Successfully reported this slideshow.
We use your LinkedIn profile and activity data to personalize ads and to show you more relevant ads. You can change your ad preferences anytime.
Review
A molecular view on signal transduction by the apoptosome
Thomas F. Reubold, Susanne Eschenburg ⁎
Institute for Bio...
2. Apaf-1
Upon discovery of the executioner caspase-3 the molecular deter-
minants for its activation were still obscure [...
1422 T.F. Reubold, S. Eschenburg / Cellular Signalling 24 (2012) 1420–1425
Different to the sensor in AAA+ ATPases, the sensor arginine of
Apaf-1 and of the related Ced-4 does not seem to have a hy...
level suggests a similar mode of autoinhibition. However, there is
still a controversy concerning the role of cytochrome c...
[50] M. Proell, S.J. Riedl, J.H. Fritz, A.M. Rojas, R. Schwarzenbacher, PLoS One 3 (4) (2008)
e2119.
[51] N. Yan, J. Chai,...
Upcoming SlideShare
Loading in …5
×
Upcoming SlideShare
Call 1 500 166 outsourcing jasa cleaning service jasa outsourcing cleaning service jakarta
Next
Download to read offline and view in fullscreen.

0

Share

Download to read offline

A molecular view on signal transduction by the apoptosome

Download to read offline

A molecular view on signal transduction by the apoptosome

Related Books

Free with a 30 day trial from Scribd

See all
  • Be the first to like this

A molecular view on signal transduction by the apoptosome

  1. 1. Review A molecular view on signal transduction by the apoptosome Thomas F. Reubold, Susanne Eschenburg ⁎ Institute for Biophysical Chemistry, Hannover Medical School, 30625 Hannover, Germany a b s t r a c ta r t i c l e i n f o Article history: Received 7 February 2012 Accepted 5 March 2012 Available online 13 March 2012 Keywords: Apoptosis Apaf-1 Apoptosome Caspase activation Apoptosomes are signaling platforms that initiate the dismantling of a cell during apoptosis. In mammals, assembly of the apoptosome is the pivotal point in the mitochondrial pathway of apoptosis, and is prompted by binding of cytochrome c to the apoptotic protease-activating factor 1 (Apaf-1) in the presence of ATP. The resulting wheel-like heptamer of seven molecules Apaf-1 and seven molecules cytochrome c binds and acti- vates the initiator caspase-9, which in turn ignites the downstream caspase cascade. In this review we discuss the molecular determinants for the formation of the mammalian apoptosome and caspase activation and describe the related signaling platforms in flies and nematodes. © 2012 Elsevier Inc. All rights reserved. Contents 1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1420 2. Apaf-1 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1421 3. Cytochrome c . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1421 4. ATP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1421 5. Structure of the Apaf-1 apoptosome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1423 6. Caspase activation by the Apaf-1 apoptosome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1423 7. The Apaf-1 orthologs Dark and Ced-4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1423 8. Concluding remarks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1424 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1424 1. Introduction Apoptosis is a highly regulated cellular mechanism in metazoans used to terminate superfluous or unwanted cells in a controlled way [1]. Apoptosis is crucial for normal human embryonic development [2,3] and the development of the immune system [4]. Deregulation of apoptosis is associated with severe pathologic conditions, e.g. can- cer and neurodegenerative diseases [5–8]. Two different pathways have emerged, which both lead to the activation of members of a class of cystein-dependent aspartate-specific proteases (caspases). Activation of the caspase cascade ultimately leads to the orderly deg- radation of the cell [9]. The extrinsic apoptotic pathway is induced by binding of death ligands to the ectodomains of their cognate death receptors [10] and leads to the activation of the initiator caspases 8 and 10. The intrinsic or mitochondrial apoptotic pathway is triggered by different kinds of cellular stress — e.g. DNA damaging agents including UV irradiation and cytotoxic drugs (reviewed in [11]). Predominance of pro-apoptotic stimuli eventually lead to the Bax (Bcl-2-associated x protein)- or Bak (Bcl-2 antagonist killer)-induced permeabilization of the outer mitochondrial membrane [12,13]. Per- meabilization is followed by the release of cytochrome c from the intermembrane space into the cytosol [13]. Binding of cytochrome c to cytosolic Apaf-1 (apoptotic protease-activating factor 1) in the presence of ATP yields a large complex of 1 MDa termed apoptosome, which serves as activation platform for procaspase-9 [14]. The holo- apoptosome formed by the apoptosome and caspase-9 in turn acti- vates the effector caspase-3 [15]. In this review we summarize the current knowledge about the constituents of the mammalian holo-apoptosome and discuss their role in the transduction of the death signal in the mitochondrial path- way of apoptosis. In doing so we highlight recent structural data and their implications for the mechanism of apoptosome formation and subsequent caspase activation. Cellular Signalling 24 (2012) 1420–1425 ⁎ Corresponding author. Tel.: +49 511 532 8655; fax: +49 511 532 2909. E-mail address: eschenburg.susanne@mh-hannover.de (S. Eschenburg). 0898-6568/$ – see front matter © 2012 Elsevier Inc. All rights reserved. doi:10.1016/j.cellsig.2012.03.007 Contents lists available at SciVerse ScienceDirect Cellular Signalling journal homepage: www.elsevier.com/locate/cellsig
  2. 2. 2. Apaf-1 Upon discovery of the executioner caspase-3 the molecular deter- minants for its activation were still obscure [16,17]. The molecules, which eventually activate caspase-3 in the mitochondrial branch of apoptosis, were subsequently identified as procaspase-9 [18], cyto- chrome c [19], and Apaf-1 [20]. Apaf-1 was recognized as human counterpart of Ced-4 of the nematode Caenorhabditis elegans [21]. Based on sequence homology Apaf-1 has been grouped into the STAND (signal transduction ATPases with numerous domains) family of proteins [22]. Whether the STAND family constitutes a subfamily of AAA+ (ATPases associated with diverse cellular activities) ATPases is debated [23,24]. Apaf-1 is expressed in various tissues and exists in at least four different splice forms [25]. The residue numbering through- out this review refers to the longest isoform termed Apaf-1-XL (resi- dues 1–1248). Apaf-1 contains three main domains: the N-terminal caspase recruitment domain (CARD), the nucleotide binding and olig- omerization domain (NOD), and the C-terminal regulatory WD40 re- peat domain (WRD) (Fig. 1). The CARD belongs to the death domain superfamily whose members share a common basic fold consisting of six bundled α-helices [26]. The Apaf-1 CARD binds its counterpart in procaspase-9 via homotypic interactions mostly involving charged residues [27]. The NOD (residues 108–586) harbors a binding site for adenine nucleotides and mediates oligomerization of Apaf-1 into the apoptosome complex. Several years ago the crystal structure of an N-terminal fragment of human Apaf-1 comprising the CARD and the NOD was determined [28]. The structure revealed a compact fold of the NOD and that the NOD can be divided into four subdomains. These subdomains were termed nucleotide-binding domain (NBD, residues 108–284), helical domain 1 (HD1, residues 285–365), winged-helix domain (WHD, res- idues 366–450), and helical domain 2 (HD2, residues 451–586). The nucleotide binding site of the NOD was occupied with an ADP mole- cule, which was in contact with residues from all four subdomains. Since the entire regulatory WRD was missing, the explanation of how Apaf-1 inhibits itself until cytochrome c binds had to wait until a crystal structure of full-length Apaf-1 was solved. The crystal structure of full-length murine Apaf-1 in the absence of cytochrome c and exogenous nucleotide [29] recently showed that the tandem β-propellers of the WRD do not block the CARD as pre- sumed previously. Rather, the WRD acts as a clamp which holds the subdomains of the NOD in place. Thus, the WRD prevents the confor- mational rearrangement of the NOD which is required for apopto- some formation [30,31] (see below for further details). 3. Cytochrome c The identification of cytochrome c as a crucial component of the mitochondrial pathway of apoptosis came as a surprise [19]. Given the well known function of cytochrome c in the respiratory chain, a role as signaling molecule appeared remarkable. However, the fact that cytochrome c is, in a healthy cell, confined to the mitochondrial intermembrane space while the executing part of the apoptosis ma- chinery is located in the cytosol makes the molecule well suited as de- cisive start signal. The strict spatial separation of signal and machinery minimizes the chance for accidental activation of the death cascade. Apaf-1 is able to bind cytochrome c both in the presence and ab- sence of nucleotide [20,32,33]. It was soon recognized that cyto- chrome c binds to the WRD and that this interaction is required to relieve the autoinhibition of Apaf-1 imposed by its WRD, since dele- tion of the WRD abolishes the need for cytochrome c in the assembly process [34,35]. Biophysical studies assessing the binding characteris- tics of cytochrome c to Apaf-1 suggested a stoichiometry of two cyto- chrome c molecules per molecule Apaf-1 [33,36]. However, subsequent cryo-EM studies clearly showed a 1:1 stoichiometry without any indication for a secondary cytochrome c binding site on Apaf-1 [30,31]. Globular density, which may be attributed to cyto- chrome c, is visible only between the β-propellers of individual Apaf-1 molecules within the apoptosome (see reference [31]). This observation is supported by the crystal structure of full length Apaf- 1 [29]. The faces of the β-propellers WD1 and WD2 show pronounced negative charges as would be expected for efficient binding of the positively charged cytochrome c. Binding of cytochrome c appears to cause a rotation of β-propeller WD1 [29], which detaches the NBD–HD1 subunit of the NOD from the propeller clamp (Fig. 1). The rotation of β-propeller WD1 enables and even actuates subsequent rotation of the NBD–HD1 subunit, which eventually leads to the extended conformation of Apaf-1 seen in the apoptosome [29,31]. Furthermore, cytochrome c might prevent re- closing of the opened NOD, since the rotated position of WD1 is steri- cally incompatible with the closed conformation of the NOD. 4. ATP A second crucial molecular event during apoptosome formation is the exchange of (d)ADP bound to Apaf-1 for exogenous (d)ATP [37]. The origin of the bound diphosphate is still a subject of controversy. According to a study performed in the lab of Xiadong Wang, mono- meric Apaf-1 is loaded with dATP [38]. Here, binding of cytochrome c prompts Apaf-1 to perform a single round of nucleotide hydrolysis. The hydrolytic energy would be needed to prime Apaf-1 for the ensu- ing large conformational changes, which in turn are then triggered by replacing dADP for exogenous dATP. Several other studies contradict these findings showing that recombinant monomeric full-length Apaf-1 as well as WD40-deleted Apaf-1 purified from different insect cell strains and from Escherichia coli contain ADP [28,29,39,40]. Con- sistently, a low but steady (d)ATPase activity of monomeric Apaf-1 in the absence of cytochrome c has been described [40–43]. For apoptosome assembly the bound ADP has to be exchanged for an exogenous nucleoside triphosphate in the presence of cytochrome c. Not only ATP or dATP but also non-hydrolyzable ATP analogs like AppNHp or AppCp fulfill this function, which emphasizes that chem- ical energy derived from nucleotide hydrolysis is not required at all during the assembly process. Rather, the presence of a γ-phosphate alone is sufficient to allow apoptosome formation [40]. The molecular effect of the nucleotide exchange is still obscure. However, comparing the nucleotide binding sites of different AAA+ ATPases and STAND proteins, the notion emerges that the determin- ing factor is a polar interaction of the γ-phosphate with a sensor res- idue. This residue is located at the tip of β-strand 4 as part of the so- called sensor 1 motif. In AAA+ ATPases the sensor residue is mostly an asparagine, but serine, threonine, aspartate, or histidine are also found [44,45]. Several studies suggest that sensor 1 is directly in- volved in ATP hydrolysis, since its mutation strongly decreases coop- erative ATPase activity in different proteins [46–48]. Most members of the AAA+ ATPase related family of STAND ATPases, to which Apaf-1 belongs, possess an arginine as sensor residue [49,50]. In Apaf-1 this arginine (R265) would be in interaction distance to the γ-phosphate group of an ATP molecule in the nucleotide binding site [29]. Such an interaction is also visible within crystal structures of Ced-4, both in the Egl-1 inhibited dimer [51] and in the apopto- some [52], as well as in the EM-derived model of the human apopto- some [31]. The importance of the sensor arginine has been confirmed for Apaf-1 and other STAND proteins. Functional analysis of an Apaf-1 R265S mutant revealed that the mutant protein does not yield func- tional apoptosomes [29]. In the bacterial transcription regulator MalT mutation of the putative sensor arginine R160 to alanine resulted in loss of activation [53]. The same was observed for the cor- responding mutation (R313A) in the plant resistance protein I-2 [54]. 1421T.F. Reubold, S. Eschenburg / Cellular Signalling 24 (2012) 1420–1425
  3. 3. 1422 T.F. Reubold, S. Eschenburg / Cellular Signalling 24 (2012) 1420–1425
  4. 4. Different to the sensor in AAA+ ATPases, the sensor arginine of Apaf-1 and of the related Ced-4 does not seem to have a hydrolytic function. Apaf-1 has no hydrolytic activity in the assembled state and also Apaf-1 assembly does not rely on nucleotide hydrolysis [39,40]. Ced-4, the only STAND protein for which atomic structures are available for both the inhibited and the assembled state, does not seem to possess hydrolytic activity at all, since in both states the bound nucleotide is ATP [51,52]. These circumstances point to a stabi- lizing role of the sensor arginine rather than a contribution to any cat- alytic activity. 5. Structure of the Apaf-1 apoptosome The first evidence for the existence of a high molecular weight complex as the caspase-activating species of Apaf-1 came from in vitro studies using recombinant purified proteins [43]. Gel filtration of Apaf-1 showed that the apparent molecular weight shifted from that of monomeric Apaf-1 to about 1 MDa if cytochrome c and dATP were added. The availability of in vitro reconstituted apoptosome en- abled the group of Christopher Akey to generate a first cryo-EM map to 27 Å that revealed that the apoptosome resembles a wheel with seven spokes. Each of the spokes consists of one molecule Apaf-1 and one molecule cytochrome c. Cytochrome c apparently docked between the tandem β-propellers of Apaf-1 at the rim of the disk. The hub was thought to be formed by the CARD and part of the NOD [55]. A subsequent EM reconstruction to 12.8 Å [30] allowed the placement of individual subdomains of the X-ray model of the human Apaf-1 CARD–NOD fragment [28] and of homology models of a six-bladed and a seven-bladed β-propeller. The resulting model dissects the hub into an inner ring formed by the seven CARDS and an outer ring formed by NBD–HD1 subunits bridged by WHDs. A modeling study of the Apaf-1 apoptosome complex challenged this view and pre- sented a model in which the protomer arrangement resembles that in AAA+ complexes [56]. Here, the hub is exclusively formed by sub- units of the NODs, and the CARDs form a crown that sits on top. A re- cent cryo-EM study to 9.5 Å confirmed these results [31]. The NBDs indeed form an inner ring reminiscent of the canonical AAA+ archi- tecture, whereas an outer ring contains HD1 and WHDs in an alternat- ing arrangement (Fig. 1). In the absence of procaspase-9 the CARDs are disordered but together with procaspase-9 CARDs form a disk that is flexibly tethered to the apoptosome. Implications of these findings for caspase stimulation will be discussed in Section 6. 6. Caspase activation by the Apaf-1 apoptosome Caspases are produced as zymogens, which are cleaved within their catalytic domains to yield a large and a small subunit [57]. Apo- ptotic caspases are divided into two classes according to their posi- tion and function within the apoptotic pathway: initiator or effector caspases [58]. Effector caspases are constitutive dimers lacking a pro- domain, whereas initiator caspases possess a prodomain and are mo- nomeric at physiological concentrations [57,58]. Effector caspases are activated by cleavage of the interdomain linker allowing for rearran- gement of the active site loops [59]. Although loop rearrangement seems to be a general necessity for caspase activation, linker cleavage is not a strict requirement for initiator caspase stimulation [60,61]. After several activator platforms for different initiator caspases had been identified, two alternative models for initiator caspase activation emerged: the proximity-induced dimerization model [15,62,63] and the induced conformation model [64–66]. The proximity-induced di- merization model is based on the observation that initiator caspase molecules are able to process and activate themselves if brought into close contact to each other [35,67–69]. It is thought that the active cas- pase species is a dimer where one monomer is catalytically active, whereas the second monomer is inactive [70]. Catalytic competence is achieved by rearrangement of the active site loops in the active mono- mer. In this view the respective activator platform, the Apaf-1 apopto- some in case of caspase-9, serves to achieve a local caspase concentration higher than the dissociation constant for dimer formation [62,63,71]. The induced conformation model assumes that the major de- terminant for caspase activation is not dimerization but an allosteric ac- tion of the activator platform on the caspase. This action induces a conformational change of the caspase which in turn increases its activi- ty. The nature of this conformational change is unknown and several variants of this model are conceivable (reviewed in [64]). The proximity-induced dimerization model is amongst others sup- ported by a study using a chimeric caspase containing the CARD of caspase-9 fused to the catalytic domain of caspase-8 [63]. The chimera was able to restore the activation of executioner caspase in caspase-9 depleted cytosolic extracts supplied with exogenous dATP and cyto- chrome c to about 70% of the wild type level. Support for the induced conformation model comes from the fact that a constitutively dimeric caspase-9 mutant is only slightly more active than wild-type caspase- 9 and can not be activated by the apoptosome [65]. Further evidence for the induced conformation model comes from a recent cryo-EM study on the caspase-9 containing holo-apoptosome of Apaf-1 [72]. A map calculated from holo-apoptosomes without symmetry restraints only showed density for a single procaspase-9 molecule located on the central hub. Moreover, the CARD disk assumes a tilted acentric ori- entation shifted away from the procaspase density. This suggests that the acentric CARD disk might impose steric hindrance on procaspase binding allowing only one catalytic procaspase-9 subunit to bind at a time. Unexpectedly, shortening of the procaspase prodomain linker by 14 or 24 residues severely compromises apoptosome dependent procas- pase activity. If activation would proceed via proximity-induced dimer- ization, a short prodomain linker of 6–10 residues should still be long enough to allow dimer assembly [73]. Moreover, there is evidence from the EM-study that the prodomain linker is involved in binding of procaspase-9 to the apoptosome [72]. This is well in line with the fact that phosphorylation of caspase-9 at position T125 in the prodomain linker leads to inhibition of apoptosis [74–76]. It is tempting to specu- late that the phosphorylation interferes with proper orientation of the catalytic domain on the apoptosome. Taken together, these data argue against a pure dimerization based activation mechanism but rather point towards a proximity-induced association with a procaspase monomer as the active species. 7. The Apaf-1 orthologs Dark and Ced-4 The components of apoptotic pathways have also been thor- oughly studied in the nematode Caenorhabditis elegans and the fruit- fly Drosophila melanogaster. The Drosophila Apaf-1 homolog Dark (Drosophila Apaf-1 related killer) shares the same principal domain structure as mammalian Apaf-1 [77–79]. A cryo-EM reconstruction to 6.9 Å of the Dark apoptosome shows structural similarities to the Apaf-1 apoptosome, i.e. a wheel-like appearance where the NODs form the hub and the WD40 tandem β-propellers form the rim of the wheel [80]. Different from the human apoptosome, the Dark apoptosome contains eight instead of seven spokes, each con- sisting of one Dark monomer. To date, no crystal structure of autoin- hibited Dark exists, but the structural similarity on the apoptosomal Fig. 1. A. Subdomain structure of Apaf-1. Subdomains are color-coded, linkers are shown in gray. B. Conformational states of Apaf-1, the cartoon representation is color coded as in 1A. Top left: Apaf-1 in its autoinhibited state. Top right: Apaf-1 monomer in its assembly competent conformation. The yellow β-propeller WD1 has adjusted its position to com- plete the binding site for cytochrome c between the inner faces of the two β-propellers; the blue-turquoise NBD–HD1 subunit is rotated to bare the oligomerization interfaces. Bot- tom: cartoon representation of the apoptosome (PDB code 3IZA). In the three states shown the CARD is not resolved [29,31] and is therefore not shown in the drawing. 1423T.F. Reubold, S. Eschenburg / Cellular Signalling 24 (2012) 1420–1425
  5. 5. level suggests a similar mode of autoinhibition. However, there is still a controversy concerning the role of cytochrome c in the activa- tion process (reviewed in [81,82]). Although Drosophila cytochrome c seems to play a role in certain developmental apoptotic situations [83–85], the bulk of the available data suggests that cytochrome c is not required for apoptosome formation [86–92]. This notion is sup- ported by the in vitro reconstitution of Dark apoptosomes, which does not require cytochrome c but occurs upon incubation with unphysiologically high dATP concentrations in the presence of EDTA [80,93]. In vivo, the caspase-9 homolog DRONC (Drosophila Nedd2-like caspase) [92,94] can be activated by ablation of DIAP1 (Drosophila inhibitor of apoptosis protein 1). DIAP1 is a fly homolog of the mam- malian IAPs (inhibitor of apoptosis proteins) and marks DRONC for proteasomal degradation via its E3-ubiquitin ligase activity [95,96]. This suggests that Dark is constitutively active. However, another study points to the existence of a yet unknown Dark activator [92]. So, the physiological regulation of Dark is far from clear and more research is necessary to elucidate the chain of events that lead to cell death in Drosophila. Ced-4 (cell-death abnormality 4), the C. elegans homolog of Apaf-1, possesses a CARD and an NOD, but no WRD. Unlike Apaf-1 Ced-4 is not autoinhibited but forms a complex with the Bcl-2 pro- tein Ced-9 to maintain its inhibited state [97]. The crystal structure of this complex reveals one molecule of Ced-9 bound to an asym- metric Ced-4 dimer [51]. There is no evidence for any mitochondrial participation in the signaling cascade and consequentially cyto- chrome c is not needed for Ced-4 activation. Activation is achieved by a third protein, Egl-1 (egg-laying defective 1), which binds to Ced-9, thereby releasing the Ced-4 dimer from the inhibitory com- plex [98]. The sole nematode caspase, Ced-3, is activated by an octa- meric apoptosome assembled from four Ced-4 dimers [52]. While the human and fly apoptosomes display a flat disk-like geometry, the Ced-4 apoptosome possesses a funnel-like structure. The CARDs of Ced-4 form two stacked tetramers, one of which contacts the narrow end of the funnel that is formed by the NODs. Investiga- tion of the mode of Ced-3 activation within the same study led to an unexpected result. A soluble CARD-deleted Ced-3 fragment (resi- dues 198–503) could be stimulated in the presence of Ced-4 apop- tosome [52]. This led to the hypothesis that the caspase binding sites responsible for stimulation are located within the funnel. If this were true, CARD interactions, which are indispensable for other caspase activation platforms, would not be required for exe- cution of nematode cell death. However, without further experi- ments employing full-length Ced-3 the proposed mechanism must remain speculative. 8. Concluding remarks The Apaf-1 apoptosome represents the central signaling hub within the mitochondrial pathway of apoptosis. In recent years con- siderable information has been gained about both the architecture and the function of apoptosomal complexes. Especially structural analyses using X-ray crystallography and cryo electron microscopy contributed substantially to the understanding of these molecular switches. However, there are still open questions about important aspects of the signal transduction process. The role of the nucleotide exchange has yet to be determined and the mechanism of caspase activation is even more subject of controversial discussion. Further- more, it is still unknown how Apaf-1 interacts with regulating pro- teins other than cytochrome c. In recent years numerous interaction partners of Apaf-1 have been identified, which inhibit or stimulate apoptosome assembly. Detailed knowledge about these interac- tions, as future studies will certainly deliver, may be the key to tar- get Apaf-1 for therapeutic purposes. The possibility to specifically modulate the function of Apaf-1 is highly desirable, given the cen- tral role of Apaf-1 in the mitochondrial pathway of apoptosis. References [1] R.C. Taylor, S.P. Cullen, S.J. Martin, Nature Reviews. Molecular Cell Biology 9 (3) (2008) 231–241. [2] P.M. Domingos, H. Steller, Current Opinion in Genetics & Development 17 (4) (2007) 294–299. [3] C. Twomey, J.V. McCarthy, Journal of Cellular and Molecular Medicine 9 (2) (2005) 345–359. [4] J.T. Opferman, Cell Death and Differentiation 15 (2) (2008) 234–242. [5] T.G. Cotter, Nature Reviews. Cancer 9 (7) (2009) 501–507. [6] K. Vermeulen, D.R. Van Bockstaele, Z.N. Berneman, Annals of Hematology 84 (10) (2005) 627–639. [7] O. Ekshyyan, T.Y. Aw, Current Neurovascular Research 1 (4) (2004) 355–371. [8] R.M. Friedlander, The New England Journal of Medicine 348 (14) (2003) 1365–1375. [9] C.J. de Almeida, R. Linden, Cellular and Molecular Life Sciences 62 (14) (2005) 1532–1546. [10] Z. Mahmood, Y. Shukla, Experimental Cell Research 316 (6) (2010) 887–899. [11] W.P. Roos, B. Kaina, Trends in Molecular Medicine 12 (9) (2006) 440–450. [12] S.W. Tait, D.R. Green, Nature Reviews. Molecular Cell Biology 11 (9) (2010) 621–632. [13] G. Kroemer, L. Galluzzi, C. Brenner, Physiological Reviews 87 (1) (2007) 99–163. [14] P. Li, D. Nijhawan, I. Budihardjo, S.M. Srinivasula, M. Ahmad, E.S. Alnemri, X. Wang, Cell 91 (4) (1997) 479–489. [15] J. Rodriguez, Y. Lazebnik, Genes & Development 13 (24) (1999) 3179–3184. [16] M. Tewari, L.T. Quan, K. O'Rourke, S. Desnoyers, Z. Zeng, D.R. Beidler, G.G. Poirier, G.S. Salvesen, V.M. Dixit, Cell 81 (5) (1995) 801–809. [17] D.W. Nicholson, A. Ali, N.A. Thornberry, J.P. Vaillancourt, C.K. Ding, M. Gallant, Y. Gareau, P.R. Griffin, M. Labelle, Y.A. Lazebnik, et al., Nature 376 (6535) (1995) 37–43. [18] H. Duan, K. Orth, A.M. Chinnaiyan, G.G. Poirier, C.J. Froelich, W.W. He, V.M. Dixit, The Journal of Biological Chemistry 271 (28) (1996) 16720–16724. [19] X. Liu, C.N. Kim, J. Yang, R. Jemmerson, X. Wang, Cell 86 (1) (1996) 147–157. [20] H. Zou, W.J. Henzel, X. Liu, A. Lutschg, X. Wang, Cell 90 (3) (1997) 405–413. [21] J. Yuan, H.R. Horvitz, Development 116 (2) (1992) 309–320. [22] D.D. Leipe, E.V. Koonin, L. Aravind, Journal of Molecular Biology 343 (1) (2004) 1–28. [23] J.P. Erzberger, J.M. Berger, Annual Review of Biophysics and Biomolecular Structure 35 (2006) 93–114. [24] M. Ammelburg, T. Frickey, A.N. Lupas, Journal of Structural Biology 156 (1) (2006) 2–11. [25] M.A. Benedict, Y. Hu, N. Inohara, G. Nunez, The Journal of Biological Chemistry 275 (12) (2000) 8461–8468. [26] H.H. Park, Y.C. Lo, S.C. Lin, L. Wang, J.K. Yang, H. Wu, Annual Review of Immunol- ogy 25 (2007) 561–586. [27] H. Qin, S.M. Srinivasula, G. Wu, T. Fernandes-Alnemri, E.S. Alnemri, Y. Shi, Nature 399 (6736) (1999) 549–557. [28] S.J. Riedl, W. Li, Y. Chao, R. Schwarzenbacher, Y. Shi, Nature 434 (7035) (2005) 926–933. [29] T.F. Reubold, S. Wohlgemuth, S. Eschenburg, Structure 19 (8) (2011) 1074–1083. [30] X. Yu, D. Acehan, J.F. Menetret, C.R. Booth, S.J. Ludtke, S.J. Riedl, Y. Shi, X. Wang, C.W. Akey, Structure 13 (11) (2005) 1725–1735. [31] S. Yuan, X. Yu, M. Topf, S.J. Ludtke, X. Wang, C.W. Akey, Structure 18 (5) (2010) 571–583. [32] A. Saleh, S.M. Srinivasula, S. Acharya, R. Fishel, E.S. Alnemri, The Journal of Biolog- ical Chemistry 274 (25) (1999) 17941–17945. [33] C. Purring-Koch, G. McLendon, Proceedings of the National Academy of Sciences of the United States of America 97 (22) (2000) 11928–11931. [34] Y. Hu, L. Ding, D.M. Spencer, G. Nunez, The Journal of Biological Chemistry 273 (50) (1998) 33489–33494. [35] S.M. Srinivasula, M. Ahmad, T. Fernandes-Alnemri, E.S. Alnemri, Molecular Cell 1 (7) (1998) 949–957. [36] C. Purring, H. Zou, X. Wang, G. McLendon, Journal of the American Chemical Soci- ety 121 (32) (1999) 7435–7436. [37] S.J. Riedl, G.S. Salvesen, Nature Reviews. Molecular Cell Biology 8 (5) (2007) 405–413. [38] H.E. Kim, F. Du, M. Fang, X. Wang, Proceedings of the National Academy of Sci- ences of the United States of America 102 (49) (2005) 17545–17550. [39] Q. Bao, W. Lu, J.D. Rabinowitz, Y. Shi, Molecular Cell 25 (2) (2007) 181–192. [40] T.F. Reubold, S. Wohlgemuth, S. Eschenburg, The Journal of Biological Chemistry 284 (47) (2009) 32717–32724. [41] X. Jiang, X. Wang, The Journal of Biological Chemistry 275 (40) (2000) 31199–31203. [42] Y. Hu, M.A. Benedict, L. Ding, G. Nunez, The EMBO Journal 18 (13) (1999) 3586–3595. [43] H. Zou, Y. Li, X. Liu, X. Wang, The Journal of Biological Chemistry 274 (17) (1999) 11549–11556. [44] A.N. Lupas, J. Martin, Current Opinion in Structural Biology 12 (6) (2002) 746–753. [45] J. Snider, G. Thibault, W.A. Houry, Genome Biology 9 (4) (2008) 216. [46] D.A. Hattendorf, S.L. Lindquist, The EMBO Journal 21 (1–2) (2002) 12–21. [47] K. Karata, T. Inagawa, A.J. Wilkinson, T. Tatsuta, T. Ogura, The Journal of Biological Chemistry 274 (37) (1999) 26225–26232. [48] G.J. Steel, C. Harley, A. Boyd, A. Morgan, Molecular Biology of the Cell 11 (4) (2000) 1345–1356. [49] O. Danot, E. Marquenet, D. Vidal-Ingigliardi, E. Richet, Structure 17 (2) (2009) 172–182. 1424 T.F. Reubold, S. Eschenburg / Cellular Signalling 24 (2012) 1420–1425
  6. 6. [50] M. Proell, S.J. Riedl, J.H. Fritz, A.M. Rojas, R. Schwarzenbacher, PLoS One 3 (4) (2008) e2119. [51] N. Yan, J. Chai, E.S. Lee, L. Gu, Q. Liu, J. He, J.W. Wu, D. Kokel, H. Li, Q. Hao, D. Xue, Y. Shi, Nature 437 (7060) (2005) 831–837. [52] S. Qi, Y. Pang, Q. Hu, Q. Liu, H. Li, Y. Zhou, T. He, Q. Liang, Y. Liu, X. Yuan, G. Luo, J. Wang, N. Yan, Y. Shi, Cell 141 (3) (2010) 446–457. [53] E. Marquenet, E. Richet, Journal of Bacteriology 192 (19) (2010) 5181–5191. [54] G. van Ooijen, G. Mayr, M.M. Kasiem, M. Albrecht, B.J. Cornelissen, F.L. Takken, Journal of Experimental Botany 59 (6) (2008) 1383–1397. [55] D. Acehan, X. Jiang, D.G. Morgan, J.E. Heuser, X. Wang, C.W. Akey, Molecular Cell 9 (2) (2002) 423–432. [56] A.V. Diemand, A.N. Lupas, Journal of Structural Biology 156 (1) (2006) 230–243. [57] J. Li, J. Yuan, Oncogene 27 (48) (2008) 6194–6206. [58] P.K. Ho, C.J. Hawkins, The FEBS Journal 272 (21) (2005) 5436–5453. [59] C. Pop, G.S. Salvesen, The Journal of Biological Chemistry 284 (33) (2009) 21777–21781. [60] S.M. Srinivasula, R. Hegde, A. Saleh, P. Datta, E. Shiozaki, J. Chai, R.A. Lee, P.D. Robbins, T. Fernandes-Alnemri, Y. Shi, E.S. Alnemri, Nature 410 (6824) (2001) 112–116. [61] H.R. Stennicke, Q.L. Deveraux, E.W. Humke, J.C. Reed, V.M. Dixit, G.S. Salvesen, The Journal of Biological Chemistry 274 (13) (1999) 8359–8362. [62] K.M. Boatright, M. Renatus, F.L. Scott, S. Sperandio, H. Shin, I.M. Pedersen, J.E. Ricci, W.A. Edris, D.P. Sutherlin, D.R. Green, G.S. Salvesen, Molecular Cell 11 (2) (2003) 529–541. [63] C. Pop, J. Timmer, S. Sperandio, G.S. Salvesen, Molecular Cell 22 (2) (2006) 269–275. [64] Q. Bao, Y. Shi, Cell Death and Differentiation 14 (1) (2007) 56–65. [65] Y. Chao, E.N. Shiozaki, S.M. Srinivasula, D.J. Rigotti, R. Fairman, Y. Shi, PLoS Biology 3 (6) (2005) e183. [66] Y. Shi, Cell 117 (7) (2004) 855–858. [67] R.A. MacCorkle, K.W. Freeman, D.M. Spencer, Proceedings of the National Academy of Sciences of the United States of America 95 (7) (1998) 3655–3660. [68] M. Muzio, B.R. Stockwell, H.R. Stennicke, G.S. Salvesen, V.M. Dixit, The Journal of Bio- logical Chemistry 273 (5) (1998) 2926–2930. [69] X. Yang, H.Y. Chang, D. Baltimore, Molecular Cell 1 (2) (1998) 319–325. [70] M. Renatus, H.R. Stennicke, F.L. Scott, R.C. Liddington, G.S. Salvesen, Proceedings of the National Academy of Sciences of the United States of America 98 (25) (2001) 14250–14255. [71] G.S. Salvesen, S.J. Riedl, Advances in Experimental Medicine and Biology 615 (2008) 13–23. [72] S. Yuan, X. Yu, J.M. Asara, J.E. Heuser, S.J. Ludtke, C.W. Akey, Structure 19 (8) (2011) 1084–1096. [73] Q. Yin, H.H. Park, J.Y. Chung, S.C. Lin, Y.C. Lo, L.S. da Graca, X. Jiang, H. Wu, Molec- ular Cell 22 (2) (2006) 259–268. [74] L.A. Allan, N. Morrice, S. Brady, G. Magee, S. Pathak, P.R. Clarke, Nature Cell Biology 5 (7) (2003) 647–654. [75] L.A. Allan, P.R. Clarke, Molecular Cell 26 (2) (2007) 301–310. [76] A. Seifert, L.A. Allan, P.R. Clarke, The FEBS Journal 275 (24) (2008) 6268–6280. [77] H. Kanuka, K. Sawamoto, N. Inohara, K. Matsuno, H. Okano, M. Miura, Molecular Cell 4 (5) (1999) 757–769. [78] A. Rodriguez, H. Oliver, H. Zou, P. Chen, X. Wang, J.M. Abrams, Nature Cell Biology 1 (5) (1999) 272–279. [79] L. Zhou, Z. Song, J. Tittel, H. Steller, Molecular Cell 4 (5) (1999) 745–755. [80] S. Yuan, X. Yu, M. Topf, L. Dorstyn, S. Kumar, S.J. Ludtke, C.W. Akey, Structure 19 (1) (2011) 128–140. [81] A. Oberst, C. Bender, D.R. Green, Cell Death and Differentiation 15 (7) (2008) 1139–1146. [82] R.J. Krieser, K. White, Apoptosis 14 (8) (2009) 961–968. [83] E. Arama, J. Agapite, H. Steller, Developmental Cell 4 (5) (2003) 687–697. [84] E. Arama, M. Bader, M. Srivastava, A. Bergmann, H. Steller, The EMBO Journal 25 (1) (2006) 232–243. [85] C.S. Mendes, E. Arama, S. Brown, H. Scherr, M. Srivastava, A. Bergmann, H. Steller, B. Mollereau, EMBO Reports 7 (9) (2006) 933–939. [86] E. Abdelwahid, T. Yokokura, R.J. Krieser, S. Balasundaram, W.H. Fowle, K. White, Developmental Cell 12 (5) (2007) 793–806. [87] L. Dorstyn, K. Mills, Y. Lazebnik, S. Kumar, The Journal of Cell Biology 167 (3) (2004) 405–410. [88] L. Dorstyn, S. Read, D. Cakouros, J.R. Huh, B.A. Hay, S. Kumar, The Journal of Cell Biology 156 (6) (2002) 1089–1098. [89] J.C. Means, I. Muro, R.J. Clem, Cell Death and Differentiation 13 (7) (2006) 1222–1234. [90] J. Varkey, P. Chen, R. Jemmerson, J.M. Abrams, The Journal of Cell Biology 144 (4) (1999) 701–710. [91] K.C. Zimmermann, J.E. Ricci, N.M. Droin, D.R. Green, The Journal of Cell Biology 156 (6) (2002) 1077–1087. [92] L. Dorstyn, S. Kumar, Cell Death and Differentiation 15 (3) (2008) 461–470. [93] X. Yu, L. Wang, D. Acehan, X. Wang, C.W. Akey, Journal of Molecular Biology 355 (3) (2006) 577–589. [94] N. Yan, J.R. Huh, V. Schirf, B. Demeler, B.A. Hay, Y. Shi, The Journal of Biological Chemistry 281 (13) (2006) 8667–8674. [95] R. Wilson, L. Goyal, M. Ditzel, A. Zachariou, D.A. Baker, J. Agapite, H. Steller, P. Meier, Nature Cell Biology 4 (6) (2002) 445–450. [96] H.D. Ryoo, A. Bergmann, H. Gonen, A. Ciechanover, H. Steller, Nature Cell Biology 4 (6) (2002) 432–438. [97] M.S. Spector, S. Desnoyers, D.J. Hoeppner, M.O. Hengartner, Nature 385 (6617) (1997) 653–656. [98] N. Yan, L. Gu, D. Kokel, J. Chai, W. Li, A. Han, L. Chen, D. Xue, Y. Shi, Molecular Cell 15 (6) (2004) 999–1006. 1425T.F. Reubold, S. Eschenburg / Cellular Signalling 24 (2012) 1420–1425

A molecular view on signal transduction by the apoptosome

Views

Total views

205

On Slideshare

0

From embeds

0

Number of embeds

4

Actions

Downloads

3

Shares

0

Comments

0

Likes

0

×