CHEMICAL BIOLOGY

Cell Death, Biological Mechanisms and Small Molecule Inhibitors of

 

Olga Korkina and Alexei Degterev, Department of Biochemistry, Tufts University, Boston, Massachusetts

doi: 10.1002/9780470048672.wecb059

 

The process of regulated or programmed cell death (PCD) executed through genetically encoded intrinsic cellular machinery is widely accepted to represent one of the key cellular responses to extrinsic and intrinsic stimulation. Extensive analysis of PCD carried out during the last decade clearly established that proper execution of PCD is important for normal mammalian development and also for homeostasis of the adult organism. Deregulation of PCD has been linked to development of many severe human diseases like cancer, autoimmunity, stroke, and some neurodegenerative diseases. Although apoptosis, which is the first discovered form of PCD, has been and remains the mainstay of PCD research, better understanding of the process of apoptosis led to the surprising discovery that other forms of PCD also exist and play multiple important roles in health and disease. Small-molecule inhibitors of apoptotic and nonapoptotic cell death have been successfully developed and proved very useful in defining the mechanisms and functional role of various PCD processes. Furthermore, some molecules have been developed extensively and represent emerging new therapies for human pathologies. In this article, we will discuss the major PCD-related protein targets of chemical inhibitors and describe the major classes of small molecule inhibitors developed to this point.

 

Introduction

The paradigm of programmed cell death (PCD) has emerged in the last decade as the critical mechanism of normal development and homeostasis of multicellular organisms. Apoptosis, which was the first discovered form of PCD (1), remains the central topic in the field of PCD research. Alterations to apoptotic signaling either through genetic or small-molecule means can cause significant developmental abnormalities, including mortality, and they can also lead to the development of serious pathologies in adult animals, such as cancer and autoimmunity.

Apoptosis is associated with several highly uniform and characteristic morphological changes, such as cell shrinkage, cell membrane blebbing, condensation of nuclear chromatin, micronuclei formation, extensive vacuolization of cytoplasm, and disintegration of the cell into small fragments (apoptotic bodies) (2), which are reflective of the highly conserved nature of the apoptotic execution machinery. In general, two different general pathways of apoptotic cell death initiation exist: extrinsic and intrinsic (Fig. 1). The extrinsic pathway is activated by external stimuli, such as engagement of death domain receptors (DRs) with their cognate ligands. The intrinsic pathway is stimulated by intracellular stress, such as DNA damage. The pathway of DR-induced apoptosis has been studied and characterized extensively. It involves recruitment of the proforms of cysteine proteases, caspase-8 and caspase-10, into the receptor-induced death inducing signaling complex (DISC), which results in their autocatalytic cleavage and activation. Apical caspases can trigger execution of apoptosis directly by processing and activation of the “effector” caspases (caspase-3, caspase-6, and caspase-7) (3). Effector caspases execute apoptosis by cleaving various cellular substrates, which leads to the orderly cell demise. In some types of cells, DR signaling relies on a mitochondrial amplification step, which is carried out through caspase-8 and caspase-10 mediated cleavage of the BH3 domain-only Bcl-2 family member Bid (4-6). Processed Bid translocates to the mitochondria, where it causes release of cytochrome c from the intermembrane space through induction of oligomerization of the proapoptotic Bcl-2 family members, Bax and Bak (7). In the cytosol, cytochrome c interacts with the Apaf-1 adaptor molecule and induces an apoptosome complex formation (8). The apoptosome is a heptamer composed of seven Apaf-1 adaptor molecules; each molecule is bound to one molecule of cytochrome c and a monomer of the initiator caspase-9 (9). In this scenario, caspase-9, which is activated through apoptosome-induced dimerization and conformational change, subsequently processes the executioner caspases (10). The intrinsic pathway of apoptotic death also proceeds through release of cytochrome c from mitochondria, and different forms of stress use different BH3-only factors for signaling (11).

Although apoptosis remains the mainstay of PCD research, rapidly accumulating evidence indicates that programmed or intrinsically regulated cell death can occur through additional pathways independent of caspase activation and other proapoptotic factors. Multiple alternative processes of regulated cell death have been described, including autophagic cell death, mitotic catastrophe, necroptosis, oncosis, and so on. In most cases, mechanisms of these processes are much less understood than that of apoptosis. Here, we will discuss some emerging small-molecule regulators of these pathways, which have proven helpful in understanding the processes of nonapoptotic cell death. We have omitted several important protein factors from our discussion, such as PI3-kinase, Akt, and p53, and the process of autophagy. Although corresponding cellular pathways make important contributions to the regulation of cell death, they play much broader role in cellular regulation and deserve separate discussion.

 

 

Figure 1. Schematic representation of the receptor-mediated (extrinsic) and the intracellular stress-mediated (intrinsic) pathways of caspase activation. Death-receptor signaling may involve direct caspase-8-mediated caspase-3 activation (type 1 cells) or a Bid-cleavage-dependent mitochondrial amplification step (type 2 cells).

 

Caspases

The family of cysteine proteases, which are called caspases, plays a major role in the execution of apoptotic cell death (12). Many studies suggest that increased apoptosis and caspase activity contribute to tissue damage in both acute (e.g., myocardial infarction, stroke, sepsis, spinal cord injury) and chronic (e.g., Alzheimer’s, Parkinson’s, Huntington’s disease) human diseases (13, 14). Caspase family members are also prominently involved in inflammatory responses and are required for processing and secretion of proinflammatory cytokines (15). For example, deficiency in caspase-1 or caspase-11 leads to significant protection from septic shock (16, 17). Thus, inhibition of caspase activity has emerged as a promising direction for cytoprotective therapies. All caspases, with the exception of caspase-9, are expressed in the form of catalytically inactive single-chain zymogens that contain a large and a small subunit (18). Activation of the proform occurs by proteolytic cleavage that releases N-terminal pro-domain and separates large and small subunits. An active enzyme is a heterotetramer composed of two large and two small subunits with two identical active sites, which are formed with the contribution from both large and small subunits (19). A small molecule activator of pro-caspase-3, PAC-1 (Fig. 2c), has been identified recently in an in vitro screen of 20,500 compounds (20). This molecule displayed an EC50 = 220 nM in an in vitro assay and EC50 < 1 μM in cancer cell lines. Furthermore, this molecule was significantly more toxic to primary colon cancer cells compared with matched normal cells (at least 10-fold difference), which suggests that activation of procaspase-3 may represent a new approach for selective killing of cancer cells. Indeed, administration of PAC-1 was found to attenuate growth of three different types of cancer in vivo. However, the precise mechanism of caspase activation by PAC-1 has not yet been described.

Unlike many other proteases, caspases possess a high degree of substrate specificity, that is, they display an almost absolute requirement for an aspartic acid residue in the P1 position of a substrate. This requirement stems from several hydrogen bonds that Asp forms within the caspase substrate-binding pocket (21). Furthermore, three preceding amino acid residues (P2-P4) of the substrate contribute to substrate recognition by specific caspase family members, although substrate selectivity of caspases is typically not absolute (22). Based on their substrate preferences, caspases can be divided into three groups (23). Group 1, including caspase-1, -4, and -5, prefers a hydrophobic amino acid in the P4 position. Group 2 displays strong preferences for Asp in the P4 position and includes caspase-2, -3, and -7. Activity of group 3 caspases, which consists of caspase-6, -8, -9, and -10, is less dependent on the identity of a P4 residue.

 

 

Figure 2. Small-molecule inhibitors of caspases. (a) Schematic of a typical caspase inhibitor; (b) caspases substrate specificities: preferred amino acids in P1-P4 positions; (c) chemical structure of PAC-1, procaspase-3 activator (EC50 of 0.22 uM); (d) low-nanomolar inhibitors of caspases-3 and -7 (compound 1) and caspases-3, -7 and -9 (compound 2); (e) caspase-1 peptidomimetic inhibitor VX-740 or pralnacasan from Vertex Pharmaceutical (Cambridge, MA); (f) peptidomimetic irreversible oxamyl dipeptide pan-caspase inhibitor IDN-6556 from IDUN Pharmaceuticals; and (g) caspase-3 inhibitor.

 

Most caspase-inhibitor design strategies target their active sites and are based on caspase substrate preferences. Tetrapeptide inhibitors, which are based on identified sequences of the four amino acid recognition motifs, were shown to inhibit caspase family members selectively (24) (Fig. 2a). Overall, a typical peptide-based caspase inhibitor consists of three major structural components:

1. the “warhead” moiety that interacts with an active site Cys of the caspase;

2. Asp in P1 position (invariable for all peptide-based inhibitors); and

3. P2-P4 sequence, which provides some selectivity toward individual caspase sub-classes (21) (Fig. 2b). Selectivity of peptide substrates is mainly defined by P4 and to a lesser extent by P2 and P3 residues (Fig. 2b) (12, 22).

The warhead is an electrophilic group that reacts with the nucleophilic Cys of the active site to form reversible or irreversible adducts. Use of aldehyde, semicarbazone, or thiomethylketone groups leads to reversible caspase inhibitors, whereas fluoromethylketones, (2, 3, 5, 6)-tetrafluorophenoxymethylketones, chloromethylketones and acyloxymethylketones are used to generate irreversible inhibitors (21). Although peptide inhibitors can be very useful in defining the functional role of apoptosis and caspases in particular cell death paradigm in vitro and even, in some cases, in vivo (25), peptide inhibitors possess several general disadvantages, including toxicity of the leaving group, limited half-life in cells and in vivo, modest selectivity toward individual caspases, and lack of oral bioavailability. These disadvantages limit their use mostly to cell-based studies. Specificity of these molecules toward caspases is somewhat limited, as for example, an irreversible pan-caspase inhibitor z-VAD-fmk was found to inhibit other unrelated cysteine proteases, such as cathepsins B and H (26).

Several promising nonpeptide inhibitors of caspases have also been developed, which may overcome many limitations of tetrapeptide molecules. Lee et al. (27, 28) used high-throughput screening to identify 5-nitroisatin as a caspase inhibitor, which was optimized significantly to obtain selective, low-nanomolar inhibitors of caspases-3 and -7 (Compound 1, Fig. 2d) and caspases-3, -7 and -9 (Compound 2, Fig. 2d). In contrast to peptide inhibitors, these compounds displayed high activity despite the lack of interaction with S1 subsite of caspase substrate pocket, and X-ray crystallography suggested that they primarily interacted with the S2 subsite. These compounds inhibited apoptosis in several different cell types, which include camptothecin-treated Jurkat T cells and chondrocytes and cycloheximide-treated neutrophiles (27, 28).

Okamoto et al. (29) pursued development of peptidomimetic caspase-1 inhibitors based on the crystal structure of caspase-1/ Ac-Tyr-Val-Ala-Asp-H complex, which resulted in an inhibitor displaying potent activity in an in vitro caspase assay (IC50 = 38 nM) and blocking IL-1β processing in the cells with EC50 = 230nM. Furthermore, this molecule blocked IL-1β release in mice in a dose-dependent fashion. X-ray analysis revealed that naphthoyl and methyl groups of the methanesulfonamidecar- bonyl made critical contacts with S4 and S1 subsites of the caspase-1 active center, which is consistent with their important role in substrate recognition by caspases.

Vertex Pharmaceuticals (Cambridge, MA) reported development of several additional caspase-1 peptidomimetic inhibitors, such as VX-740 (Pralnacasan, Fig. 2e), for the treatment of rheumatoid arthritis (30-32). VX-740 is a pro-drug, which is converted in vivo into an aldehyde active form with Ki of 1 nM against caspase-1. VX-740 was found to reduce inflammation and disease symptoms significantly in patients during Phase 2 clinical trials (33, 34). However, clinical trials also demonstrated significant liver toxicity associated with long-term dosing of this molecule, which is likely associated with reactivity of the warhead group. This finding led to premature termination of clinical trials. Synthesis of improved and highly selective caspase-1 inhibitor, VX-765, which is a prodrug resulting in 4-hydroxybutyrolactone “warhead” moiety, has also been reported recently (35, 36). This molecule efficiently blocked release of inflammatory cytokines in the cells and in vivo; however, no data from clinical trials has been reported thus far.

IDUN Pharmaceuticals (San Diego, CA) also reported development of a potent peptidomimetic irreversible oxamyl dipeptide inhibitor IDN-6556 that uses 2,3,5,6-tetrafluro-phenoxymethylketone warhead (Fig. 2f) (37, 38). This molecule preferentially accumulates in the liver, which results in pronounced liver protection in animal models [for example, after Fas-induced liver injury (39)], and showed significant promise in clinical trials of acute alcoholic hepatitis, human liver preservation injury, and chronic hepatitis C (38, 40, 41).

To overcome limitations of peptidomimetic inhibitors, such as limited central bioavailability caused by accumulation in the liver, a novel strategy for rapid identification of non-peptidomimetic caspase inhibitors, tethering has been recently proposed (42, 43). Tethering is based on covalent capture of sulfohydril-containing small molecules that interact within the active site of caspase-3. For this assay, caspase is modified to contain a free thiol-bearing “extender” attached covalently to the active site cysteine, which is used for small molecule capture. Using this approach, a caspase-3/extender complex was screened against a library of fragments modified to contain a free-sulfhydryl group. Selected fragments were subsequently combined with a reversibly binding form of extender, which resulted in rapid selection of potent and reversible caspase-3-specific inhibitor with Ki of 2.8 μM. Additional chemical modifications enhanced its potency into low nM range (Fig. 2g) (42).

 

BCL-2 Family

Members of the Bcl-2 protein family are key regulators of the mitochondrial step in apoptotic pathway (44). Upregulation of antiapoptotic Bcl-2 family members is commonly observed in many types of cancers and is well established to play a major role in apoptosis evasion of cancer cells under chemotherapeutic treatment conditions (45).The Bcl-2 family can be further subdivided into three classes of proteins. The proapoptotic members Bax and Bak activate apoptosis through formation of a pore in the outer mitochondrial membrane, which results in cytochrome c release and activation of the apoptosome (46). The primary function of antiapoptotic proteins Bcl-2, Bcl-xL, Bcl-w, Mcl-1, and Bfl-1 is to inhibit the functions of Bak and Bax by preventing their oligomerization (47). The members of the third “BH3-only” group have homology with the other family members only in the BH3 domain and serve as upstream sensors of apoptotic signaling. Once activated by an apoptotic signal, BH3-only proteins are proposed to act through two different mechanisms. Some BH3-only factors, which are termed “sensitizers” (Bad, Bik), may act primarily by inhibiting antiapoptotic Bcl-2 family members through BH3-mediated binding to the hydrophobic cleft formed by BH1, BH2, and BH3 domains of antiapoptotic factors. Another subgroup of BH3-only factors, termed “activators,” including Bid and Bim, were proposed to activate Bax and Bak directly, which induces their oligomerization (48).

Initial studies of BH3-dependent heterodimerization found that an isolated 16 a.a. BH3 peptides derived from several Bcl-2 family proteins can bind antiapoptotic Bcl-2 family members with submicromolar affinity (49) and antagonize Bcl-xL heterodimerization with proapoptotic proteins Bax and Bad (50, 51). The ability of synthetic BH3 peptides to trigger apoptosis was first demonstrated in a cell-free system based on extracts of Xenopus eggs (52). In these studies, BH3 peptides derived from Bak, Bax, and Bid were all found to induce apoptosis through rapid activation of caspases. Because BH3 peptides cannot permeate the cells readily, several strategies were used to generate cell-permeable BH3 peptide-based proapoptotic agents. Wang et al. (53) demonstrated that attachment of decanoic acid allows generation of cell-permeable BH3 peptides. One such peptide, termed CPM-1285 and containing the BH3 domain of mouse Bad, was shown to compete with a fluorescein-labeled Bak BH3 peptide for binding to Bcl-2 (IC50 of 130 nM) in vitro and to trigger apoptosis in human myeloid leukemia HL-60 cells. In another approach, Bak BH3 peptide was fused to Antennapedia cell-penetrating peptide, which resulted in activation of apoptosis in the cells in the presence of 50-μM peptide (54). Similarly, fusion of Bim BH3 peptide with cell-penetrating TAT peptide led to a molecule that can induce apoptosis in different types of cancer cells (55). Finally, in a highly innovative approach, Walensky et al. (56) described introducing internal “crosslinks” into BH3 peptides, which led to stabilization of a helixes, increased cell permeability, and bioavailability. Such “stapled” Bid BH3 peptide efficiently induced apoptosis in leukemia cells in vitro and in vivo, and this activity was increased even more by membrane targeting of the peptide (57).

The first nonpeptidic inhibitor of Bcl-2 family proteins was identified by Wang et al. (58) in 2000 using a virtual-screening strategy. This method relies on the high-resolution three-dimensional structure of a targeted receptor protein and computer-aided techniques to screen a large number of organic compounds for a potential ligand. Virtual screening of more than 190,000 organic molecules resulted in identification HA14-1 (Fig. 3b) and subsequent in vitro binding assay demonstrated the interaction of HA14-1 with the surface pocket of Bcl-2 with an IC50 value of 9 μM. Subsequently, multiple research groups reported activation of apoptosis by HA14-1 in a variety of cell types through mechanisms related to regulation of the Bcl-2 family as well as retardation of glioblastoma tumor growth in vivo when this molecule was combined with etoposide (59). The group of Dr. Shaomeng Wang (University of Michigan) also successfully used computational strategies to identify a number of different submicromolar small molecule antagonists of antiapop- totic Bcl-2 family members [TW-37, Fig. 3c, Ki = 290 nM (60); pyrogallol-based inhibitors, Fig. 3d, Ki = 110nM (Bcl-2) (61); flavanoid compound BI-33, Fig. 3e, Ki = 17nM (Bcl-2)(62)]. These molecules were all found to trigger apoptosis in tissue culture, with some molecules displaying very potent effect (IC50 for BI-33 in MDA-MB-231 breast cancer cells = 110 nM). Furthermore, one of these molecules, TW-37, was found to enhance the antitumor effect of standard chemotherapy (cyclophosphamide-doxorubicin-vincristine-prednisone, CHOP) in mouse lymphoma model (63). Overall, these data suggest that computational approaches can be very powerful in designing proapoptotic inhibitors of the Bcl-2 family.

Using a competitive binding assay based on fluorescence polarization (FP), Degterev et al. (64) screened a chemical library of 16,320 compounds to identify two classes of small molecule ligands of Bcl-xL, which are termed BH3I-1 and BH3I-2 (Fig. 3a). These compounds were shown to inhibit BH3 peptide binding to Bcl-xL and Bcl-2 with Ki values in the low micromolar range (Ki of 2.4-15.6 μM) as determined by NMR titration assays (64, 65). The NMR titration experiments suggested that BH3I molecules directly interact with the BH3 binding pocket in disrupting Bcl-xL heterodimerization. These compounds were found to induce apoptosis in Jurkat cells through disruption of Bcl-2/Bcl-xL heterodimerization measured in intact cells (64). In addition to affecting Bcl-2-dependent regulation of outer mitochondrial membrane permeability, BH3I-2 was also found to induce damage to inner mitochondrial membrane, likely also through interaction with Bcl-2 (63). In another report, small scale FP-based screening of polyphenols identified gossypol (Fig. 3f) as a novel Bcl-2 and Bcl-xL inhibitor (66). This molecule was found to possess significant activity as a sensitizer when combined with CHOP in a mouse lymphoma model (67) and ionizing radiation in prostate cancer xenograft (68). Using a similar FP screen, PKC inhibitor chelerythrine (Fig. 3g) was identified as a low micromolar Bcl-xL/BH3 inhibitor (69). Curiously, NMR analysis and molecular docking suggested that unlike BH3I-1, this and related sanguinarine molecules do not bind into the hydrophobic cleft of Bcl-xL, but rather to the BH groove and BH1 domain, respectively. This analysis suggests a distinct mechanism of BH3 domain displacement, which was proposed to explain increased cytotoxicity of these molecules compared with BH3I-1 (70).

Researchers at Abbott Laboratories used a different approach for discovering high-affinity protein ligands, the “structure-activity relationships by nuclear magnetic resonance” (“SAR by NMR”) (71). In this method, the relatively large site is divided into two smaller half-sites that are targeted individually by small molecules. The two lead molecules are then chemically linked to improve affinity. In this approach, although the two molecules that target each half displayed Ki values of only 0.3 and 4.3 mM, the combined molecule displayed a Ki of 36 nM against Bcl-2. Subsequent chemical modifications to improve affinity and decrease nonspecific binding to human serum albumin yielded ABT-737 molecule with a Ki < 1 nM for Bcl-xL, Bcl-2, and Bcl-w (Fig. 3i) (72). ABT-737 was found to kill the cells efficiently through Bcl-2 or Bcl-xL-dependent mechanism. Furthermore, ABT-737 induced cytochrome c release from isolated mitochondria, which was dependent on inhibition of Bax and Bak by Bcl-2 (72). In other words, ABT-737 was found to antagonize prosurvival activity of Bcl-2 aimed at inhibition of Bax and Bak. Furthermore, Oltersdorf et al. (72) demonstrated that ABT-737 can act as a selective cancer therapeutic drug that displays potent single-agent efficacy against small cell lung cancer (SCLC) cells and cells from lymphoid malignancies, which are known to express high levels of Bcl-2, with EC50 as low as 10nM. In mouse xenograft models, ABT-737 treatment provided significantly improved survival in mice injected with either lymphoma or SCLC cell lines (73). Curiously, very high specificity of ABT-737 binding to Bcl-2 may actually limit efficacy of this molecule as it has been found not to inhibit activity of the antiapoptotic Bcl-2 family member, Mcl-1, which results in reduced activity in multiple cancer cell lines (74, 75).

Another small-molecule pan-Bcl-2 inhibitor that mimicks BH3-only proteins, GX15-070 (developed by GeminX, Montreal, Canada) (Fig. 3k), was shown to be a potent apoptosis inducer in breast cancer, chronic lymphocytic leukemia, multiple myeloma, and mantle cell lymphoma cell lines (76-79). GeminX is currently conducting several clinical trials of GX15- 070 in multiple cancer types as a single agent and in combination with other agents.

The laboratory of Dr. David Hockenberry discovered that increased sensitivity of Bcl-2 and Bcl-xL-expressing cells to mitochondrial respiratory chain inhibitor, antimycin A, is caused by its direct interaction with Bcl-2 family members (80). Furthermore, this affect is retained by 2-methoxy antimycin A (2MAA) (Fig. 3j), an analog lacking ability to inhibit complex 3 of respiratory chain. Curiously, activity of 2MAA seems very different from that of all of the abovementioned Bcl-2/Bcl-xL inhibitors, as 2MAA displays preferential toxicity toward Bcl-2/Bcl-xL-overexpressing cells. This activity may be caused by the unique ability of 2MAA to antagonize Bcl-xL dependent changes in cell metabolism, namely reduction in oxidative phosphorylation and activation of glycolysis (80). Such “gain-of-function” Bcl-2 antagonists may be very beneficial for preferentially inducing cell death in Bcl-2 overexpressing cancers.

 

 

Figure 3. Inhibitors of Bcl-2 and Bcl-XL. (a) BH3I-1 and BH3I-2, (b) HA14-1, (c) TW-37, (d) pyrogallol-based inhibitor, (e) flavonoid BI-33, (f) R-(-)-gossypol, (g) chelerythrine, (h) sanguinarine, (i) ABT-737, (j) GX15-070, (k) 2-methoxy antimycin A (2MAA).

 

The lap, Xiap, and Smac Mimetic Peptides

The inhibitor of apoptosis proteins (IAPs), which are characterized by the presence of one or more baculovirus IAP repeat (BIR) domains, are a family of endogenous apoptosis inhibitors that possess multiple antiapoptotic activities, including binding and inhibition of active caspases 3, 7, and 9. By inhibiting the downstream caspases 3 and 7, IAPs block the convergence point of multiple caspase activation pathways and thus inhibit apoptosis induced by various stimuli (81). At least eight human IAP members have been identified, of which XIAP (X-linked IAP) and survivin have received the most attention as therapeutic targets (82).

Survivin is a bifunctional protein that acts as a suppressor of apoptosis and plays a central role in the regulation of cell division. Survivin is preferentially expressed in malignant cells, and its expression is frequently responsible for radioresistance of malignancies (83, 84). However, this effect may not be linked to the direct regulation of apoptosis, but rather to the regulation of cell division. Cell-cycle-dependent transcriptional regulation of the survivin gene (85) as well as posttranslational modifications, including phosphorylation by the p34cdc2 (86), were found to be essential for the cell-cycle control. Based on the finding that pharmacologic inhibition of mitotic phosphorylation of survivin accelerated the protein destruction and counteracted its function (87), CDK inhibitors such as flavopiridol (Fig. 4a) and purvalanol A (Fig. 4b), which is a more specific p34cdc2 inhibitor, were tested in tumor cells arrested at mitosis with taxol, which induces hyperphosphorylation of survivin (88). Administration of CDK inhibitors resulted in escape from the mitotic block imposed by taxol, activation of mitochondria-dependent apoptosis, and anticancer activity in vivo (87). The stability and function of survivin depends on physical interaction between its BIR domain and ATPase domain of the molecular chaperone heat shock protein 90 (HSP90). Targeted antibody-mediated disruption of the survivin-Hsp90 complex in cancer cells resulted in proteosomal degradation of survivin, mitotic arrest, and mitochondria-dependent apoptosis (89). A structure-based rational screening for antagonists of the survivin-HSP90 complex identified a cell-permeable peptidomimetic derived from the Lys79-Leu87 sequence of the survivin called shepherdin (90). Shepherdin inhibited HSP90 chaperone function by competing with ATP binding and destabilized several HSP90 client proteins, including Akt, CDK6, and telomerase, to induce cell death via apoptotic and nonapoptotic mechanisms in various tumor cell lines. Shepherdin (79-83), which is a cell-permeable five-residue peptide that contains the Lys79-Gly83 sequence of shepherdin essential for HSP90 binding (91), induced rapid killing of different types of human acute myeloid leukemia (AML) cell lines, but not of normal mononuclear cells. Moreover, shepherdin (79-83) efficiently inhibited the growth of AML xenograft tumors without systemic or organ toxicity (91). More recently, a combined structure- and dynamics-based computational design strategy using shepherdin as a scaffold identified the nonpeptidic small molecule that targeted the HSP90 function, 5-aminoimidazole-4-carboxamide-1-P-D-ribofuranoside (AICAR, also a known activator of AMP kinase (AMPK)) (Fig. 4c) (92). AICAR was shown to destabilize several HSP90 client proteins in vivo, including survivin, and to exhibit antiproliferative and proapoptotic activity in multiple tumor cell lines, but not in normal human fibroblasts. Finally, a small molecule that selectively inhibits survivin gene transcription and protein expression has been identified, YM155 (Fig. 4d), and is currently being evaluated in a Phase 2 study for patients with stage 3 and stage 4 melanoma. It showed marked antiproliferative activity in the nanomolar range in a broad spectrum of human tumor cell lines and induced tumor regression in lymphoma, prostate cancer, and non-small cell lung cancer xenografts (93).

XIAP is the best characterized human IAP and is the only member of this family shown to inhibit both the initiator (caspase-9) and executioner (caspases-3 and -7) caspases directly. Structural and functional studies of XIAP have demonstrated that a groove in its BIR3 domain is required for binding and inhibition of caspase-9, whereas two surfaces of the BIR2 domain and the juxtaposed linker region bind and inhibit active caspases-3 and -7 (94, 95). The natural inhibitor of XIAP, cIAPl, and cIAP2, the proapoptotic protein SMAC/DIABLO, is released into the cytosol from the mitochondrial intermembrane space during apoptosis activation (96, 97). SMAC dimers cooperatively bind and inhibit both BIR3 and BIR2 domains of IAPs and thus relieve their caspase inhibitory function (98). Peptides that correspond to the four N-terminal amino acids of SMAC (AVPI) were shown to be sufficient for binding to XIAP and preventing XIAP-mediated inhibition of caspase-9 (99, 100). On the other hand, the SMAC peptides cannot relieve the inhibition of caspase-3 by XIAP, because they do not change the XIAP conformation around the linker region (101). When delivered into the cells either by conjugation with the TAT protein-transduction domain (102) or to polyarginine tail (103), SMAC peptides sensitized the SHEP neuroblastoma (102) and non-small cell lung carcinoma H460 (103) cell lines to apoptotic cell death induced by chemotherapeutic drug treatment. In addition, cell-permeable SMAC peptide delayed tumor growth of lung cancer and glioma xenografts (102, 103). These studies provided the proof-of-concept that small-molecule SMAC-mimics can be effective as the anticancer agents (82).

 

 

Figure 4. Structures of small molecule inhibitors of (a, b) CDK, (c) HSP90, (d) survivin gene expression, (e-l) XIAP, and (m) omi/HtrA2. (a) flavopiridol, (b) purvalanol, (c) 5-aminoimidazole-4-carboxamide-1-beta-D-ribofuranoside (AICAR), (d) YM155, (e-h) SMAC-mimetics, (i) Smac mimetic tetrazoyl thioether, (j) benzoquinone embelin, (k) aryl sulfonamide, (l) XIAP inhibitor of Polyphenylurea series, (m) Omi/HtrA2 inhibitor Ucf-101.

 

Nonpeptidic small molecule inhibitors of BIR3 domain of XIAP with micromolar binding affinities were synthesized at Abbott Laboratories (Abbott Park, IL)using structure-based design (104). Having conserved the first amino acid residue of SMAC peptide alanine, the substituted five-membered heterocycles such as thiazoles and imidazoles were identified to serve as a replacement for peptide fragments of the lead (Fig. 4e). Several research groups have also reported the discovery of nonpeptide XIAP inhibitors active in the cells. Using a high-throughput fluorescent polarization assay, pentapeptides competing with the binding of SMAC-like protein HID to BIR3 domain of XIAP with affinities in 40-60-nM range were identified (105). Tripeptide peptidomimetics (Fig. 4f) based on these leads were shown to inhibit the interactions between the BIR3 domain of XIAP and SMAC, caspase-9, and SMAC-derived peptide (105). When the cytotoxicity of selected peptidomimetics was assessed in various cancer cell lines (105), compounds revealed a wide range of potencies from low nanomolar activity in some cell lines to no activity at 50 μM in most others. The toxicity exhibited by peptidomimetic BIR3 ligands in the sensitive cell lines (breast cancer cell lines BT-549 and MDA-MB-231, melanoma cell line SK-MEL-5, and human myeloid leukemia HL-60 cell line) was observed in the absence of additional apoptotic stimulation (105). Furthermore, the selected peptidomimetics were found to slow the growth of tumors in a MDA-MB-231 breast cancer xenograft model (105). Another research group reported synthesis of a SMAC-mimetic, which was approximately 23 times more potent than SMAC peptide in binding the BIR3 domain of XIAP (Fig. 4g) (106). This compound efficiently inhibited the growth of etoposide-treated Jurkat leukemia T cells stably transfected with XIAP vector, which protects the cells from etoposide-induced apoptosis.

The broad-spectrum peptidomimetic IAP family inhibitors were also recently developed (107). Designed based on (7, 5)-bicyclic scaffold, these SMAC-mimetics were found to antagonize the protein interactions that involve XIAP, melanoma IAP (ML-IAP), cIAPl, andcIAP2. The most potent SMAC-mimeticvation of TRAIL receptor-2 (DR5/TRAIL-R2) via an anti-DR5 was more specific for cIAP1-BIR3 and ML-IAP-BIR with Ki ~50 nM (Fig. 4h). The compounds were demonstrated to activate caspase-3 and -7, to reduce cell viability in assays using MDA-MB-231 breast cancer cells and A2058 melanoma cells and to enhance doxorubicin-induced apoptosis in MDA-MB-231 cells.

Computer-based rational drug design was successfully used to synthesize a tetrazoyl thioether (Fig. 4j), a dimeric SMAC-mimic that binds both the BIR3 and BIR2 domains of XIAP with nanomolar affinity (108). Furthermore, this molecule also cross-reacted with cIAP1 and cIAP2. The potency of this dimeric inhibitor is consistent with the recently reported synergistic BIR2/BIR3 inhibition by dimeric SMAC peptide, which is explained by close proximity (<45 angstrom) between BIR2 and BIR3 binding sites (109). The tetrazoyl thioether sensitized cells to the death induced by death receptor ligands [tumor necrosis factor alpha (TNFα) and TNF-related apoptosis-inducing ligand (TRAIL)] and promoted the activation of caspase-8. This result was unexpected because XIAP is not known to inhibit caspase-8 directly. Rather, recent analyses suggested that killing by this molecule is primarily mediated by targeting cIAP1 and cIAP2, which promotes their autoubiquitination and degradation (110). This, in turn, leads to stabilization of NIK kinase, NF-kB activation, and TNFa production, triggering apoptosis in an autocrine mode. Furthermore, cIAP degradation promotes TRAF2-dependent RIP1 recruitment to TNFR1, which promotes formation of RIP1-dependent caspase-8-activating complex. These unexpected findings suggest that cIAPs may be more important targets for SMAC mimetic compounds than XIAP, for which this class of molecules was originally developed.

Virtual screening was also used to identify the BIR3 domain inhibitors (111). In this case, a library of Chinese herbal remedies was docked into the BIR3 domain in silico, which resulted in selection of natural compound benzoquinone embelin (Fig. 4i). Subsequent studies confirmed that embelin binds to the XIAP BIR3 domain, which resulted in the inhibition of its interaction with caspase-9, and induced apoptosis in the prostate cancer cells expressing high levels of XIAP. In stably XIAP-transfected Jurkat cells, embelin was shown to overcome the protective effect of XIAP effectively, which enhanced etoposide-induced apoptosis. At the same time, this molecule had a minimal effect in Jurkat cells transfected with vector control.

The BIR2 domain of XIAP has also been specifically targeted for inhibition. The linker region immediately to the N-terminus of the BIR 2 domain binds the catalytic domain of caspase-3 and blocks the active site of the enzyme through steric hindrance (94, 112, 113). This interaction is relatively weak and is stabilized by a stronger interaction between the binding groove of BIR 2 and a site on a small subunit of caspase 3 (94, 112). Using a high-throughput enzymatic de-repression assay based on caspase-3 proteolytic activity, Wu et al. (114) identified a series of aryl sulphonamide inhibitors of XIAP (Fig. 4k). These molecules bind the BIR2-linker region and were found to sensitize resistant cells lines to death triggered by the acti-specific antibody (114). Using a similar enzymatic assay, another research group screened a combinatorial library of approximately 1 million diverse small molecules and identified several active compounds including the polyphenylurea series (Fig. 4l) (115). These molecules also derepressed XIAP- and BIR 2-mediated inhibition of caspases-3 and -7 in vitro but not the BIR3-dependent inhibition of caspase-9 (115, 116). Consistent with this result, inhibitors specifically interacted with BIR2 but not BIR3 domain of XIAP in cell-free binding studies. The active compounds, but not inactive controls, were directly toxic to several hematologic and solid tumor cell lines. In other tumor cell lines, these agents were mostly nontoxic as single agents but still sensitized them to death induced by death receptor ligands (115).

 

HtrA2/Omi (High Temperature Recruitment A2)

Similar to SMAC, serine protease HtrA2/Omi is localized to the mitochondrial intermembrane space and is released into cytosol in response to apoptotic stimuli. HtrA2 can induce cell death in a caspase-dependent manner by interacting with IAPs, in a manner similar to SMAC, and in a caspase-independent manner through its intrinsic serine protease activity (117, 118). A reversible, cell-permeable small molecule inhibitor of HtrA2 protease, UCF-101, has been identified through high-throughput screening of a combinatorial library using recombinant Omi protease (residues 134-458) and fluorescein-labeled casein as a generic substrate (specific substrates of HtrA2 have not been known until recently) (119, 120) (Fig. 4m). Although this molecule displays only micromolar activity in vitro (IC5o = 9.5 μM), it has a significant selectivity for HtrA2 over other serine proteases (IC50 ≥ 200 μM). Quite impressively, UCF-101 was shown to ameliorate heart dysfunction following ischemia/reperfusion injury in in situ and in vivo in rat models (121). Furthermore, in this study UCF-101 was found to block mitochondria-to-cytosol translocation of HtrA2/Omi and degradation of XIAP (consistent with the previously reported role of the serine protease activity of HtrA2/Omi in XIAP degradation), which resulted in suppression of caspase-3, -7, and -9 processing. In addition, treatment with UCF-101 also led to the suppression of FLIP degradation and to the reduced Fas receptor expression. These data establish a specific role of HtrA2/Omi in ischemia/reperfusion injury through regulation of XIAP degradation and Fas-induced apoptosis. It should be mentioned, however, that another research group recently suggested that the cytoprotective effect of UCF-101 in the cell-culture experiments could be partially attributed to the activation of stress response pathways, rather than direct HtrA2/Omi inhibition. This finding suggested the possibility that HtrA2/Omiindependent effect of UCF-101 might also contribute to the in vivo cytoprotection (122).

 

Poly(Adp-Ribose) Polymerase (Parp)

The polyADP-ribosylation reactions play important roles in many cellular processes, which include regulation of DNA repair, transcriptional control, cellular transformation, and cell death (123). PARP is an enzyme-sensing single and double strand DNA nicks, which catalyzes addition of ADP-ribose units to DNA, histones, and various DNA repair enzymes (using NAD+ as a substrate) to promote DNA repair. Mouse knockout studies showed that combined deletion of just 2 of the 17 PARP family members (PARP-1 and PARP-2) is sufficient to block DNA repair (124, 125). Curiously, multiple studies suggested that combining inhibition of DNA repair with the use of DNA-damaging agents hypersensitizes cancer cells to cell death, which prompted development of PARP inhibitors as the general sensitizing anticancer agents (126). In addition, tumors deficient in DNA repair-associated factors BRCA1, BRCA2, and ATM, were all shown to be hypersensitive to PARP inhibitors, which suggests that PARP inhibitors can be useful in killing these types of cancer cells (127, 128).

At the same time, overactivation of PARP, which is frequently observed in various pathologies, including cardiovascular, neurological, and inflammatory diseases, was shown to result in the depletion of NAD+, leading to the loss of ATP and necrotic cell death. Cell-based studies showed that overactivated PARP-1 mediates both mitochondria-dependent apoptosis and necrosis (129, 130). Consistent with this notion, genetic deletion of PARP rendered mice resistant to experimental stroke (131), providing rationale for developing PARP inhibitors as the cytoprotective agents.

Several PARP-1 inhibitors were designed based on 3-amino-benzamide (132) (Fig. 5a), but these molecules lacked specificity and potency (133). Another early inhibitor, benadrostin is a natural product of actinomyces, and it was isolated from the culture broth of Streptomyces flavovirens as a competitive PARP inhibitor with Ki of 34 μM (134, 135) (Fig. 5b). Compounds PD128763 and Nu1025 (Fig. 5c) (136), based on benadrostin, showed high in vitro affinity for PARP-1, but still required high concentrations (10-100 μM) for chemopotentiation (137, 138). Subsequently, significant effort has been spent on structure-based drug design utilizing information generated using known inhibitors, which ultimately led to the identification of a number of potent PARP inhibitors. Five of these molecules are currently in clinical trials for oncologic indications, that is, AG014699 (139) (Fig. 5d), KU59436 (As- traZeneca/KuDOS, London, UK), BSI-201 (BiPar, Brisbane, CA), INO-1001 (140-142), and GPI 21016 (MGI Pharma, Bloomington, MN). ABT-888 (143, 144) (Fig. 5e) is expected to enter clinical trials shortly (145). All of these molecules are low-nanomolar PARP inhibitors, which sensitize cancer cells effectively to chemotherapy or radiotherapy at nanomolar concentrations. In animal studies, all of these molecules are well tolerated and effectively synergize with multiple DNA-damaging anticancer agents.

Conversely, PARP inhibitors showed significant promise as cytoprotective agents in animal models of inflammation, stroke, Parkinson’s disease, spinal cord injury, and myocardial infarction (146-148). In addition, PARP inhibitors showed activity in inhibiting various types of injury associated with type 1 and 2 diabetes, including neuropathy, retinopathy, and renopathy, as well as beta cell death in a streptozotocin-injection model of type 1 diabetes (149). Overall, PARP has emerged as a very promising therapeutic target, especially in treating cancer, whereas a general lack of success in developing cytoprotective treatments has made this direction of PARP inhibitor development more challenging.

 

 

Figure 5. Structures of small-molecule inhibitors of nonapoptotic cell death. Inhibitors of: (a-e) PARP-1, (f) HSP90, (i, j) mitochondrial respiratory complexes I and (k) complexes 2, (l) phosphatidylcholine-specific phospholipase C, (m) acid sphingomyelinase, (n) NADPH oxidase, (o) JNK kinase, (p-s) necroptosis, (t-w) MPTP, and (g, h) antioxidants. (a) 3-aminobenzamide, (b) benadrostin, (c) Nu1025, 8-hydroxy-2-methylquinazolin-4(3H)-one, (d) AG014699, (e) ABT-888, (f) geldanamycin, (g) 3-tert-butyl-4-hydroxyanisol, (h) butylhydroxytoluene, (i) amytal, (j) rotenone, (k) thenoyltrifluoroacetone, (l) D609, (m) desipramine, (n) diphenyleneiodonium chloride, (o) SP600125, (p) necrostatin-1, (q) 7-Cl-Necrostatin-1, (r) necrostatin-3, (s) necrostatin-5, (t) cyclosporin A (CsA), (u) bongkrekig acid, (v) rasagiline, and (w) promethazine.

 

Necroptosis

Necroptosis or programmed necrosis is a novel type of regulated nonapoptotic cell death that was recently described by several research groups (150-153). It was found that in some cell types, stimulation of DR with their cognate ligands (TNFa, FasL, or TRAIL) under specific conditions where apoptosis is inhibited leads to the cellular demise with necrotic morphological features. Similar observations were also reported for cell death induced by some oncogenes (Ras, cMyc) (154, 155), chemotherapeutic agents (etoposide, camptothecin, and stau- rosporine) (156-158), and viral and bacterial agents (159, 160). This unique type of cell demise shares the characteristics of both apoptosis (as a regulated form of cell death) and unregulated pathologic necrosis (by the cellular morphology) (161). Thus, discovery of necroptosis may offer an opportunity for therapeutic targeting the pathologic necrosis, because, in contrary to previously accepted views, it may represent a regulated and, therefore, specifically inhibitable form of cell death. Activation of necrosis-like death under apoptosis-suppressive conditions has been observed in various mouse models of acute pathologic death, including experimental pancreatitis (162) and multiple organ failure (163).

The signaling pathway of necroptosis is just beginning to emerge with Holler et al. (150) establishing that Ser/Thr kinase activity of DR-associated adaptor molecule, RIP1, is a key specific upstream activator of necroptosis. RIP1 kinase is a client protein of the molecular chaperone heat shock protein 90 (Hsp90), and inhibition of Hsp90 in the cells by the small molecule geldanamycin was found to result in efficient proteasome-mediated degradation of the RIP1 (164). As a result, geldanamycin (Fig. 5f) has been found to inhibit the activation of necroptosis in human Jurkat T cells (150). However, this effect is not specific to necroptosis as geldanamycin also was shown to block the RIP1-dependent NF-kB activation efficiently (164), which is independent of the RIP1 kinase activity, unlike necroptosis (165). In another study, Temkin et al. (166) have suggested that RIP1 may translocate to the mitochondria and lead to disruption of the VDAC/ANT/Cyclophilin D complex through an unidentified indirect mechanism.

Although the mechanisms of necrotic cell death downstream of RIP1 are mostly unclear, inhibition of certain cell-signaling pathways has been found to attenuate necroptotic cell death. Overproduction of reactive oxygen species (ROS) is a hallmark of necrotic cell death and is a prominent part of necroptosis in some systems (161). ROS production in conditions of necrotic cell death is mediated by the mitochondrial respiratory chain complexes 1 and 2 (167) and/or through formation of RIP1/Rac1/ nicotinamide adenine dinucleotide phosphate (NADPH) oxidase complex resulting in an oxidative burst (168). The antioxidant 3-tert-butyl-4-hydroxyanisol (BHA) (Fig. 5g) has been found to be effective in blocking necroptosis in mouse fibrosarcoma L929 cells and in mouse embryonic fibroblasts (169), but not in human Jurkat T cells (161). Based on the data that closely related antioxidant butylhydroxytoluene (BHT) (Fig. 5H) was not as effective in inhibiting necrotic cell death as BHA, it was proposed that additional inhibitory activities of BHA, like inhibition of mitochondrial Complex 1 and/or of lipid peroxidation, may be critical for its inhibition of necroptosis (169). Consistent with the role of respiratory chain in ROS generation during necroptosis, inhibitors of complexes 1 (rotenone, amytal) (Fig. 5j, 5i) and 2 (thenoyltrifluoroacetone) (Fig. 5k), but not of complex 4 (cytochrome c oxidase) of the mitochondrial respiratory chain, have provided marked attenuation of cell death (167).

Autophagy is an important large-scale cellular catabolic process (see Reference 170 for review), and it is prominently activated as a part of necroptosis in many systems (161, 171-173). However, inhibition of autophagy with 3-methyladenine has been found to inhibit necroptosis in some cell lines, such as mouse fibrosacroma L929 cells, but not in the other cell lines (human Jurkat T cells or mouse embryonic fibroblasts), which suggests cell type-specific contribution of autophagy to necroptotic cell demise (161).

Activation of acid sphingomyelinase (A-SMase) and ceramide production were also shown to contribute to DR-induced necrosis (174). Inhibition of A-SMase activation by small-molecule inhibitors D609 (Fig. 5l) and desipramine (Fig. 5m) has been reported to attenuate necroptosis (174). D609 inhibits A-SMase induction indirectly by inhibiting the upstream-acting phosphatidylcholine-specific phospholipase C, whereas desipramine causes rapid and irreversible degradation of A-SMase. Finally, inhibition of NADPH oxidase/ C-Jun N-terminal kinases (JNK) axis of the pathway using siRNA tools has been shown to attenuate necroptosis (168). Although it has not yet been tested directly, it is likely that small-molecule inhibitors of NADPH oxidase, such as diphenyleneiodonium chloride (DPI) (Fig. 5n), and of JNK kinase, such as SP600125, (Fig. 5o), may interfere with necroptosis activation. It is important to note that none of the above-mentioned inhibitors is specific for necroptosis, because their target proteins are involved in a wide range of cellular regulatory networks. Additional analysis of the mechanisms of specific activation of the downstream pathways of necroptosis by RIP1 kinase will be important for developing more specific strategies for necroptosis inhibition.

The first specific inhibitor of necroptosis, necrostatin-1 (Nec-1), was identified by Degterev et al. (161) in a cell-based screen of ~15,000 compounds. Nec-1 (Fig. 5p) efficiently blocked necrotic death of human monocytic U937 cells stimulated with TNFa in the presence of broad-spectrum caspase inhibitor zVAD-fmk and other instances of necroptotic death (161). Although Nec-1 did not inhibit either activation of apoptosis or NF-kB by TNFa, it completely eliminated all the manifestations of cellular necrosis. Furthermore, optimized derivatives of this molecule were reported with EC50 in the cells of 50 nM (Fig. 5q) (175). Additional screening resulted in identification of the other potent inhibitors of necroptosis, termed Nec-3 and Nec-5 (Fig. 5r, s) (175, 176), which created a unique panel of nanomolar inhibitors of this process. Based on implication that necroptosis is responsible, at least partially, for the pathologic necrosis, the necrostatins were tested for the cytoprotective effects in in vivo rodent models of acute organ injury, which included cerebral (161) and cardiac (177) ischemia and brain trauma (178). Necrostatins were found to provide significant cytoprotective effect and functional improvement in multiple paradigms of acute injury. In the case of brain ischemia, Nec-1 displayed protection when administered up to 6 hours after 2-hour middle cerebral artery occlusion, which suggested that necroptosis may represent a delayed and, hence, therapeutically targetable injury component. Furthermore, recent data showed that Nec-1 also inhibits necrotic cell death provoked in response to other pathologic stimuli in cellular assays, which includes high doses of glutamate (179), plant sterols (180), and the chemotherapeutic agent shikonin (180). These results suggest that importance of necroptosis likely extends beyond DR signaling, and it may represent a major novel component of acute pathologic injuries.

 

Mitochondrial PTP

The mitochondrial permeability transition (MPT) is the loss of the inner mitochondrial membrane impermeability to solutes caused by opening of the MPT pore (MPTP). In turn, this action results in a loss of mitochondrial function and provides a common mechanism implicated in activation of mitophagy/autophagy, apoptosis, and necrosis in different cell systems. Although the composition of MPTP is not fully settled, multiple studies suggest involvement of adenine nucleotide translocase (ANT) in the inner mitochondrial membrane, voltage-dependent anion channel (VDAC or porin) in the outer membrane, and cyclophilin D (CypD) in the matrix. Involvement of other proteins such as benzodiazepine receptor, hexokinase, creatine kinase, and Bax has also been proposed (181-183).

The first identified potent inhibitor of MPT is the cyclosporine A (CsA) (184) (Fig. 5t), which inhibits the interaction between CypD and ANT (180). Using isolated mitochondria, CsA was shown to inhibit MPT at submicromolar concentrations (185, 186). CsA is also a potent inhibitor of necrotic death in the cells, for example, induced by oxidative stress and in vivo, notably in models of ischemia/reperfusion injury of liver (187), brain and central nervous system (188, 189), and myocardium (190). This finding suggests that inhibition of MPT may be a promising general direction for treating ischemia/reperfusion injury. This notion is supported by the resistance of observed in CypD-deficient mice to this form of injury (191, 192). The major drawbacks of therapeutic use of CsA are its transient and incomplete PTP inhibition as well as immunosuppressive side effects (193). In addition to CsA, other cytoprotective agents were also found to act as PTP inhibitors. These agents include bogkrekic acid, rasagiline, and promethazine (Fig. 5u, 5w), with both of the latter exerting neuroprotective effects in vivo (194, 195). However, in the case of these agents, the exact mechanism of the cytoprotective effects in vivo is yet to be established.

 

Conclusion

After the groundbreaking discoveries establishing that the cell death can result from intrinsic cellular regulation, rather than excessive external stress, significant focus has been placed on characterization of the molecular mechanism of this process. Several interesting targets, some of which are described in our review, have been identified, and multiple classes of small molecules have been developed and optimized successfully for use as the research tools and, in particular cases, even as the drug development candidates for various disorders, for which efficient treatments are currently unavailable. Compounds that target the enzymatic activities of cell death regulators, such as caspases, PARP, and HtrA2 inhibitors, as well as compounds that modulate the protein-protein interactions and protein conformations were developed successfully. Inhibitors have been identified using a variety of different screening approaches from tethering and SAR-by-NMR to rational drug design and high-throughput screening using enzymatic assays or whole-cell phenotypic readouts. Therefore, chemical inhibitors of cell death represent a very extensive and diverse small-molecule development effort, which attests to the importance of cell death regulatory pathways. In some instances, for example, of necroptosis inhibitors, small molecules provided very useful means for uncovering functional significance of the regulation of novel modes of cell death, even in the absence of the specific protein target information. In the other cases, for example, of the PARP inhibitors, small molecules proved very useful in showcasing the importance of the activity of their targets in the cells and in vivo, prompting additional analysis of the biological regulation associated with these protein factors. Finally, in the case of well-validated targets, such as caspases or Bcl-2 family, specific small-molecule inhibitors confirmed validity of targeting these regulators and the apoptotic pathway in general for the treatment of human disease. They provide exciting new lead compounds for therapeutic development. Overall, these findings emphasize that the availability of tools and methods for fast and efficient identification of small molecule modulators of important biological processes, stemming from rapid development of synthetic chemistry and small molecule screening technologies, provide a very valuable compliment to biologic research.

 

References

1. Kerr JF, Wyllie AH, Currie AR. Apoptosis: a basic biological phenomenon with wide-ranging implications in tissue kinetics. Br. J. Cancer 1972; 26:239-257.

2. Wyllie AH, Kerr JF, Currie AR. Cell death: the significance of apoptosis. Int. Rev. Cytol. 1980; 68:251-306.

3. Scaffidi C, et al. Two CD95 (APO-1/Fas) signaling pathways. EMBO J. 1998; 17:1675-1687.

4. Li H, et al. Cleavage of BID by caspase 8 mediates the mitochondrial damage in the Fas pathway of apoptosis. Cell 1998; 94:491-501.

5. Luo X, et al. Bid, a Bcl2 interacting protein, mediates cytochrome c release from mitochondria in response to activation of cell surface death receptors. Cell 1998; 94:481-490.

6. Fischer U, Stroh C, Schulze-Osthoff K. Unique and overlapping substrate specificities of caspase-8 and caspase-10. Oncogene 2006; 25:152-159.

7. Korsmeyer SJ, et al. Pro-apoptotic cascade activates BID, which oligomerizes BAK or BAX into pores that result in the release of cytochrome c. Cell Death Differ. 2000; 7:1166-1173.

8. Li P, et al. Cytochrome c and dATP-dependent formation of Apaf-1/caspase-9 complex initiates an apoptotic protease cascade. Cell 1997; 91:479-489.

9. Acehan D, et al. Three-dimensional structure of the apoptosome: implications for assembly, procaspase-9 binding, and activation. Mol. Cell 2002; 9:423-432.

10. Slee EA, et al. Ordering the cytochrome c-initiated caspase cascade: hierarchical activation of caspases-2, -3, -6, -7, -8, and -10 in a caspase-9-dependent manner. J. Cell Biol. 1999; 144:281-292.

11. Roset, R., L. Ortet, and G. Gil-Gomez, Role of Bcl-2 family members on apoptosis: what we have learned from knock-out mice. Front. Biosci. 2007; 12:4722-4730.

12. Degterev A, Boyce M, Yuan J. A decade of caspases. Oncogene 2003; 22:8543-8567.

13. Ekshyyan O, Aw TY. Apoptosis in acute and chronic neurological disorders. Front. Biosci. 2004; 9:1567-1576.

14. Kreuter M, et al. Stroke, myocardial infarction, acute and chronic inflammatory diseases: caspases and other apoptotic molecules as targets for drug development. Arch. Immunol. Ther. Exp. (Warsz), 2004; 52:141-155.

15. Thornberry NA, et al. A novel heterodimeric cysteine protease is required for interleukin-1 beta processing in monocytes. Nature 1992; 356:768-774.

16. Kang SJ, et al. Distinct downstream pathways of caspase-11 in regulating apoptosis and cytokine maturation during septic shock response. Cell Death Differ. 2002; 9:1115-1125.

17. Wang S, et al. Murine caspase-11, an ICE-interacting protease, is essential for the activation of ICE. Cell 1998; 92:501-509.

18. Earnshaw WC, Martins LM, Kaufmann SH. Mammalian caspases: structure, activation, substrates, and functions during apoptosis. Annu. Rev. Biochem. 1999; 68:383-424.

19. Hengartner MO. The biochemistry of apoptosis. Nature 2000; 407:770-776.

20. Putt KS, et al. Small-molecule activation of procaspase-3 to caspase-3 as a personalized anticancer strategy. Nat. Chem. Biol. 2006; 2:543-550.

21. O’Brien T, Lee D. Prospects for caspase inhibitors. Mini. Rev. Med. Chem. 2004; 4:153-165.

22. Thornberry NA, et al. A combinatorial approach defines specificities of members of the caspase family and granzyme B. Functional relationships established for key mediators of apoptosis. J. Biol. Chem. 1997; 272:17907-17911.

23. Thornberry NA. Caspases: key mediators of apoptosis. Chem. Biol. 1998; 5:R97-R103.

24. Garcia-Calvo M, et al. Inhibition of human caspases by peptidebased and macromolecular inhibitors. J. Biol. Chem. 1998; 273:32608-32613.

25. Hara H, et al. Inhibition of interleukin 1beta converting enzyme family proteases reduces ischemic and excitotoxic neuronal damage. Proc. Natl. Acad. Sci. U. S. A., 1997; 94:2007-2012.

26. Schotte P, et al. Non-specific effects of methyl ketone peptide inhibitors of caspases. FEBS Lett. 1999; 442:117-121.

27. Lee D, et al. Potent and selective nonpeptide inhibitors of caspases 3 and 7. J. Med. Chem. 2001; 44:2015-2026.

28. Lee D, et al. Potent and selective nonpeptide inhibitors of caspases 3 and 7 inhibit apoptosis and maintain cell functionality. J. Biol. Chem. 2000; 275:16007-16014.

29. Okamoto Y, et al. Peptide based interleukin-1 beta converting enzyme (ICE) inhibitors: synthesis, structure activity relationships and crystallographic study of the ICE-inhibitor complex. Chem. Pharm. Bull. 1999; 47:11-21.

30. Cornelis S, et al. Inflammatory caspases: targets for novel therapies. Curr. Pharm. Des. 2007; 13:367-385.

31. Linton SD. Caspase inhibitors: a pharmaceutical industry perspective. Curr. Top. Med. Chem. 2005; 5:1697-1717.

32. Randle JC, et al. ICE/Caspase-1 inhibitors as novel anti-inflammatory drugs. Expert Opin. Investig. Drugs 2001; 10:1207-1209.

33. Bayes M, Rabasseda X, and Prous JR. Gateways to clinical trials. Methods Find Exp. Clin. Pharmacol. 2003; 25:53-76.

34. Siegmund B, Zeitz M. Pralnacasan (vertex pharmaceuticals). IDrugs 2003; 6:154-158.

35. Stack JH, et al. IL-converting enzyme/caspase-1 inhibitor VX-765 blocks the hypersensitive response to an inflammatory stimulus in monocytes from familial cold autoinflammatory syndrome patients. J. Immunol. 2005; 175:2630-2634.

36. Wannamaker W, et al. (S)-1-((S)-2-{(1-(4-amino-3-chloro-phenyl)-methanoyl)-amino}-3,3-dimethyl-butanoy 1)-pyrrolidine-2-carboxylic acid ((2R,3S)-2-ethoxy-5-oxo-tetrahydro-furan-3-yl)-amide (VX-765), an orally available selective interleukin (IL)-converting enzyme/caspase-1 inhibitor, exhibits potent antiinflammatory activities by inhibiting the release of IL-1beta and IL-18. J. Pharmacol. Exp. Ther. 2007; 321:509-516.

37. Hoglen NC, et al. A caspase inhibitor, IDN-6556, ameliorates early hepatic injury in an ex vivo rat model of warm and cold ischemia. Liver Transpl. 2007; 13:361-366.

38. Linton SD, et al. First-in-class pan caspase inhibitor developed for the treatment of liver disease. J. Med. Chem. 2005; 48:6779-6782.

39. Ueno Y, et al. Orally-administered caspase inhibitor PF- 03491390 is retained in the liver for prolonged periods with low systemic exposure, exerting a hepatoprotective effect against alpha-Fas-induced liver injury in a mouse model. J. Pharmacol. Sci. 2007; 105:201-205.

40. Pockros PJ, et al. Oral IDN-6556, an antiapoptotic caspase inhibitor, may lower aminotransferase activity in patients with chronic hepatitis C. Hepatology 2007; 46:324-329.

41. Poordad FF. IDN-6556 Idun Pharmaceuticals Inc. Curr. Opin. Investig. Drugs 2004; 5:1198-1204.

42. Erlanson DA, et al. In situ assembly of enzyme inhibitors using extended tethering. Nat. Biotechnol. 2003; 21:308-314.

43. Choong IC, et al. Identification of potent and selective small-molecule inhibitors of caspase-3 through the use of extended tethering and structure-based drug design. J. Med. Chem. 2002; 45:5005-5022.

44. Youle RJ, Strasser A. The BCL-2 protein family: opposing activities that mediate cell death. Nat. Rev. Mol. Cell Biol. 2008; 9:47-59.

45. O’Neill J, et al. Promises and challenges of targeting Bcl-2 anti-apoptotic proteins for cancer therapy. Biochim. Biophys. Acta 2004; 1705:43-51.

46. Scorrano L, Korsmeyer SJ. Mechanisms of cytochrome c release by proapoptotic BCL-2 family members. Biochem. Biophys. Res. Commun. 2003; 304:437-444.

47. Adams JM, Cory S. The Bcl-2 apoptotic switch in cancer development and therapy. Oncogene 2007; 26:1324-1337.

48. Chipuk JE, Green DR. How do BCL-2 proteins induce mitochondrial outer membrane permeabilization? Trends Cell. Biol. In press.

49. Sattler M, et al. Structure of Bcl-xL-Bak peptide complex: recognition between regulators of apoptosis. Science 1997; 275:983-986.

50. Diaz JL, et al. A common binding site mediates heterodimerization and homodimerization of Bcl-2 family members. J. Biol. Chem. 1997; 272:11350-11355.

51. Ottilie S, et al. Dimerization properties of human BAD. Identification of a BH-3 domain and analysis of its binding to mutant BCL-2 and BCL-XL proteins. J. Biol. Chem. 1997; 272:30866-30872.

52. Cosulich SC, et al. Regulation of apoptosis by BH3 domains in a cell-free system. Curr. Biol. 1997; 7:913-920.

53. Wang JL, et al. Cell permeable Bcl-2 binding peptides: a chemical approach to apoptosis induction in tumor cells. Cancer Res. 2000; 60:1498-1502.

54. Holinger EP, Chittenden T, Lutz RJ. Bak BH3 peptides antagonize Bcl-xL function and induce apoptosis through cytochrome c-independent activation of caspases. J. Biol. Chem. 1999; 274:13298-13304.

55. Kashiwagi H, et al. TAT-Bim induces extensive apoptosis in cancer cells. Ann. Surg. Oncol. 2007; 14:1763-1771.

56. Walensky LD, et al. Activation of apoptosis in vivo by a hydrocarbon-stapled BH3 helix. Science 2004; 305:1466-1470.

57. Oh KJ, et al. A membrane-targeted BID BCL-2 homology 3 peptide is sufficient for high potency activation of BAX in vitro. J. Biol. Chem. 2006; 281:36999-37008.

58. Wang JL, et al. Structure-based discovery of an organic compound that binds Bcl-2 protein and induces apoptosis of tumor cells. Proc. Natl. Acad. Sci. U. S. A. 2000; 97:7124-7129.

59. Manero F, et al. The small organic compound HA14-1 prevents Bcl-2 interaction with Bax to sensitize malignant glioma cells to induction of cell death. Cancer Res. 2006; 66:2757-2764.

60. Wang G, et al. Structure-based design of potent small-molecule inhibitors of anti-apoptotic Bcl-2 proteins. J. Med. Chem. 2006; 49:6139-6142.

61. Tang G, et al. Pyrogallol-based molecules as potent inhibitors of the antiapoptotic Bcl-2 proteins. J. Med. Chem. 2007; 50:1723-1726.

62. Tang G, et al. Structure-based design of flavonoid compounds as a new class of small-molecule inhibitors of the anti-apoptotic Bcl-2 proteins. J. Med. Chem. 2007; 50:3163-3166.

63. Mohammad RM, et al. Preclinical studies of TW-37, a new nonpeptidic small-molecule inhibitor of Bcl-2, in diffuse large cell lymphoma xenograft model reveal drug action on both Bcl-2 and Mcl-1. Clin. Cancer Res. 2007; 13:2226-2235.

64. Degterev A, et al. Identification of small-molecule inhibitors of interaction between the BH3 domain and Bcl-xL. Nat. Cell Biol. 2001; 3:173-182.

65. Lugovskoy AA, et al. A novel approach for characterizing protein ligand complexes: molecular basis for specificity of small-molecule Bcl-2 inhibitors. J. Am. Chem. Soc. 2002; 124:1234-1240.

66. Kitada S, et al. Discovery, characterization, and structure-activity relationships studies of proapoptotic polyphenols targeting B-cell lymphocyte/leukemia-2 proteins. J. Med. Chem. 2003; 46:4259-4264.

67. Mohammad RM, et al. Preclinical studies of a nonpeptidic small-molecule inhibitor of Bcl-2 and Bcl-X(L) ((-)-gossypol) against diffuse large cell lymphoma. Mol. Cancer Ther. 2005; 4:13-21.

68. Xu L, et al. (-)-Gossypol enhances response to radiation therapy and results in tumor regression of human prostate cancer. Mol. Cancer Ther. 2005; 4:197-205.

69. Chan SL, et al. Identification of chelerythrine as an inhibitor of BclXL function. J. Biol. Chem. 2003; 278:20453-20456.

70. Zhang YH, et al. Chelerythrine and sanguinarine dock at distinct sites on BclXL that are not the classic BH3 binding cleft. J. Mol. Biol. 2006; 364:536-549.

71. Shuker SB, et al. Discovering high-affinity ligands for proteins: SAR by NMR. Science 1996; 274:1531-1534.

72. Oltersdorf T, et al. An inhibitor of Bcl-2 family proteins induces regression of solid tumours. Nature 2005; 435:677-681.

73. Letai A. BCL-2: found bound and drugged! Trends Mol. Med. 2005; 11:442-444.

74. Konopleva M, et al. Mechanisms of apoptosis sensitivity and resistance to the BH3 mimetic ABT-737 in acute myeloid leukemia. Cancer Cell 2006; 10:375-388.

75. van Delft MF, et al. The BH3 mimetic ABT-737 targets selective Bcl-2 proteins and efficiently induces apoptosis via Bak/Bax if Mcl-1 is neutralized. Cancer Cell 2006; 10:389-399.

76. Campas C, et al. Bcl-2 inhibitors induce apoptosis in chronic lymphocytic leukemia cells. Exp. Hematol. 2006; 34:1663-1669.

77. Perez-Galan P, et al. The BH3-mimetic GX15-070 synergizes with bortezomib in mantle cell lymphoma by enhancing Noxa-mediated activation of Bak. Blood 2007; 109:4441-4449.

78. Trudel S, et al. Preclinical studies of the pan-Bcl inhibitor obatoclax (GX015-070) in multiple myeloma. Blood 2007;109: 5430-5438.

79. Witters LM, et al. Synergistic inhibition of breast cancer cell lines with a dual inhibitor of EGFR-HER-2/neu and a Bcl-2 inhibitor. Oncol. Rep. 2007; 17:465-469.

80. Tzung SP, et al. Antimycin A mimics a cell-death-inducing Bcl-2 homology domain 3. Nat. Cell Biol. 2001; 3:183-191.

81. Schimmer AD, et al. Receptor- and mitochondrial-mediated apoptosis in acute leukemia: a translational view. Blood 2001; 98:3541-3553.

82. Schimmer AD, Dalili S. Targeting the IAP family of caspase inhibitors as an emerging therapeutic strategy. Hematol. Am. Soc. Hematol. Educ. Prog. 2005: 215-219.

83. Altieri DC. The molecular basis and potential role of survivin in cancer diagnosis and therapy. Trends Mol. Med. 2001 ;7:542-547.

84. Ambrosini G, Adida C, Altieri DC. A novel anti-apoptosis gene, survivin, expressed in cancer and lymphoma. Nat. Med. 1997; 3:917-921.

85. Li F, et al. Control of apoptosis and mitotic spindle checkpoint by survivin. Nature 1998; 396:580-584.

86. O’Connor DS, et al. Regulation of apoptosis at cell division by p34cdc2 phosphorylation of survivin. Proc. Natl. Acad. Sci. U. S. A. 2000; 97:13103-13107.

87. O’Connor DS, et al. A p34(cdc2) survival checkpoint in cancer. Cancer Cell 2002; 2:43-54.

88. Zaffaroni N, et al. Expression of the anti-apoptotic gene survivin correlates with taxol resistance in human ovarian cancer. Cell Mol. Life Sci. 2002; 59:1406-1412.

89. Fortugno P, et al. Regulation of survivin function by Hsp90. Proc. Natl. Acad. Sci. U. S. A. 2003; 100(24): p. 13791-6.

90. Plescia J, et al. Rational design of shepherdin, a novel anticancer agent. Cancer Cell. 2005; 7:457-468.

91. Gyurkocza B, et al. Antileukemic activity of shepherdin and molecular diversity of hsp90 inhibitors. J. Natl. Cancer Inst. 2006; 98:1068-1077.

92. Meli M, et al. Small-molecule targeting of heat shock protein 90 chaperone function: rational identification of a new anticancer lead. J. Med. Chem. 2006; 49:7721-7730.

93. Nakahara T, et al. YM155, a novel small-molecule survivin suppressant, induces regression of established human hormone- refractory prostate tumor xenografts. Cancer Res. 2007; 67:8014-8021.

94. Scott FL, et al. XIAP inhibits caspase-3 and -7 using two binding sites: evolutionarily conserved mechanism of IAPs. EMBO J. 2005; 24:645-655.

95. Shiozaki EN, et al. Mechanism of XIAP-mediated inhibition of caspase-9. Mol. Cell 2003; 11:519-527.

96. Du C, et al. Smac, a mitochondrial protein that promotes cytochrome c-dependent caspase activation by eliminating IAP inhibition. Cell 2000; 102:33-42.

97. Verhagen AM, et al. Identification of DIABLO, a mammalian protein that promotes apoptosis by binding to and antagonizing IAP proteins. Cell 2000; 102:43-53.

98. Huang Y, et al. Requirement of both the second and third BIR domains for the relief of X-linked inhibitor of apoptosis protein (XIAP)-mediated caspase inhibition by Smac. J. Biol. Chem. 2003; 278:49517-49522.

99. Liu Z, et al. Structural basis for binding of Smac/DIABLO to the XIAP BIR3 domain. Nature 2000; 408:1004-1008.

100. Wu G, et al. Structural basis of IAP recognition by Smac/ DIABLO. Nature 2000; 408:1008-1012.

101. Huang Y, Lu M, Wu H. Antagonizing XIAP-mediated caspase-3 inhibition. Achilles’ heel of cancers? Cancer Cell 2004; 5:1-2.

102. Fulda S, et al. Smac agonists sensitize for Apo2L/TRAIL- or anticancer drug-induced apoptosis and induce regression of malignant glioma in vivo. Nat. Med. 2002; 8:808-815.

103. Yang L, et al. Predominant suppression of apoptosome by inhibitor of apoptosis protein in non-small cell lung cancer H460 cells: therapeutic effect of a novel polyarginine-conjugated Smac peptide. Cancer Res, 2003; 63:831-837.

104. Park CM, et al. Non-peptidic small molecule inhibitors of XIAP. Bioorg Med Chem Lett. 2005; 15:771-775.

105. Oost TK, et al. Discovery of potent antagonists of the antiapop- totic protein XIAP for the treatment of cancer. J. Med. Chem. 2004; 47:4417-4426.

106. Sun H, et al. Structure-based design of potent, conformationally constrained Smac mimetics. J. Am. Chem. Soc. 2004; 126:16686-16687.

107. Zobel K, et al. Design, synthesis, and biological activity of a potent Smac mimetic that sensitizes cancer cells to apoptosis by antagonizing IAPs. ACS Chem Biol, 2006; 1:525-533.

108. Li L, et al. A small molecule Smac mimic potentiates TRAIL- and TNFalpha-mediated cell death. Science 2004; 305:1471-1474.

109. Splan KE, Allen JE, McLendon GL. Biochemical basis for enhanced binding of Peptide dimers to x-linked inhibitor of apoptosis protein. Biochemistry 2007; 46:11938-11944.

110. Wu H, Tschopp J, Lin SC. Smac mimetics and TNFalpha: a dangerous liaison? Cell, 2007; 131:655-658.

111. Nikolovska-Coleska Z, et al. Discovery of embelin as a cell-permeable, small-molecular weight inhibitor of XIAP through structure-based computational screening of a traditional herbal medicine three-dimensional structure database. J. Med. Chem. 2004; 47:2430-2440.

112. Huang Y, et al. Structural basis of caspase inhibition by XIAP: differential roles of the linker versus the BIR domain. Cell 2001; 104:781-790.

113. Riedl SJ, et al. Structural basis for the inhibition of caspase-3 by XIAP. Cell, 2001; 104:791-800.

114. Wu TY, et al. Development and characterization of nonpeptidic small molecule inhibitors of the XIAP/caspase-3 interaction. Chem. Biol. 2003; 10:759-767.

115. Schimmer AD, et al. Small-molecule antagonists of apoptosis suppressor XIAP exhibit broad antitumor activity. Cancer Cell 2004; 5:25-35.

116. Wang Z, et al. Cellular, biochemical, and genetic analysis of mechanism of small molecule IAP inhibitors. J. Biol. Chem. 2004; 279:48168-48176.

117. Suzuki Y, et al. A serine protease, HtrA2, is released from the mitochondria and interacts with XIAP, inducing cell death. Mol. Cell 2001; 8:613-621.

118. Verhagen AM, et al. HtrA2 promotes cell death through its serine protease activity and its ability to antagonize inhibitor of apoptosis proteins. J. Biol. Chem. 2002; 277:445-454.

119. Vande Walle L, et al. Proteome-wide Identification of HtrA2/Omi Substrates. J. Proteome Res. 2007; 6:1006-1015.

120. Cilenti L, et al. Characterization of a novel and specific inhibitor for the pro-apoptotic protease Omi/HtrA2. J. Biol. Chem. 2003; 278:11489-11494.

121. Bhuiyan MS, Fukunaga K. Inhibition of HtrA2/Omi ameliorates heart dysfunction following ischemia/reperfusion injury in rat heart in vivo. Eur. J. Pharmacol. 2007; 557:168-177.

122. Klupsch K, Downward J. The protease inhibitor Ucf-101 induces cellular responses independently of its known target, HtrA2/Omi. Cell Death Differ. 2006; 13:2157-2159.

123. Boye AB, Hergenrother PJ. The promises and pitfalls of small molecule inhibition of poly (ADP-Ribose) glycohydrolase (PARG). Drug Discovery Research: New Frontiers in the PostGenomic Era. Huang Z, ed. 2007. John Wiley & Sons, Hoboken, NJ.

124. Menissier de Murcia J, et al. Functional interaction between PARP-1 and PARP-2 in chromosome stability and embryonic development in mouse. EMBO J. 2003; 22:2255-2263.

125. Schreiber V, et al. Poly(ADP-ribose) polymerase-2 (PARP-2) is required for efficient base excision DNA repair in association with PARP-1 and XRCC1. J. Biol. Chem. 2002; 277:23028-23036.

126. Zaremba T, Curtin NJ. PARP inhibitor development for systemic cancer targeting. Anticancer Agents Med. Chem. 2007; 7:515-523.

127. Bryant HE, et al. Specific killing of BRCA2-deficient tumours with inhibitors of poly(ADP-ribose) polymerase. Nature 2005; 434:913-917.

128. Farmer H, et al. Targeting the DNA repair defect in BRCA mutant cells as a therapeutic strategy. Nature 2005; 434:917-921.

129. Bouchard VJ, Rouleau M, Poirier GG. PARP-1, a determinant of cell survival in response to DNA damage. Exp. Hematol. 2003; 31:446-454.

130. Koh DW, Dawson TM, Dawson VL. Mediation of cell death by poly(ADP-ribose) polymerase-1. Pharmacol. Res. 2005; 52:5-14.

131. Eliasson MJ, et al. Poly(ADP-ribose) polymerase gene disruption renders mice resistant to cerebral ischemia. Nat. Med. 1997; 3:1089-1095.

132. Durkacz BW, et al. (ADP-ribose)n participates in DNA excision repair. Nature 1980; 283:593-596.

133. Banasik M, Ueda K. Inhibitors and activators of ADP-ribosylation reactions. Mol. Cell Biochem. 1994; 138:185-197.

134. Aoyagi T, et al. Benadrostin, new inhibitor of poly(ADP-ribose) synthetase, produced by actinomycetes. I. Taxonomy, production, isolation, physico-chemical properties and biological activities. J. Antibiot. 1988; 41:1009-1014.

135. Yoshida S, et al. Benadrostin, new inhibitor of poly(ADP-ribose) synthetase, produced by actinomycetes. II. Structure determination. J. Antibiot. 1988; 41:1015-1018.

136. Griffin RJ, et al. Resistance-modifying agents. 5. Synthesis and biological properties of quinazolinone inhibitors of the DNA repair enzyme poly(ADP-ribose) polymerase (PARP). J. Med. Chem. 1998; 41:5247-5256.

137. Delaney CA, et al. Potentiation of temozolomide and topotecan growth inhibition and cytotoxicity by novel poly(adenosine diphosphoribose) polymerase inhibitors in a panel of human tumor cell lines. Clin. Cancer Res. 2000; 6:2860-2867.

138. Suto, M.J., et al., Dihydroisoquinolinones: the design and synthesis of a new series of potent inhibitors of poly(ADP-ribose) polymerase. Anticancer Drug Des, 1991. 6(2): p. 107-17.

139. Thomas HD, et al. Preclinical selection of a novel poly(ADP-ribose) polymerase inhibitor for clinical trial. Mol. Cancer Ther. 2007; 6:945-956.

140. Clark RS, et al. Local administration of the poly(ADP-ribose) polymerase inhibitor INO-1001 prevents NAD+depletion and improves water maze performance after traumatic brain injury in mice. J. Neurotrauma 2007; 24:1399-1405.

141. Mason KA, et al. INO-1001, a novel inhibitor of poly(ADP-ribose) polymerase, enhances tumor response to doxorubicin. Invest. New Drugs. 2007.

142. Radovits T, et al. Poly(ADP-Ribose) polymerase inhibition improves endothelial dysfunction induced by hypochlorite. Exp. Biol. Med. 2007; 232:1204-1212.

143. Albert JM, et al. Inhibition of poly(ADP-ribose) polymerase enhances cell death and improves tumor growth delay in irradiated lung cancer models. Clin. Cancer Res. 2007; 13:3033-3042.

144. Donawho CK, et al. ABT-888, an orally active poly(ADP-ribose) polymerase inhibitor that potentiates DNA-damaging agents in preclinical tumor models. Clin. Cancer Res. 2007; 13:2728-2737.

145. Ratnam K, Low JA. Current development of clinical inhibitors of poly(ADP-ribose) polymerase in oncology. Clin. Cancer Res. 2007; 13:1383-1388.

146. de la Lastra CA, Villegas I, Sanchez-Fidalgo S. Poly(ADP-ribose) polymerase inhibitors: new pharmacological functions and potential clinical implications. Curr. Pharm. Des. 2007; 13:933-962.

147. Skaper SD. Poly(ADP-ribosyl)ation enzyme-1 as a target for neuroprotection in acute central nervous system injury. Curr. Drug Targets CNS Neurol. Disord. 2003; 2:279-291.

148. Szabo G, Bahrle S. Role of nitrosative stress and poly(ADP-ribose) polymerase activation in myocardial reperfusion injury. Curr. Vasc. Pharmacol. 2005; 3:215-220.

149. Gale EA. Molecular mechanisms of beta-cell destruction in IDDM: the role of nicotinamide. Horm. Res. 1996; 45(suppl 1):39-43.

150. Holler N, et al. Fas triggers an alternative, caspase-8-independent cell death pathway using the kinase RIP as effector molecule. Nat. Immunol. 2000; 1:489-495.

151. Kawahara A, et al. Caspase-independent cell killing by Fas-associated protein with death domain. J. Cell Biol. 1998; 143:1353-1360.

152. Vercammen D, et al. Inhibition of caspases increases the sensitivity of L929 cells to necrosis mediated by tumor necrosis factor. J. Exp. Med. 1998; 187:1477-1485.

153. Vercammen D, et al. Dual signaling of the Fas receptor: initiation of both apoptotic and necrotic cell death pathways. J. Exp. Med. 1998; 188:919-930.

154. Chi S, et al. Oncogenic Ras triggers cell suicide through the activation of a caspase-independent cell death program in human cancer cells. Oncogene 1999; 18:2281-2290.

155. McCarthy NJ, et al. Inhibition of Ced-3/ICE-related proteases does not prevent cell death induced by oncogenes, DNA damage, or the Bcl-2 homologue Bak. J. Cell Biol. 1997; 136:215-227.

156. Hirsch T, et al. The apoptosis-necrosis paradox. Apoptogenic proteases activated after mitochondrial permeability transition determine the mode of cell death. Oncogene 1997; 15:1573-1581.

157. Leist M, et al. Inhibition of mitochondrial ATP generation by nitric oxide switches apoptosis to necrosis. Exp. Cell Res. 1999; 249:396-403.

158. Sane, A.T. and R. Bertrand, Caspase inhibition in camptothecin-treated U-937 cells is coupled with a shift from apoptosis to transient G1 arrest followed by necrotic cell death. Cancer Res, 1999. 59(15): p. 3565-9.

159. Kalai M, et al. The caspase-generated fragments of PKR cooperate to activate full-length PKR and inhibit translation. Cell Death Differ. 2007; 14:1050-1059.

160. Xu Y, et al. Autophagy contributes to caspase-independent macrophage cell death. J. Biol. Chem. 2006; 281:19179-19187.

161. Degterev A, et al. Chemical inhibitor of nonapoptotic cell death with therapeutic potential for ischemic brain injury. Nat. Chem. Biol. 2005; 1:112-119.

162. Mareninova OA, et al. Cell death in pancreatitis: caspases protect from necrotizing pancreatitis. J. Biol. Chem. 2006; 281:3370-3381.

163. Cauwels A, et al. Caspase inhibition causes hyperacute tumor necrosis factor-induced shock via oxidative stress and phospholipase A2. Nat. Immunol. 2003; 4:387-393.

164. Lewis J, et al. Disruption of hsp90 function results in degradation of the death domain kinase, receptor-interacting protein (RIP), and blockage of tumor necrosis factor-induced nuclear factor-kappaB activation. J. Biol. Chem. 2000; 275:10519-10526.

165. Lee TH, et al. The kinase activity of Rip1 is not required for tumor necrosis factor-alpha-induced IkappaB kinase or p38 MAP kinase activation or for the ubiquitination of Rip1 by Traf2. J. Biol. Chem. 2004; 279:33185-33191.

166. Temkin V, et al. Inhibition of ADP/ATP exchange in receptorinteracting protein-mediated necrosis. Mol. Cell Biol. 2006; 26:2215-2225.

167. Fiers W, et al. More than one way to die: apoptosis, necrosis and reactive oxygen damage. Oncogene 1999; 18:7719-7730.

168. Kim, Y.S., et al., TNF-induced activation of the Nox1 NADPH oxidase and its role in the induction of necrotic cell death. Mol. Cell 2007; 26:675-687.

169. Festjens N, et al. Butylated hydroxyanisole is more than a reactive oxygen species scavenger. Cell Death Differ. 2006; 13:166-169.

170. Mizushima N. Autophagy: process and function. Genes Dev. 2007; 21:2861-2873.

171. Thorburn J, et al. Selective inactivation of a Fas-associated death domain protein (FADD)-dependent apoptosis and autophagy pathway in immortal epithelial cells. Mol. Biol. Cell 2005; 16:1189-1199.

172. Xue L, Fletcher GC, Tolkovsky AM. Autophagy is activated by apoptotic signalling in sympathetic neurons: an alternative mechanism of death execution. Mol. Cell Neurosci. 1999; 14:180/198.

173. Yu L, et al. Regulation of an ATG7-beclin 1 program of autophagic cell death by caspase-8. Science 2004; 304:1500-1502.

174. Thon L, et al. Ceramide mediates caspase-independent programmed cell death. FASEB J. 2005; 19:1945-1956.

175. Teng X, et al. Structure-activity relationship study of novel necroptosis inhibitors. Bioorg. Med. Chem. Lett. 2005; 15:5039-5044.

176. Wang K, et al. Structure-activity relationship analysis of a novel necroptosis inhibitor, Necrostatin-5. Bioorg. Med. Chem. Lett. 2007; 17:1455-1465.

177. Smith, C.C., et al., Necrostatin: a potentially novel cardioprotective agent? Cardiovasc Drugs Ther, 2007. 21(4): p. 227-33.

178. You Z, et al. Necrostatin-1 reduces histopathology and improves functional outcome after controlled cortical impact in mice. J. Cereb. Blood Flow Metab. 2007. In press.

179. Xu X, et al. Necrostatin-1 protects against glutamate-induced glutathione depletion and caspase-independent cell death in HT-22 cells. J. Neurochem. In press.

180. Bao L, et al. Sitosterol-containing lipoproteins trigger free sterol-induced caspase-independent death in ACAT-competent macrophages. J. Biol. Chem. 2006; 281:33635-33649.

181. Crompton M. The mitochondrial permeability transition pore and its role in cell death. Biochem. J. 1999; 341(Pt 2): 233-249.

182. Kim JS, He L, Lemasters JJ. Mitochondrial permeability transition: a common pathway to necrosis and apoptosis. Biochem. Biophys. Res. Commun. 2003; 304:463-470.

183. Zoratti M, Szabo I. The mitochondrial permeability transition. Biochim. Biophys. Acta 1995; 1241:139-176.

184. Crompton M, Virji S, Ward JM. Cyclophilin-D binds strongly to complexes of the voltage-dependent anion channel and the adenine nucleotide translocase to form the permeability transition pore. Eur. J. Biochem. 1998; 258:729-735.

185. Halestrap AP, et al. Cyclosporin A binding to mitochondrial cyclophilin inhibits the permeability transition pore and protects hearts from ischaemia/reperfusion injury. Mol. Cell Biochem. 1997; 174:167-172.

186. Nicolli A, et al. Interactions of cyclophilin with the mitochondrial inner membrane and regulation of the permeability transition pore, and cyclosporin A-sensitive channel. J. Biol. Chem. 1996; 271:2185-2192.

187. Saxton, N.E., et al., Cyclosporin A pretreatment in a rat model of warm ischaemia/reperfusion injury. J Hepatol, 2002. 36(2): p. 241-7.

188. Domanska-Janik, K., et al., Neuroprotection by cyclosporin A following transient brain ischemia correlates with the inhibition of the early efflux of cytochrome C to cytoplasm. Brain Res. Mol. Brain Res. 2004; 121:50-59.

189. Sato M, et al. Cyclosporin A reduces delayed motor neuron death after spinal cord ischemia in rabbits. Ann. Thorac. Surg. 2003; 75:1294-1299.

190. Squadrito F, et al. Cyclosporin-A reduces leukocyte accumulation and protects against myocardial ischaemia reperfusion injury in rats. Eur. J. Pharmacol. 1999; 364:159-168.

191. Nakagawa T, et al. Cyclophilin D-dependent mitochondrial permeability transition regulates some necrotic but not apoptotic cell death. Nature 2005; 434:652-658.

192. Schinzel AC, et al. Cyclophilin D is a component of mitochondrial permeability transition and mediates neuronal cell death after focal cerebral ischemia. Proc. Natl. Acad. Sci. U. S. A. 2005; 102:12005-12010.

193. Kroemer G, Galluzzi L, Brenner C. Mitochondrial membrane permeabilization in cell death. Physiol. Rev. 2007; 87:99-163.

194. Mandel S, et al. Mechanism of neuroprotective action of the anti-Parkinson drug rasagiline and its derivatives. Brain Res. Brain Res. Rev. 2005; 48:379-387.

195. Cleren C, et al. Promethazine protects against 1-methyl-4-phenyl- 1,2,3,6-tetrahydropyridine neurotoxicity. Neurobiol. Dis. 2005; 20:701-708.

 

Further Reading

Gangadhar NM, Stockwell BR. Chemical genetic approaches to probing cell death. Curr. Opin. Chem. Biol. 2007; 11:83-87.

Ghosh S, Nie A, An J, Huang Z. Structure-based virtual screening of chemical libraries for drug discovery. Curr. Opin. Chem. Biol. 2006; 10:194-202.

Horvath EM, Szabo C. Poly(ADP-ribose) polymerase as a drug target for cardiovascular disease and cancer: an update. Drug News Perspect. 2007; 20:171-181.

Huang Z. The chemical biology of apoptosis. Exploring protein-protein interactions and the life and death of cells with small molecules. Chem Biol. 2002; 9:1059-1072.

Hunter AM, LaCasse EC, Korneluk RG. The inhibitors of apoptosis (IAPs) as cancer targets. Apoptosis 2007; 12:1543-1568.

Degterev A, Boyce M, Yuan J. A decade of caspases. Oncogene 2003; 22:8543-8567.

Leary A, Johnston SR. Small molecule signal transduction inhibitors for the treatment of solid tumors. Cancer Invest. 2007; 25:347-365.

Rajapakse HA. Small molecule inhibitors of the XIAP protein-protein interaction. Curr. Top. Med. Chem. 2007; 7:966-971.

Reed JC. Apoptosis mechanisms: implications for cancer drug discovery. Oncology 2004; 18:11-20.

Stauffer SR. Small molecule inhibition of the Bcl-X(L)-BH3 protein-protein interaction: proof-of-concept of an in vivo chemopotentiator ABT-737. Curr. Top. Med. Chem. 2007; 7:961-965.

Talanian RV, Brady KD, Cryns VL. Caspases as targets for anti-inflammatory and anti-apoptotic drug discovery. J. Med. Chem. 2000; 43:3351-3371.

Wang S, Yang D, Lippman ME. Targeting Bcl-2 and Bcl-XL with non-peptidic small-molecule antagonists. Semin. Oncol. 2003; 30:133-142.

 

See Also

Kinases, selective and Nonselective Inhibitors of

Mitochondria: Topics in Chemical Biology

Cysteine Proteases and Cysteine Protease Inhibitors

Protein-Protein Interactions, Tools to Study

DNA Damage, Sensing of