BH 3‐in‐groove dimerization initiates and helix 9 dimerization expands Bax pore assembly in membranes
2015; Springer Nature; Volume: 35; Issue: 2 Linguagem: Inglês
10.15252/embj.201591552
ISSN1460-2075
AutoresZhi Zhang, Sabareesh Subramaniam, Justin Kale, Chenyi Liao, Bo Huang, Hetal Brahmbhatt, Samson G.F. Condon, Suzanne M. Lapolla, Franklin A. Hays, Jingzhen Ding, Feng He, Xuejun C. Zhang, Jianing Li, Alessandro Senes, David W. Andrews, Jialing Lin,
Tópico(s)Mitochondrial Function and Pathology
ResumoArticle23 December 2015free access Source Data BH3-in-groove dimerization initiates and helix 9 dimerization expands Bax pore assembly in membranes Zhi Zhang Zhi Zhang Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Search for more papers by this author Sabareesh Subramaniam Sabareesh Subramaniam Department of Biochemistry, University of Wisconsin-Madison, Madison, WI, USA Search for more papers by this author Justin Kale Justin Kale Biological Sciences, Sunnybrook Research Institute, Department of Biochemistry, University of Toronto, Toronto, ON, Canada Search for more papers by this author Chenyi Liao Chenyi Liao Department of Chemistry, University of Vermont, Burlington, VT, USA Search for more papers by this author Bo Huang Bo Huang Institute of Biophysics, Chinese Academy of Sciences, Beijing, China Search for more papers by this author Hetal Brahmbhatt Hetal Brahmbhatt Biological Sciences, Sunnybrook Research Institute, Department of Biochemistry, University of Toronto, Toronto, ON, Canada Search for more papers by this author Samson GF Condon Samson GF Condon Department of Biochemistry, University of Wisconsin-Madison, Madison, WI, USA Search for more papers by this author Suzanne M Lapolla Suzanne M Lapolla Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Search for more papers by this author Franklin A Hays Franklin A Hays Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Peggy and Charles Stephenson Cancer Center, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Search for more papers by this author Jingzhen Ding Jingzhen Ding Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Search for more papers by this author Feng He Feng He Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Search for more papers by this author Xuejun C Zhang Xuejun C Zhang Institute of Biophysics, Chinese Academy of Sciences, Beijing, China Search for more papers by this author Jianing Li Jianing Li Department of Chemistry, University of Vermont, Burlington, VT, USA Search for more papers by this author Alessandro Senes Alessandro Senes Department of Biochemistry, University of Wisconsin-Madison, Madison, WI, USA Search for more papers by this author David W Andrews David W Andrews Biological Sciences, Sunnybrook Research Institute, Department of Biochemistry, University of Toronto, Toronto, ON, Canada Search for more papers by this author Jialing Lin Corresponding Author Jialing Lin Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Peggy and Charles Stephenson Cancer Center, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Search for more papers by this author Zhi Zhang Zhi Zhang Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Search for more papers by this author Sabareesh Subramaniam Sabareesh Subramaniam Department of Biochemistry, University of Wisconsin-Madison, Madison, WI, USA Search for more papers by this author Justin Kale Justin Kale Biological Sciences, Sunnybrook Research Institute, Department of Biochemistry, University of Toronto, Toronto, ON, Canada Search for more papers by this author Chenyi Liao Chenyi Liao Department of Chemistry, University of Vermont, Burlington, VT, USA Search for more papers by this author Bo Huang Bo Huang Institute of Biophysics, Chinese Academy of Sciences, Beijing, China Search for more papers by this author Hetal Brahmbhatt Hetal Brahmbhatt Biological Sciences, Sunnybrook Research Institute, Department of Biochemistry, University of Toronto, Toronto, ON, Canada Search for more papers by this author Samson GF Condon Samson GF Condon Department of Biochemistry, University of Wisconsin-Madison, Madison, WI, USA Search for more papers by this author Suzanne M Lapolla Suzanne M Lapolla Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Search for more papers by this author Franklin A Hays Franklin A Hays Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Peggy and Charles Stephenson Cancer Center, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Search for more papers by this author Jingzhen Ding Jingzhen Ding Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Search for more papers by this author Feng He Feng He Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Search for more papers by this author Xuejun C Zhang Xuejun C Zhang Institute of Biophysics, Chinese Academy of Sciences, Beijing, China Search for more papers by this author Jianing Li Jianing Li Department of Chemistry, University of Vermont, Burlington, VT, USA Search for more papers by this author Alessandro Senes Alessandro Senes Department of Biochemistry, University of Wisconsin-Madison, Madison, WI, USA Search for more papers by this author David W Andrews David W Andrews Biological Sciences, Sunnybrook Research Institute, Department of Biochemistry, University of Toronto, Toronto, ON, Canada Search for more papers by this author Jialing Lin Corresponding Author Jialing Lin Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Peggy and Charles Stephenson Cancer Center, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA Search for more papers by this author Author Information Zhi Zhang1, Sabareesh Subramaniam2,‡, Justin Kale3,‡, Chenyi Liao4,‡, Bo Huang5,‡, Hetal Brahmbhatt3, Samson GF Condon2, Suzanne M Lapolla1, Franklin A Hays1,6, Jingzhen Ding1, Feng He1, Xuejun C Zhang5, Jianing Li4, Alessandro Senes2, David W Andrews3 and Jialing Lin 1,6 1Department of Biochemistry and Molecular Biology, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA 2Department of Biochemistry, University of Wisconsin-Madison, Madison, WI, USA 3Biological Sciences, Sunnybrook Research Institute, Department of Biochemistry, University of Toronto, Toronto, ON, Canada 4Department of Chemistry, University of Vermont, Burlington, VT, USA 5Institute of Biophysics, Chinese Academy of Sciences, Beijing, China 6Peggy and Charles Stephenson Cancer Center, University of Oklahoma Health Sciences Center, Oklahoma City, OK, USA ‡These authors contributed equally to this work *Corresponding author. Tel: +1 405 271 2227, Ext. 61216; E-mail: [email protected] The EMBO Journal (2016)35:208-236https://doi.org/10.15252/embj.201591552 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract Pro-apoptotic Bax induces mitochondrial outer membrane permeabilization (MOMP) by forming oligomers through a largely undefined process. Using site-specific disulfide crosslinking, compartment-specific chemical labeling, and mutational analysis, we found that activated integral membrane Bax proteins form a BH3-in-groove dimer interface on the MOM surface similar to that observed in crystals. However, after the α5 helix was released into the MOM, the remaining interface with α2, α3, and α4 helices was rearranged. Another dimer interface was formed inside the MOM by two intersected or parallel α9 helices. Combinations of these interfaces generated oligomers in the MOM. Oligomerization was initiated by BH3-in-groove dimerization, without which neither the other dimerizations nor MOMP occurred. In contrast, α9 dimerization occurred downstream and was required for release of large but not small proteins from mitochondria. Moreover, the release of large proteins was facilitated by α9 insertion into the MOM and localization to the pore rim. Therefore, the BH3-in-groove dimerization on the MOM nucleates the assembly of an oligomeric Bax pore that is enlarged by α9 dimerization at the rim. Synopsis Apoptotic Bax oligomers permeabilize the mitochondrial outer membrane. Structural analyses and modeling of Bax interactions at mitochondria show that BH3-in-groove dimerization on membranes initiates the pore assembly, which is followed by helix 9 dimerization-mediated expansion. Bax protein oligomerization initiates with helices α2-α5 forming a BH3-in-groove dimer interface on the membrane surface. The BH3-in-groove dimer interface is rearranged after α5 insertion into the membrane. α9 helices from neighboring Bax molecules form another dimer interface inside the membrane linking the BH3-in-groove dimers to higher order oligomers. α9 insertion and dimerization facilitate pore enlargement required to release large mitochondrial intermembrane space proteins. Introduction In most cells, Bax is a monomeric protein in the cytosol. During apoptosis initiation, Bax is activated and targeted to the mitochondrial outer membrane (MOM). The active Bax undergoes a series of conformational changes and eventually forms oligomeric pores in the MOM that release cytochrome c and other mitochondrial intermembrane space proteins to activate caspases and nucleases, thereby dismantling the cells (Borner & Andrews, 2014; Chi et al, 2014; Moldoveanu et al, 2014; Volkmann et al, 2014; Westphal et al, 2014b). Previous studies revealed a multi-step process that transforms Bax from a dormant soluble protein to an active integral membrane protein. Bax cycles on and off membranes by a process called retrotranslocation (Edlich et al, 2011; Schellenberg et al, 2013). The Bax is activated and bound to mitochondria by BH3-only proteins such as tBid (Lovell et al, 2008) or by previously activated Bax (Tan et al, 2006). The activated Bax partially embeds helices α5, α6, and α9 into the MOM as cysteines positioned in these helices become inaccessible to a membrane-impermeant sulfhydryl-specific modifying agent when Bax inserts into membranes (Annis et al, 2005; Westphal et al, 2014a). Inaccessibility was observed for Bax bound to the mitochondria isolated from etoposide-treated Myc-null cells where Bax did not oligomerize, and from Myc-expressing cells where Bax did oligomerize, suggesting that the helices are inserted into the MOM before oligomerization (Annis et al, 2005). Consistent with this model, residue inaccessibility was also observed before the tBid-induced oligomerization of Bax mutants that were constitutively bound to mitochondria due to a mutation in α9 (Westphal et al, 2014a). The α5 insertion was also supported by an increase of fluorescence of an environment-sensing fluorophore attached to the α5 of Bax after it was activated by membrane-bound tBid (Lovell et al, 2008). Kinetic analysis of the fluorescence changes associated with α5 insertion and Bax oligomerization indicated that the insertion occurred earlier than the oligomerization. However, while the early chemical labeling study concluded that the α5, α6, and α9 are deeply inserted into the lipid bilayer, the later study concluded that the α5 and α6 are shallowly inserted into the cytosolic leaflet of the bilayer with some α5 residues buried in the cytosolic domain of Bax oligomer after Bax is activated by tBid. Therefore, a more rigorous topology survey is required to differentiate the models for the three membrane-embedded helices, and to ascertain the topology of the other regions, particularly the BH3 region and its binding groove, which are critical to Bax interaction and function (Bleicken et al, 2010; Zhang et al, 2010; Dewson et al, 2012). The structure of the oligomeric Bax pore was largely unexplored until recently. Our photocrosslinking study revealed two interdependent interfaces in the Bax oligomer formed in detergent micelles (Zhang et al, 2010). A double electron–electron resonance (DEER) study of Bax oligomer formed in detergent micelle and liposomal membrane suggested an antiparallel helical dimer interface formed by α2-α3 region of neighboring Bax molecules in the oligomer (Bleicken et al, 2010). A disulfide-crosslinking study indicated that the antiparallel α2-α3 interface was extended to include α4 which binds to the other side of α2, and this interface together with a parallel α6 interface could generate Bax oligomers in the MOM (Dewson et al, 2012). Another disulfide-crosslinking study detected an α9 interface formed by a Bax mutant constitutively bound to mitochondria (Iyer et al, 2015). Whether wild-type Bax can form this α9 interface was unknown. Moreover, the contribution of all of these interfaces to the oligomeric pore assembly has not been assessed. A crystallographic study revealed structures of three Bax complexes (Czabotar et al, 2013). The first is a domain-swapped dimer in which two Bax polypeptides lacking the C-terminal α9 helix (BaxΔα9) swap their α6-α8 helices, resulting in two globular units, each comprising α1-α5 helices from one monomer (the core domain), plus α6-α8 helices from the other monomer (the latch domain), that are bridged by two extended antiparallel α5-α6 helices. The second is the domain-swapped BaxΔα9 dimer with a BH3 peptide of tBid bound to each globular unit via a hydrophobic groove that is occupied by α9 in full-length Bax monomer (Suzuki et al, 2000). The third was formed by a GFP fusion protein containing Bax α2-α5 helices. A symmetric dimer interface exists in this complex, in which the BH3 region or α2 of one Bax fragment engages a groove in the other Bax fragment and vice versa, resulting in two reciprocal BH3-in-groove interfaces (Fig 1A). Figure 1. Activated Bax proteins integrate into the MOM and dimerize via a BH3-in-groove interface The crystal structure of the BH3-in-groove Bax homodimer (PDB entry 4BDU) is shown with one monomer colored gray and the other colored green, magenta, blue, and red for its α2, α3, α4, and α5 helices, respectively, as indicated. The residue pairs that were replaced with cysteine pairs in (B) are presented in stick form, and their β-carbon atoms linked by dashed lines with the distances ranging from 5.0 to 6.0 Å. The in vitro synthesized [35S]Met-labeled single-cysteine Bax proteins were activated and targeted to the mitochondria that were pretreated with NEM to block the sulfhydryls of mitochondrial proteins. The resulting mitochondria were isolated and oxidized by CuPhe for 30 min. NEM and EDTA were then added to stop the oxidation. For the "0 min" controls, NEM and EDTA were added before the addition of CuPhe. The resulting samples were analyzed by phosphorimaging after non-reducing or reducing SDS–PAGE (see Appendix Fig S2A). Oxidized mitochondria with the radioactive single-cysteine Bax protein pair or double-cysteine Bax protein were prepared and analyzed as in (B). Oxidized mitochondria with the radioactive single-cysteine Bax protein pair or double-cysteine Bax protein were prepared as in (B). After an aliquot was withdrawn as input, another aliquot was extracted by Na2CO3 (pH 11.5) and centrifuged through a sucrose cushion to separate the integral proteins in the membrane pellet from the soluble and peripheral proteins in the supernatant. The input, pellet, and supernatant were analyzed by non-reducing SDS–PAGE and phosphorimaging. In a parallel control experiment, the pellet and supernatant were analyzed by reducing SDS–PAGE and immunoblotting with an antibody specific to PDHE1α, a soluble mitochondrial matrix protein. Data information: In (B–D), protein standards are indicated on the side of phosphor images or immunoblots by their molecular mass (Mr). Open circles indicate Bax monomers. Upward arrows indicate disulfide-linked dimers of the same single-cysteine Bax mutant (e.g., M79C), and downward arrows disulfide-linked dimers of two different single-cysteine Bax mutants (e.g., L59C + M79C) or of the same double-cysteine Bax mutant (L59C,M79C). Closed circles indicate the disulfide-linked heterodimer of Bax M79C and the BH3 peptide (with a cysteine at position 62). The number of independent replicates done for each Bax mutant in (B–C), n = 2 for C62+M79C, T56C, and R94C; 3 for L63C, L63C+A112C, A112C, K64C, K64C+M99C, and M99C; 4 for L59C+M79C, M79C, C62, C62+L76C, and T56C+R94C; 6 for L59C, and L76C; and 8 for L59C,M79C. For each Bax mutant and PDHE1α in (D), n = 2. Source data are available online for this figure. Source Data for Figure 1 [embj201591552-sup-0006-SDataFig1.pdf] Download figure Download PowerPoint Based on these crystal structures, the following model was proposed for Bax activation and oligomerization (Czabotar et al, 2013). Binding of a BH3-only protein to the hydrophobic groove of Bax after α9 is released from the groove and inserted into the MOM triggers the release of the latch domain from the core domain and the exposure of the BH3 region. The exposed BH3 region of one Bax replaces the BH3-only protein from the groove of other Bax and vice versa, resulting in a Bax homodimer with the BH3-in-groove interface that nucleates the oligomerization process. In addition, a hydrophobic patch consisting of aromatic residues from α4 to α5 that is located on one side of the BH3-in-groove dimer engages the MOM to promote MOMP (Fig 2A, top-left). However, these inferences were made based on the structure of a domain-swapped dimer that is acknowledged to be an off mechanism dead-end complex and the structures of Bax deletion mutants, some of them fused with GFP and all of them formed without membranes. Although the core–latch separation and the BH3-in-groove dimerization were confirmed recently by a DEER study that measured the intra- and intermolecular distances of tBid-activated liposome-bound Bax molecules with spin-labeled cysteines (Bleicken et al, 2014), the broad distance distributions implied that the Bax oligomer structure might be dynamic and flexible. Moreover, only three intramolecular distances were obtained from mitochondrion-bound Bax. Thus, it was uncertain whether the domain separation and dimerization actually occurred for the Bax at mitochondria. Figure 2. The BH3 region and the groove are partially embedded in the MOM A. Top-left panel, structure of the BH3-in-groove dimer (PDB entry 4BDU) with the predicted MOM-engaging residues presented in stick form. Bottom-left panel, structure of the BH3-in-groove dimer with the predicted protein-buried residues presented in stick form. Top-right panel, structure of the BH3-in-groove dimer without helix α5 (modified from PDB entry 4BDU), and bottom-right panel, structure of α5 (extracted from the NMR structure of Bax monomer, PDB entry 1F16). In both structures in the right panels, the residues that are buried in the MOM or exposed to the aqueous milieu as the IASD-labeling data in (B–D) confirmed are presented in yellow or cyan stick form, respectively. In all panels, the color codes for the two monomers are the same as that in Fig 1A, and indicated. B. The in vitro synthesized radioactive Bax proteins, each with a single cysteine positioned in helix α2, α3, α4, or α5, were activated and targeted to the mitochondria. The resulting mitochondria were isolated and treated with IASD in the absence or presence of CHAPS, urea, or both. After 30 min, the labeling reactions were stopped by mercaptoethanol. For the "0 min" controls, the samples were pretreated with mercaptoethanol before addition of IASD. The IASD-labeled radioactive Bax proteins were resolved from the unlabeled ones using either isoelectric focusing (IEF; as indicated) or gradient SDS–PAGE and detected by phosphorimaging. Triangles and arrows indicate the unlabeled and IASD-labeled Bax proteins, respectively. n = 3 for V111C, and K119C; 4 for W107C, and A117C; 2 for other mutants. C, D. The phosphorimaging data for IASD labeling of Bax mutants in (B) and the similar data from the independent replicates were quantified to derive the membrane or protein burial indices as described and shown in Appendix Fig S3. The membrane burial indices were normalized by that of G179C in α9, a reference for the membrane-buried residues. The resulting relative membrane burial indices are shown in (C). The residues with the relative membrane burial index ≥40% of that of Gly179 are considered as buried in the MOM. Similarly, the relative protein burial indices shown in (D) were obtained by using Y115C in α5 as a reference for the protein-buried residues. The residues with the relative protein burial index ≥40% of that of Tyr115 are considered as buried in the protein or its complex. Source data are available online for this figure. Source Data for Figure 2 [embj201591552-sup-0007-SDataFig2.pdf] Download figure Download PowerPoint Förster resonance energy transfer (FRET) was used to measure intramolecular distances within a Bax molecule and intermolecular distances between two Bax molecules in the cytosol or the mitochondria of live and apoptotic cells (Gahl et al, 2014). The intramolecular distance from the donor dye-labeled α2 to the acceptor dye-labeled α7 was increased after Bax translocation from the cytosol to the mitochondria, consistent with the core–latch separation model. However, a significant decrease in FRET efficiency between the donor at α2 and the acceptor at α5 suggested a large conformational change within the core that separates the two helices further apart, inconsistent with the subtle distance changes between the two sites during a transition from the monomer NMR structure to the domain-swapped and the BH3-in-groove dimer crystal structures (Suzuki et al, 2000; Czabotar et al, 2013). In addition, the FRET distance between α9 and α2 indicated that α9 was distant from the canonical groove in the cytosolic Bax, thereby exposing the groove and BH3 region for potential homo- and hetero-interactions. After migrating to the mitochondria, while the intramolecular distance between α9 and α2 remained large and in accordance with the proposed core–latch separation, the intermolecular distance measured by the FRET between a donor-labeled α9 in one Bax molecule and an acceptor-labeled α9 in other Bax molecule was comparable with a homodimerization between the two α9 helices, in line with a model proposed in the DEER and crosslinking studies (Bleicken et al, 2014; Iyer et al, 2015). However, the DEER study proposed an antiparallel α9 dimer model, different from the parallel dimer model proposed by the other studies. Furthermore, intermolecular FRET measurements supported a Bax oligomer model in which in addition to the α9 dimer interface, α2 and α3 form the other dimer interface that unlike the crystallographic BH3-in-groove interface does not involve α5. To further refine the mechanism of Bax activation and oligomerization, several important questions must be addressed. Does reciprocal binding of the BH3 region of one Bax to the groove of other Bax indeed occur at the MOM resulting in a BH3-in-groove dimer interface as revealed by crystallography? Does formation of the BH3-in-groove interface nucleate Bax oligomerization? For an oligomer to form, there must be additional dimer interfaces, but what regions of Bax are involved? In particular, does α9 form an additional interface, which together with the BH3-in-groove interface mediates Bax oligomerization? Are these dimer interfaces located above, on or in the MOM? If the BH3-in-groove dimer interface is located on the membrane surface with the hydrophobic patch engaging the membrane as the crystallography study proposed and the DEER and one chemical labeling study concluded (Czabotar et al, 2013; Bleicken et al, 2014; Westphal et al, 2014a), could that drive the hydrophobic patch, particularly the hydrophobic α5 helix, more deeply into the membrane as the other chemical labeling study suggested (Annis et al, 2005)? If the α5 inserts into the membrane and thereby separates from the rest of the BH3-in-groove dimer as the FRET study indicated (Gahl et al, 2014), would the remnant interface rearrange to another conformation that is more stable? In other words, does the BH3-in-groove dimer represent a transient intermediate state? Most importantly, how do Bax oligomers form pores in the MOM? And how flexible is the pore structure? Accumulating evidence suggests that the pores are proteolipidic with some regions of Bax embedded into one leaflet of the bilayer to increase the membrane tension to a point that the bilayer would fuse to a highly curved monolayer, resulting in a toroidal pore with polar and charged lipid head groups lining the rim (Garcia-Saez, 2012). Other regions of Bax might localize to the rim to decrease the line tension, thereby stabilizing the lipidic pore. However, what are the Bax regions that induce, and that stabilize the lipidic pore? Finally, the size of Bax pore is tunable (Bleicken et al, 2013), but does higher order oligomerization expand the pore? To answer these questions, we used disulfide crosslinking to map the dimer interfaces in the active mitochondrial Bax oligomer, and compartment-specific chemical labeling to determine the membrane topology. We built structural models and conducted molecular dynamics (MD) simulations to fit the experimental data, and generated mutations to test the functional relevance of the models. Based on these data and models, we propose a molecular scheme for how active Bax proteins are assembled into oligomers to induce and expand lipidic pores in the MOM. Results Bax mutants for interface and topology mapping are functional Bax mutants with single, double, or triple cysteines located at specific positions were generated from a Bax cysteine-null mutant (Fig EV1A). The mutant proteins were synthesized by using a coupled in vitro transcription and translation (TNT) system, and their tBid-dependent MOMP activity was measured in an in vitro cytochrome c release assay (Ding et al, 2014) using Bax and Bak double deficient mitochondria (Bax−/−/Bak−/− mitochondria) (Billen et al, 2008). Addition of both tBid protein and the wild-type (WT) Bax protein-producing TNT mixture resulted in a synergistic increase in cytochrome c released above the protein-independent background release (Fig EV1B, compare the open bars from "+Bax WT, +tBid" and "+Vector" samples). The releases above the background, indicated as the "corrected" releases (hatched bars) calculated from the "raw" releases (open bars) as described in the legend, showed the protein-dependent releases of cytochrome c by tBid (~20%), Bax (~0%), and both (~50%). With the background release common to these samples subtracted, the "corrected" data in Fig EV1C demonstrated that the cytochrome c release due to Bax alone is ~10% or lower for the wild-type Bax and all the mutants, except for L76C and V110C. These two mutants are "autoactive" as they released ~30–60% of cytochrome c in the absence of tBid. As expected, addition of tBid to the "non-autoactive" Bax mutants increased the release to ~30–60%, comparable to the wild-type Bax. Even though some of the mitochondria are somewhat leaky in our in vitro lysate-based system (Fig EV1B, "Mito-only" sample), the intact mitochondria still respond to the tBid and Bax proteins appropriately. Click here to expand this figure. Figure EV1. Sequence and in vitro MOMP activity of Bax mutants Bax sequence is shown with BH motifs highlighted by dashed lines above and helices identified by arrows below. The native cysteines (closed circle-indicated Cs) were changed to alanine to create the cysteine-null mutant. Single-cysteine Bax mutants were created from the cysteine-null mutant by individually replacing the red-colored residues with cysteine. Triangles indicate Gly108, Gly179, Thr182, and Ser184 of Bax that were changed to glutamate, isoleucine, isoleucine, and valine in the G108E, G179I, T182I, and S184V mutants, respectively. The GxxxA (GVLTA) motif in α9 is in bold font. The cytochrome c release from the Bax−/−/Bak−/− mitochondria-only sample (Mito only), and from the mitochondrial plus purified recombinant tBid protein (+tBid), the TNT reaction programmed by pSPUTK vector (+Vector) or wild-type Bax gene inserted after SP6 promoter in the vector (+Bax WT), or their combinations was measured in n independent replicates as indicated. The raw data shown as open bars are the means with the standard deviations (s.d.). A background release ˜20% was observed in the mitochondria-only sample, which might be due to the mitochondria that were frozen and thawed once before used in the assay according to an established protocol (Yamaguchi et al, 2007). Addition of bacterial expressed and purified tBid protein increased the release slightly. The TNT mixture containing the vector plasmid, or the WT Bax plasmid that produced the WT Bax protein (see Appendix Fig S1) also increased the release slightly. While addition of both tBid protein and the vector-containing TNT mixture showed a marginally additive release, addition of both tBid protein and the WT Bax protein-producing TNT mixture resulted in a synergistic increase in cytochrome c release. After the "raw" release of "+Vector" control was subtracted from the "raw" releases of "+Vector, +tBid", "+Bax WT", and "+Bax WT, +tBid" samples, which all contained the vector (with or without the WT Bax
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