Safety Catch” Regulatory Tripeptide. Proc. Natl. Acad. Sci. U. S. A. 2001, 98, 6132−6137. (116) Vaidya, S.; Velázquez-Delgado, E. M.; Abbruzzese, G.; Hardy, J. A. Substrate-Induced Conformational Changes Occur in All Cleaved Forms of Caspase-6. J. Mol. Biol. 2011, 406, 75−91. (117) Thomsen, N. D.; Koerber, J. T.; Wells, J. A. Structural Snapshots Reveal Distinct Mechanisms of Procaspase-3 and -7 Activation. Proc. Natl. Acad. Sci. U. S. A. 2013, 110, 8477−8482. (118) Chai, J.; Wu, Q.; Shiozaki, E.; Srinivasula, S. M.; Alnemri, E. S.; Shi, Y. Crystal Structure of a Procaspase-7 Zymogen: Mechanisms of Activation and Substrate Binding. Cell 2001, 107, 399−407. (119) Wang, X.-J.; Cao, Q.; Liu, X.; Wang, K.-T.; Mi, W.; Zhang, Y.; Li, L.-F.; LeBlanc, A. C.; Su, X.-D. Crystal Structures of Human Caspase 6 Reveal a New Mechanism for Intramolecular Cleavage SelfActivation. EMBO Rep. 2010, 11, 841−847. (120) MacKenzie, S. H.; Schipper, J. L.; England, E. J.; Thomas, M. E., III; Blackburn, K.; Swartz, P.; Clark, A. C. Lengthening the Intersubunit Linker of Procaspase 3 Leads to Constitutive Activation. Biochemistry 2013, 52, 6219−6321. (121) Pop, C.; Feeney, B.; Tripathy, A.; Clark, A. C. Mutations in the Procaspase-3 Dimer Interface Affect the Activity of the Zymogen. Biochemistry 2003, 42, 12311−12320. (122) Walters, J.; Pop, C.; Scott, F. L.; Drag, M.; Swartz, P.; Mattos, C.; Salvesen, G. S.; Clark, A. C. A Consitutively Active and Uninhibitable Caspase-3 Zymogen Efficiently Induces Apoptosis. Biochem. J. 2009, 424, 335−345. (123) Schipper, J. L.; MacKenzie, S. H.; Sharma, A.; Clark, A. C. A Bifunctional Allosteric Site in the Dimer Interface of Procaspase-3. Biophys. Chem. 2011, 159, 100−109. (124) Keller, N.; Mares, J.; Zerbe, O.; Grutter, M. G. Structural and Biochemical Studies on Procaspase-8: New Insights on Initiator Caspase Activation. Structure 2009, 17, 438−448. (125) Klaiman, G.; Champagne, N.; LeBlanc, A. C. Self-Activation of Caspase-6 in Vitro and in Vivo: Caspase-6 Activation Does Not Induce Cell Death in HEK293T Cells. Biochim. Biophys. Acta, Mol. Cell Res. 2009, 1793, 592−601. (126) Monod, J.; Jacob, F. Teleonomic Mechanisms in Cellular Metabolism, Growth, and Differentiation. Cold Spring Harbor Symp. Quant. Biol. 1961, 26, 389−401. (127) Changeux, J. P. 50th Anniversary of the Word “Allosteric. Protein Sci. 2011, 20, 1119−1124. (128) Hilser, V. J.; Wrabl, J. O.; Motlagh, H. N. Structural and Energetic Basis of Allostery. Annu. Rev. Biophys. 2012, 41, 585−609. (129) Tsai, C. J.; Nussinov, R. A Unified View of “How Allostery Works. PLoS Comput. Biol. 2014, 10, 1−12. (130) Sharp, K. Allostery in the Lac Operon: Population Selection or Induced Dissociation? Biophys. Chem. 2011, 159, 66−72. 6703


Chemical Reviews

Review

(131) Wright, P. E.; Dyson, H. J. Intrinsically Disordered Proteins in Cellular Signalling and Regulation. Nat. Rev. Mol. Cell Biol. 2015, 16, 18−29. (132) Motlagh, H.; Wrabl, J.; Li, J.; Hilser, V. The Ensemble Nature of Allostery. Nature 2014, 508, 331−339. (133) Hilser, V. J. An Ensemble View of Allostery. Science 2010, 327, 653−654. (134) Nussinov, R.; Tsai, C.-J. Allostery in Disease and in Drug Discovery. Cell 2013, 153, 293−305. (135) Mackinnon, J. A. G.; Gallastegui, N.; Osguthorpe, D. J.; Hagler, A. T.; Estébanez-Perpiña,́ E. Allosteric Mechanisms of Nuclear Receptors: Insights from Computational Simulations. Mol. Cell. Endocrinol. 2014, 393, 75−82. (136) Liu, J.; Nussinov, R. The Role of Allostery in the UbiquitinProteasome System. Crit. Rev. Biochem. Mol. Biol. 2013, 48, 89−97. (137) Davydov, D. R.; Baas, B. J.; Sligar, S. G.; Halpert, J. R. Allosteric Mechanisms in Cytochrome P450 3A4 Studied by HighPressure Spectroscopy: Pivotal Role of Substrate-Induced Changes in the Accessibility and Degree of Hydration of the Heme Pocket. Biochemistry 2007, 46, 7852−7864. (138) Colombo, M. F.; Rau, D. C.; Parsegian, V. A. Protein Solvation in Allosteric Regulaton: A Water Effect on Hemoglobin. Science 1992, 256, 655−659. (139) Colombo, M. F.; Seixas, F. A. V. Novel Allosteric Conformation of Human HB Revealed by the Hydration and Anion Effects on O2 Binding. Biochemistry 1999, 38, 11741−11748. (140) Salvay, A. G.; Grigera, J. R.; Colombo, M. F. The Role of Hydration on the Mechanism of Allosteric Regulation: In Situ Measurements of the Oxygen-Linked Kinetics of Water Binding to Hemoglobin. Biophys. J. 2003, 84, 564−570. (141) Ayers, S. D.; Nedrow, K. L.; Gillilan, R. E.; Noy, N. Continuous Nucleocytoplasmic Shuttling Underlies Transcriptional Activation of PPAR Gamma by FABP4. Biochemistry 2007, 46, 6744− 6752. (142) Zhuravlev, P. I.; Papoian, G. Protein Functional Landscapes, Dynamics, Allostery: A Tortuous Path towards a Universal Theoretical Framework. Q. Rev. Biophys. 2010, 43, 295−332. (143) Gardino, A. K.; Villali, J.; Kivenson, A.; Lei, M.; Liu, C. F.; Steindel, P.; Eisenmesser, E. Z.; Labeikovsky, W.; Wolf-Watz, M.; Clarkson, M. W.; et al. Transient Non-Native Hydrogen Bonds Promote Activation of a Signaling Protein. Cell 2009, 139, 1109−1118. (144) Boehr, D. D. During Transitions Proteins Make Fleeting Bonds. Cell 2009, 139, 1049−1051. (145) Nussinov, R.; Ma, B.; Tsai, C. J. Multiple Conformational Selection and Induced Fit Events Take Place in Allosteric Propagation. Biophys. Chem. 2014, 186, 22−30. (146) Moorthy, B. S.; Anand, G. S. Multistate Allostery in Response Regulators: Phosphorylation and Mutagenesis Activate RegA via Alternate Modes. J. Mol. Biol. 2012, 417, 468−487. (147) Silva, D. A.; Bowman, G. R.; Sosa-Peinado, A.; Huang, X. A Role for Both Conformational Selection and Induced Fit in Ligand Binding by the Lao Protein. PLoS Comput. Biol. 2011, 7, 1−11. (148) Zhuravleva, A.; Gierasch, L. M. Substrate-Binding Domain Conformational Dynamics Mediate Hsp70 Allostery. Proc. Natl. Acad. Sci. U. S. A. 2015, 112, E2865−E2873. (149) Fenton, A. W. Allostery: An Illustrated Definition for the “Second Secret of Life. Trends Biochem. Sci. 2008, 33, 420−425. (150) Koshland, E. Application of a Theory of Enzyme Specificity To Protein. Proc. Natl. Acad. Sci. U. S. A. 1958, 44, 98−104. (151) Vogt, A. D.; Pozzi, N.; Chen, Z.; Di Cera, E. Essential Role of Conformational Selection in Ligand Binding. Biophys. Chem. 2014, 186, 13−21. (152) Ma, B.; Kumar, S.; Tsai, C. J.; Nussinov, R. Folding Funnels and Binding Mechanisms. Protein Eng., Des. Sel. 1999, 12, 713−720. (153) Daniels, K. G.; Suo, Y.; Oas, T. G. Conformational Kinetics Reveals Affinities of Protein Conformational States. Proc. Natl. Acad. Sci. U. S. A. 2015, 112, 9352−9357.

(154) Ma, L.; Cui, Q. Activation Mechanism of a Signaling Protein at Atomic Resolution from Advanced Computations. J. Am. Chem. Soc. 2007, 129, 10261−10268. (155) Lee, S. Y.; Cho, H. S.; Pelton, J. G.; Yan, D.; Berry, E. A.; Wemmer, D. E. Crystal Structure of Activated CheY: Comparison with Other Activated Receiver Domains. J. Biol. Chem. 2001, 276, 16425− 16431. (156) Amor, B.; Yaliraki, S. N.; Woscholski, R.; Barahona, M. Uncovering Allosteric Pathways in Caspase-1 with Markov Transient Analysis and Multiscale Community Detection. Mol. BioSyst. 2014, 10, 2247−2258. (157) Cho, H. S.; Lee, S. Y.; Yan, D.; Pan, X.; Parkinson, J. S.; Kustu, S.; Wemmer, D. E.; Pelton, J. G. NMR Structure of Activated CheY. J. Mol. Biol. 2000, 297, 543−551. (158) Lockless, S. W.; Ranganathan, R. Evolutionarily Conserved Pathways of Energetic Connectivity in Protein Families. Science 1999, 286, 295−299. (159) Voss, O. H.; Batra, S.; Kolattukudy, S. J.; Gonzalez-Mejia, M. E.; Smith, J. B.; Doseff, A. I. Binding of Caspase-3 Prodomain to Heat Shock Protien 27 Regulates Monocyte Apoptosis by Inhibiting Caspase-3 Proteolytic Activation. J. Biol. Chem. 2007, 282, 25088− 25099. (160) Onuchic, J. N.; Wolynes, P. G.; Luthey-Schulten, Z.; Socci, N. D. Toward an Outline of the Topography of a Realistic ProteinFolding Funnel. Proc. Natl. Acad. Sci. U. S. A. 1995, 92, 3626−3630. (161) Li, W.; Wang, W.; Takada, S. Energy Landscape Views for Interplays among Folding, Binding, and Allostery of Calmodulin Domains. Proc. Natl. Acad. Sci. U. S. A. 2014, 111, 10550−10555. (162) Bose, K.; Clark, A. C. Dimeric Procaspase-3 Unfolds via a Four-State Equilibrium Process. Biochemistry 2001, 40, 14236−14242. (163) Feeney, B.; Pop, C.; Tripathy, A.; Clark, A. C. Ionic Interactions near Loop L4 Are Important for Maintaining the Active Site Environment and the Dimer Stability of (pro)caspase-3. Biochem. J. 2004, 384, 515−525. (164) Walters, J.; Swartz, P.; Mattos, C.; Clark, A. C. Thermodynamic, Enzymatic and Structural Effects of Removing a Salt Bridge at the Base of Loop 4 in (pro)caspase-3. Arch. Biochem. Biophys. 2011, 508, 31−38. (165) MacKenzie, S. H.; Clark, A. C. Slow Folding and Assembly of a Procaspase-3 Interface Variant. Biochemistry 2013, 52, 3415−3427. (166) Mittl, P. R. E.; DiMarco, S.; Krebs, J. F.; Bai, X.; Karanewsky, D. S.; Priestle, J. P.; Tomaselli, K. J.; Grutter, M. G. Structure of Recombinant Human CPP32 in Complex with the Tetrapeptide Acetyl-Asp-Val-Ala-Asp Fluoromethyl Ketone. J. Biol. Chem. 1997, 272, 6539−6547. (167) Walters, J.; Milam, S. L.; Clark, A. C. Practical Approaches to Protein Folding and Assembly. Spectroscopic Strategies in Thermodynamics and Kinetics. Methods Enzymol. 2009, 455, 1−39. (168) Paduch, M.; Koide, A.; Uysal, S.; Rizk, S.; Koide, S.; Kossiakoff, A. A. Generating Conformation-Specific Synthetic Antibodies to Trap Proteins in Selected Functional States. Methods 2013, 60, 3−14. (169) Pop, C.; Fitzgerald, P.; Green, D. R.; Salvesen, G. S. Role of Proteolysis in Caspase-8 Activation and Stabilization. Biochemistry 2007, 46, 4398−4407. (170) Ma, C.; MacKenzie, S. H.; Clark, A. C. Redesigning the Procaspase-8 Dimer Interface for Improved Dimerization. Protein Sci. 2014, 23, 442−453. (171) Richardson, J. S.; Richardson, D. C. Natural B-Sheet Proteins Use Negative Design to Avoid Edge-to-Edge Aggregation. Proc. Natl. Acad. Sci. U. S. A. 2002, 99, 2754−2759. (172) Lamkanfi, M.; Declercq, W.; Kalai, M.; Saelens, X.; Vandenabeele, P. Alice in Caspase Land. A Phylogenetic Analysis of Caspases from Worm to Man. Cell Death Differ. 2002, 9, 358−361. (173) Oberst, A.; Bender, C.; Green, D. R. Living with Death: The Evolution of the Mitochondrial Pathway of Apoptosis in Animals. Cell Death Differ. 2008, 15, 1139−1146. (174) Muñoz-Pinedo, C. Signaling Pathways That Regulate Life and Cell Death: Evolution of Apoptosis in the Context of Self-Defense. Adv. Exp. Med. Biol. 2012, 738, 124−143. 6704


Chemical Reviews

Review

(175) Chang, D. W.; Xing, Z.; Pan, Y.; Algeciras-Schimnich, A.; Barnhart, B. C.; Yaish-Ohad, S.; Peter, M. E.; Yang, X. C-FLIPL Is a Dual Function Regulator for Caspase-8 Activation and CD95Mediated Apoptosis. EMBO J. 2002, 21, 3704−3714. (176) Alvarado-Kristensson, M.; Melander, F.; Leandersson, K.; Rönnstrand, L.; Wernstedt, C.; Andersson, T. p38-MAPK Signals Survival by Phosphorylation of Caspase-8 and Caspase-3 in Human Neutrophils. J. Exp. Med. 2004, 199, 449−458. (177) Matthess, Y.; Raab, M.; Knecht, R.; Becker, S.; Strebhardt, K. Sequential Cdk1 and Plk1 Phosphorylation of Caspase-8 Triggers Apoptotic Cell Death during Mitosis. Mol. Oncol. 2014, 8, 596−608. (178) Hopkins-donaldson, S.; Bodmer, J.; Bourloud, K. B.; Apoptosis, A. L.; Brognara, C. B. Loss of Caspase-8 Expression in Highly Malignant Human Neuroblastoma Cells Correlates with Resistance to Tumor Necrosis Factor-Related Apoptosis-Inducing Ligand-Induced Apoptosis Advances in Brief Loss of Caspase-8 Expression in Highly Malignant Human Neu. Cancer Res. 2000, 60, 4315−4319. (179) Eggert, A.; Grotzer, M. A.; Zuzak, T. J.; Wiewrodt, B. R.; Ikegaki, N.; Brodeur, G. M. Resistance to TRAIL-Induced Apoptosis in Neuroblastoma Cells Correlates with a Loss of Caspase-8 Expression. Med. Pediatr. Oncol. 2000, 35, 603−607. (180) Grotzer, M. a; Eggert, A.; Zuzak, T. J.; Janss, a J.; Marwaha, S.; Wiewrodt, B. R.; Ikegaki, N.; Brodeur, G. M.; Phillips, P. C. Resistance to TRAIL-Induced Apoptosis in Primitive Neuroectodermal Brain Tumor Cells Correlates with a Loss of Caspase-8 Expression. Oncogene 2000, 19, 4604−4610. (181) Zuzak, T. J.; Steinhoff, D. F.; Sutton, L. N.; Phillips, P. C.; Eggert, A.; Grotzer, M. a. Loss of Caspase-8 mRNA Expression Is Common in Childhood Primitive Neuroectodermal Brain Tumour/ medulloblastoma. Eur. J. Cancer 2002, 38, 83−91. (182) Soung, Y. H.; Lee, J. W.; Kim, S. Y.; Jang, J.; Park, Y. G. CASPASE-8 Gene Is Inactivated by Somatic Mutations in Gastric Carcinomas. Cancer Res. 2005, 65, 815−821. (183) Mandruzzato, S.; Brasseur, F.; Andry, G.; Boon, T.; van der Bruggen, P. A CASP-8 Mutation Recognized by Cytolytic T Lymphocytes on a Human Head and Neck Carcinoma. J. Exp. Med. 1997, 186, 785−793. (184) Schafer, Z. T.; Kornbluth, S. The Apoptosome: Physiological, Developmental, and Pathological Modes of Regulation. Dev. Cell 2006, 10, 549−561. (185) Olsson, M.; Zhivotovsky, B. Caspases and Cancer. Cell Death Differ. 2011, 18, 1441−1449. (186) Crowder, R. N.; El-Deiry, W. S. Caspase-8 Regulation of TrailMediated Cell Death. Exp. Oncol. 2012, 34, 160−164. (187) Hassan, M.; Watari, H.; Abualmaaty, A.; Ohba, Y.; Sakuragi, N. Apoptosis and Molecular Targeting Therapy in Cancer. BioMed Res. Int. 2014, 2014, 1−23. (188) Indran, I. R.; Tufo, G.; Pervaiz, S.; Brenner, C. Recent Advances in Apoptosis, Mitochondria and Drug Resistance in Cancer Cells. Biochim. Biophys. Acta, Bioenerg. 2011, 1807, 735−745. (189) Putt, K. S.; Chen, G. W.; Pearson, J. M.; Sandhorst, J. S.; Hoagland, M. S.; Kwon, J.-T.; Hwang, S.-K.; Jin, H.; Churchwell, M. I.; Cho, H.-H.; et al. Small-Molecule Activation of Procaspase-3 to Caspase-3 as a Personalized Anticancer Strategy. Nat. Chem. Biol. 2006, 2, 543−550. (190) Wolan, D. W.; Zorn, J. A.; Gray, D. C.; Wells, J. A. SmallMolecule Activators of a Proenzyme. Science 2009, 326, 853−858. (191) Zorn, J. A.; Wolan, D. W.; Agard, N. J.; Wells, J. A. Fibrils Colocalize Caspase-3 with Procaspase-3 to Foster Maturation. J. Biol. Chem. 2012, 287, 33781−33795. (192) Zorn, J.; Wille, H.; Wolan, D. W.; Wells, J. A. Self-Assembling Small Molecules Form Nanofibrils That Bind Procaspase-3 to Promote Activation. J. Am. Chem. Soc. 2011, 133, 19630−19633. (193) Boatright, K. M.; Renatus, M.; Scott, F. L.; Sperandio, S.; Shin, H.; Pedersen, I. M.; Ricci, J.-E.; Edris, W. A.; Sutherlin, D. P.; Green, D. R.; et al. A Unified Model for Apical Caspase Activation. Mol. Cell 2003, 11, 529−541.

(194) Kersse, K.; Verspurten, J.; Berghe, T.; Vanden; Vandenabeele, P. The Death-Fold Superfamily of Homotypic Interaction Motifs. Trends Biochem. Sci. 2011, 36, 541−552. (195) Deveraux, Q. L.; Leo, E.; Stennicke, H. R.; Welsh, K.; Salvesen, G. S.; Reed, J. C. Cleavage of Human Inhibitor of Apoptosis Protein XIAP Results in Fragments with Distinct Specificities for Caspases. EMBO J. 1999, 18, 5242−5251. (196) Shiozaki, E. N.; Chai, J.; Rigotti, D. J.; Riedl, S. J.; Li, P.; Srinivasula, S. M.; Alnemri, E. S.; Fairman, R.; Shi, Y. Mechanism of XIAP-Mediated Inhibition of Caspase-9. Mol. Cell 2003, 11, 519−527. (197) Yu, J. W.; Jeffrey, P. D.; Shi, Y. Mechanism of Procaspase-8 Activation by c-Flipl. Proc. Natl. Acad. Sci. U. S. A. 2009, 106, 8169− 8174. (198) Oberst, A.; Dillon, C. P.; Weinlich, R.; McCormick, L. L.; Fitzgerald, P.; Pop, C.; Hakem, R.; Salvesen, G. S.; Green, D. R. Catalytic Activity of the Caspase-8-FLIP(L) Complex Inhibits RIPK3Dependent Necrosis. Nature 2011, 471, 363−367. (199) Oberst, A.; Green, D. R. It Cuts Both Ways: Reconciling the Dual Roles of Caspase 8 in Cell Death and Survival. Nat. Rev. Mol. Cell Biol. 2011, 12, 757−763. (200) Pop, C.; Oberst, A.; Drag, M.; Van Raam, B. J.; Riedl, S. J.; Green, D. R.; Salvesen, G. S. FLIP-L Induces Caspase 8 Activity in the Absence of Interdomain Caspase 8 Cleavage and Alters Substrate Specificity. Biochem. J. 2011, 433, 447−457. (201) Safa, A. R. c-FLIP, A Master Anti-apoptotic Regulator. Exp. Oncol. 2012, 34, 176−184. (202) R Safa, A. Roles of c-FLIP in Apoptosis, Necroptosis, and Autophagy. J. Carcinog. Mutagen. 2013, S6, 1−9. (203) Flütsch, A.; Ackermann, R.; Schroeder, T.; Lukarska, M.; Hausammann, G. J.; Weinert, C.; Briand, C.; Grütter, M. G. Combined Inhibition of Caspase 3 and Caspase 7 by Two Highly Selective DARPins Slows down Cellular Demise. Biochem. J. 2014, 461, 279− 290. (204) Schroeder, T.; Barandun, J.; Flütsch, A.; Briand, C.; Mittl, P. R. E.; Grütter, M. G. Specific Inhibition of Caspase-3 by a Competitive DARPin: Molecular Mimicry between Native and Designed Inhibitors. Structure 2013, 21, 277−289. (205) Schweizer, A.; Roschitzki-Voser, H.; Amstutz, P.; Briand, C.; Gulotti-Georgieva, M.; Prenosil, E.; Binz, H. K.; Capitani, G.; Baici, A.; Plückthun, A.; et al. Inhibition of Caspase-2 by a Designed Ankyrin Repeat Protein: Specificity, Structure, and Inhibition Mechanism. Structure 2007, 15, 625−636. (206) Schweizer, A.; Briand, C.; Grutter, M. G. Crystal Structure of Caspase-2, Apical Initiator of the Intrinsic Apoptotic Pathway. J. Biol. Chem. 2003, 278, 42441−42447. (207) Martin, M. C.; Allan, L. a.; Mancini, E. J.; Clarke, P. R. The Docking Interaction of Caspase-9 with ERK2 Provides a Mechanism for the Selective Inhibitory Phosphorylation of Caspase-9 at Threonine 125. J. Biol. Chem. 2008, 283, 3854−3865. (208) Yaoita, Y. Inhibition of Nuclear Transport of Caspase-7 by Its Prodomain. Biochem. Biophys. Res. Commun. 2002, 291, 79−84. (209) Choi, Y. E.; Butterworth, M.; Malladi, S.; Duckett, C. S.; Cohen, G. M.; Bratton, S. B. The E3 Ubiquitin Ligase cIAP1 Binds and Ubiquitinates Caspase-3 and -7 via Unique Mechanisms at Distinct Steps in Their Processing. J. Biol. Chem. 2009, 284, 12772−12782. (210) Cowling, V.; Downward, J. Caspase-6 Is the Direct Activator of Caspase-8 in the Cytochrome c-Induced Apoptosis Pathway: Absolute Requirement for Removal of Caspase-6 Prodomain. Cell Death Differ. 2002, 9, 1046−1056. (211) Katsogiannou, M.; Andrieu, C.; Rocchi, P. Heat Shock Protein 27 Phosphorylation State Is Associated with Cancer Progression. Front. Genet. 2014, 5, 1−5. (212) Vidyasagar, A.; Wilson, N. A.; Djamali, A. Heat Shock Protein 27 (HSP27): Biomarker of Disease and Therapeutic Target. Fibrog. Tissue Repair 2012, 5, 1−7. (213) Wang, X.; Chen, M.; Zhou, J.; Zhang, X. HSP27, 70 and 90, Anti-Apoptotic Proteins, in Clinical Cancer Therapy. Int. J. Oncol. 2014, 45, 18−30. 6705


Chemical Reviews

Review

(214) Elliott, J. M.; Rouge, L.; Wiesmann, C.; Scheer, J. M. Crystal Structure of Procaspase-1 Zymogen Domain Reveals Insight into Inflammatory Caspase Autoactivation. J. Biol. Chem. 2009, 284, 6546− 6553. (215) Romanowski, M. J.; Scheer, J. M.; O’Brien, T.; McDowell, R. S. Crystal Structure of a Ligand-Free and Malonate-Bound Human Caspase-1: Implications for the Mechanism of Substrate Binding. Structure 2004, 12, 1361−1371. (216) Velazquez-Delgado, E. M.; Hardy, J. A. Phosphorylation Regulates Assembly of the Caspase-6 Substrate-Binding Groove. Structure 2012, 20, 742−751. (217) Cao, Q.; Wang, X.-J.; Liu, C.-W.; Liu, D.-F.; Li, L.-F.; Gao, Y.Q.; Su, X.-D. Inhibitory Mechanism of Caspase-6 Phosphorylation Revealed by Crystal Structures, Molecular Dynamics Simulations, and Biochemical Assays. J. Biol. Chem. 2012, 287, 15371−15379. (218) Stanger, K.; Steffek, M.; Zhou, L.; Pozniak, C. D.; Quan, C.; Franke, Y.; Tom, J.; Tam, C.; Elliott, J. M.; Lewcock, J. W.; et al. Allosteric Peptides Bind a Caspase Zymogen and Mediate Caspase Tetramerization. Nat. Chem. Biol. 2012, 8, 655−660. (219) Baumgartner, R.; Meder, G.; Briand, C.; Decock, A.; D’arcy, A.; Hassiepen, U.; Morse, R.; Renatus, M. The Crystal Structure of Caspase-6, a Selective Effector of Axonal Degeneration. Biochem. J. 2009, 423, 429−439. (220) Agniswamy, J.; Fang, B.; Weber, I. Plasticity of S2−S4 Specificity Pockets of Executioner caspase-7 Revealed by Structural and Kinetic Analysis. FEBS J. 2007, 274, 4752−4765. (221) Datta, D.; McClendon, C. L.; Jacobson, M. P.; Wells, J. A. Substrate and Inhibitor-Induced Dimerization and Cooperativity in Caspase-1 but Not Caspase-3. J. Biol. Chem. 2013, 288, 9971−9981. (222) Peng, C.; Cho, Y.; Zhu, F.; Zhang, J.; Wen, W.; Xu, Y.; Yao, K.; Ma, W.-Y.; Bode, A. M.; Dong, Z. Phosphorylation of Caspase-8 (Thr263) by Ribosomal S6 Kinase 2 (RSK2) Mediates Caspase-8 Ubiquitination and Stability. J. Biol. Chem. 2011, 286, 6946−6954. (223) Dix, M.; Simon, G.; Wang, C.; Okerberg, E.; Patricelli, M. P.; Cravatt, B. F. Functional Interplay between Caspase Cleavage and Phosphorylation Sculpts the Apoptotic Proteome. Cell 2012, 150, 426−440. (224) Li, X.; Wen, W.; Liu, K.; Zhu, F.; Malakhova, M.; Peng, C.; Li, T.; Kim, H. G.; Ma, W.; Cho, Y. Y.; et al. Phosphorylation of Caspase7 by p21-Activated Protein Kinase (PAK) 2 Inhibits Chemotherapeutic Drug-Induced Apoptosis of Breast Cancer Cell Lines. J. Biol. Chem. 2011, 286, 22291−22299. (225) Cardone, M. H.; Roy, N.; Stennicke, H. R.; Salvesen, G. S.; Franke, T. F.; Stanbridge, E.; Frisch, S.; Reed, J. C. Regulation of Cell Death Protease Caspase-9 by Phosphorylation. Science 1998, 282, 1318−1321. (226) Allan, L. A.; Morrice, N.; Brady, S.; Magee, G.; Pathak, S.; Clarke, P. R. Inhibition of Caspase-9 through Phosphorylation at Thr 125 by ERK MAPK. Nat. Cell Biol. 2003, 5, 647−654. (227) Nutt, L. K.; Margolis, S. S.; Jensen, M.; Herman, C. E.; Dunphy, W. G.; Rathmell, J. C.; Kornbluth, S. Metabolic Regulation of Oocyte Cell Death through the CaMKII-Mediated Phosphorylation of Caspase-2. Cell 2005, 123, 89−103. (228) Pastore, A.; Piemonte, F. S-Glutathionylation Signaling in Cell Biology: Progress and Prospects. Eur. J. Pharm. Sci. 2012, 46, 279− 292. (229) Klomsiri, C.; Karplus, P. A.; Poole, L. B. Cysteine-Based Redox Switches in Enzymes. Antioxid. Redox Signaling 2011, 14, 1065−1077. (230) Jiang, Z. L.; Fletcher, N. M.; Diamond, M. P.; Abu-, H. M.; Saed, G. M. S-Nitrosylation of Caspase-3 Is the Mechanism by Which Adhesion Fibroblasts Manifest Lower Apoptosis. Wound Repair Regen. 2009, 17, 224−229. (231) Huber, K. L.; Hardy, J. A. Mechanism of Zinc-Mediated Inhibition of Caspase-9. Protein Sci. 2012, 21, 1056−1065.

6706


Caspase Allostery and Conformational Selection - American Chemical

Recommend Documents