Proteolipid domains form in biomimetic and cardiac mitochondrial vesicles and are regulated by cardiolipin concentration but not monolyso-cardiolipin

Cardiolipin (CL) is an anionic phospholipid mainly located in the inner mitochondrial membrane, where it helps regulate bioenergetics, membrane structure, and apoptosis. Localized, phase-segregated domains of CL are hypothesized to control mitochondrial inner membrane organization. However, the existence and underlying mechanisms regulating these mitochondrial domains are unclear. Here, we first isolated detergent-resistant cardiac mitochondrial membranes that have been reported to be CL-enriched domains. Experiments with different detergents yielded only nonspecific solubilization of mitochondrial phospholipids, suggesting that CL domains are not recoverable with detergents. Next, domain formation was investigated in biomimetic giant unilamellar vesicles (GUVs) and newly synthesized giant mitochondrial vesicles (GMVs) from mouse hearts. Confocal fluorescent imaging revealed that introduction of cytochrome c into membranes promotes macroscopic proteolipid domain formation associated with membrane morphological changes in both GUVs and GMVs. Domain organization was also investigated after lowering tetralinoleoyl-CL concentration and substitution with monolyso-CL, two common modifications observed in cardiac pathologies. Loss of tetralinoleoyl-CL decreased proteolipid domain formation in GUVs, because of a favorable Gibbs-free energy of lipid mixing, whereas addition of monolyso-CL had no effect on lipid mixing. Moreover, murine GMVs generated from cardiac acyl-CoA synthetase-1 knockouts, which have remodeled CL acyl chains, did not perturb proteolipid domains. Finally, lowering the tetralinoleoyl-CL content had a stronger influence on the oxidation status of cytochrome c than did incorporation of monolyso-CL. These results indicate that proteolipid domain formation in the cardiac mitochondrial inner membrane depends on tetralinoleoyl-CL concentration, driven by underlying lipid-mixing properties, but not the presence of monolyso-CL.

Cardiolipin (CL) is an anionic phospholipid mainly located in the inner mitochondrial membrane, where it helps regulate bioenergetics, membrane structure, and apoptosis. Localized, phase-segregated domains of CL are hypothesized to control mitochondrial inner membrane organization. However, the existence and underlying mechanisms regulating these mitochondrial domains are unclear. Here, we first isolated detergent-resistant cardiac mitochondrial membranes that have been reported to be CL-enriched domains. Experiments with different detergents yielded only nonspecific solubilization of mitochondrial phospholipids, suggesting that CL domains are not recoverable with detergents. Next, domain formation was investigated in biomimetic giant unilamellar vesicles (GUVs) and newly synthesized giant mitochondrial vesicles (GMVs) from mouse hearts. Confocal fluorescent imaging revealed that introduction of cytochrome c into membranes promotes macroscopic proteolipid domain formation associated with membrane morphological changes in both GUVs and GMVs. Domain organization was also investigated after lowering tetralinoleoyl-CL concentration and substitution with monolyso-CL, two common modifications observed in cardiac pathologies. Loss of tetralinoleoyl-CL decreased proteolipid domain formation in GUVs, because of a favorable Gibbs-free energy of lipid mixing, whereas addition of monolyso-CL had no effect on lipid mixing. Moreover, murine GMVs generated from cardiac acyl-CoA synthetase-1 knockouts, which have remodeled CL acyl chains, did not perturb proteolipid domains. Finally, lowering the tetralinoleoyl-CL content had a stronger influence on the oxidation status of cytochrome c than did incorporation of monolyso-CL. These results indicate that proteolipid domain formation in the cardiac mitochondrial inner mem-brane depends on tetralinoleoyl-CL concentration, driven by underlying lipid-mixing properties, but not the presence of monolyso-CL.
Cardiolipin (CL) 2 is a unique anionic phospholipid that is localized to the inner mitochondrial membrane, where it plays a role in regulating bioenergetics, membrane structure, and apoptosis (1)(2)(3)(4)(5)(6). Several mechanisms are proposed to explain how CL regulates inner mitochondrial membrane structure-function. These mechanisms include CL directly binding select enzymes, anchoring cytochrome c, promoting correct cristae morphology, and enhancing supercomplex formation (7)(8)(9)(10)(11)(12)(13). One poorly studied hypothesis is that CL facilitates the formation of distinct phase-segregated domains (14,15). The biological advantage of CL-enriched domains would be to concentrate respiratory protein complexes for enhanced electron channeling, to augment membrane curvature, and to serve as docking platforms for apoptotic signals (16 -22).
The limited understanding on the existence of CL-enriched domains comes largely from two-or three-component model membrane studies. Calorimetric studies with binary phospholipid mixtures in the presence of cytochrome c, truncated Bid, or creatine kinase suggested segregation of CL into distinct clusters (18). Studies with Brewster angle microscopy of phosphatidylcholine/phosphatidylethanolamine monolayers containing creatine kinase have also provided direct support for CL domains (23). Furthermore, microscopy studies with biomimetic CL-containing mixtures have demonstrated the unique This work was supported by National Institutes of Health Grant R01HL123647 (to D. B. and S. R. S.) and internal funds from the University of North Carolina, Chapel Hill (to S. R. S.). The authors declare that they have no conflicts of interest with the contents of this article. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. This article contains Figs. S1-S3. 1 To whom correspondence should be addressed. Tel.: 919-843-4348; E-mail: shaikhsa@email.unc.edu.
ability of cytochrome c to regulate membrane morphology and induce CL domain formation (24,25). CL domains have also been identified biochemically with detergent-extraction methods; similar to approaches used for the study of detergent-resistant "lipid raft" membranes (17). There are several major gaps in knowledge regarding CLenriched domain organization. The first major limitation is that many studies have not effectively modeled the inner mitochondrial composition and even relied on lipid raft models (24,26). Approximately 70% of the inner mitochondrial membrane is accounted for by phosphatidylcholine (40%) and phosphatidylethanolamine (30%), whereas CL makes up about 15-20% of total phospholipids (7). Phosphatidylinositol, phosphatidic acid, and phosphatidylserine account for the remaining phospholipids, with very low levels of cholesterol. Thus, CL domain formation should be studied in the context of inner mitochondrial-mimicking membranes. Second, the underlying mechanistic factors that regulate CL domain formation are not defined. These are essential to establish because in a range of diseases, CL is subject to considerable physiochemical modifications that impair mitochondrial inner membrane structurefunction (3,(27)(28)(29). For instance, with heart failure and type II diabetes, cardiac CL acyl chains undergo extensive remodeling that could disrupt CL domain formation (30 -37). In Barth syndrome, there is a significant loss of cardiac tetralinoleoyl-CL and increased levels of monolyso-CL (MLCL) (27, 38 -42).
The first objective of this study was to determine the utility of biochemical detergent extraction to isolate CL domains. Subsequently, we investigated if CL domains can form in the absence or presence of select proteins. These proteins included cytochrome c, creatine kinase, and ␣-synuclein, which are all associated with mitochondrial membranes. The approach relied on quantitative imaging of biomimetic giant unilamellar vesicles (GUVs) modeling the cardiac inner mitochondrial membrane, and the use of giant mitochondrial vesicles (GMVs) constructed for the first time with phospholipids extracted from rodent cardiac mitochondria. Next, we determined if decreased CL levels, or the accumulation of MLCL, as observed in various diseases, disrupted the organization of proteolipid domains. These studies relied on biomimetic mitochondrial GUVs as well as lipid monolayers mimicking the inner mitochondrial membrane lipid composition to quantify lipid-lipid mixing. In parallel, we constructed GMVs using cardiac tissue harvested from control and acyl-CoA synthetase 1 knockout mice to determine how CL remodeling impacted the organization of native phospholipids and thereby proteolipid domains.

Mitochondrial CL microdomains cannot be isolated with detergent-extraction methods
We first determined if different detergents were equally efficacious at extracting CL-enriched microdomains from rat cardiac mitochondria. Rat myocardial tissue was utilized to ensure a high abundance of material for microdomain isolation. We relied on Triton X-100, Nonidet P-40, and digitonin at various concentrations (0.01, 0.05, and 0.5% w/v). Increasing the concentration of all three detergents led to an overall decrease of phospholipid levels, as measured by TLC, in the insoluble detergent-resistant membrane (DRM) (I) fractions (Fig. 1, A-C). This decrease was paralleled by an increase in the amount of lipids in the detergent-soluble membrane (DSM (S)) fractions ( Fig. 1, A-C). The increase in the concentration of Triton X-100 from 0.01 to 0.05% caused a larger reduction in the amount of phospholipids present in the DRM fraction than Nonidet P-40 or digitonin (Fig. 1, A-C). At a concentration of 0.5%, phospholipids in the DRM fraction were appreciably reduced with all three detergents (Fig. 1, A-C).
Both DSM (S) and DRM (I) fractions were assayed for protein content. Triton X-100 and Nonidet P-40 had a higher concentration of protein in their DSM fractions than their DRM fractions, at all detergent concentrations (Fig. 1, D and E). Digitonin had more protein in the DRM fraction than the DSM fraction at lower detergent concentrations, but at 0.5% digitonin, very little protein was found in the DRM fraction (Fig. 1F). Overall, these findings challenge the notion that CL microdomains can be effectively isolated with biochemical detergent extraction. Therefore, we used a more controlled and less invasive model system to study CL microdomains.

The presence of cytochrome c induces proteolipid microdomain formation and morphological changes in GUVs and GMVs
We constructed GUVs, modeling the inner mitochondrial membrane, and GMVs to study CL microdomain organization. GMVs were constructed from native phospholipids extracted from isolated cardiac mitochondria. To visualize GUVs and GMVs, we employed two fluorescent probes, Texas Red DHPE (Texas Red), which is known to report on liquid-disordered domains of lipid membranes (43,44), and nonyl acridine orange (NAO), which is suggested to report on CL (45,46). Structures of the fluorescent probes are presented in Fig. S1, A and B.
We first visualized GUVs and GMVs in the absence of cytochrome c (Ϫcyt c) using Texas Red (Fig. 2, A and B, left panels), which revealed a homogenous distribution of the fluorophore throughout the membrane. The addition of 17 M cyt c to GUVs and GMVs containing Texas Red (Fig. 2, A and B, right panels) revealed fluorophore sequestration and the formation of proteolipid microdomains. The average fluorophore area was calculated using the equatorial cross-sectional view of GUVs and GMVs and was determined to decrease by 1.3-1.4fold upon the addition of cyt c (Fig. 2, C and D). The formation of phase-segregated domains was accompanied by clear membrane morphological changes with the GUVs and GMVs. To exemplify, the addition of cyt c decreased the average diameter of GUVs and GMVs with Texas Red (Fig. 2, E and F, respectively) by 1.8-fold.
We next visualized vesicles using the fluorescent probe NAO. Similar observations to Texas Red were made when GUVs and GMVs were visualized by NAO. In the absence of cyt c, the NAO fluorophore was homogenously distributed throughout the membrane of GUVs and GMVs, and no microdomains were observed (Fig. 3, A and B, left panels). Upon the addition of 17 M cyt c, strong phase segregation of NAO (Fig. 3, A and B, middle panels) was observed. Quantification revealed that 17 M cyt c decreased the average fluorophore Proteolipid domains are regulated by CL concentration area by up to 2.2-3.5-fold for GUVs and GMVs visualized by NAO (Fig. 3, C and D, respectively). Similarly, the diameter of the GUVs and GMVs also decreased upon the addition of cyt c by 1.5-2.5 (Fig. 3, E and F, respectively). We conducted colocalization analyses to confirm that the phase-separated domains were forming in regions where cytochrome c was associating. These studies revealed overlap (average Pearson's correlation coefficient of 45.1%) between NAO and an Alexa 647 antibody against cytochrome c (Fig. S2).
We also tested if cytochrome c-induced lipid microdomain formation depended on protein concentration. The addition of 3.8 M cyt c promoted lipid microdomain formation as observed by phase segregation of NAO in GUVs and GMVs (Fig. 3, A and B, right panels). The addition of 3.8 M cyt c to GUVs and GMVs also decreased the average area occupied by NAO (Fig. 3, C and D) as well as the average vesicle diameter (Fig. 3, E and F). Overall, Texas Red and NAO showed similar results (Figs. 2 and 3); therefore, for the subsequent imaging studies we used NAO given that is suggested to be CL specific (45,46).

Cytochrome c interactions with biomimetic and native mitochondrial lipid monolayers decreases the elasticity modulus
Given that cyt c decreased the diameters of GUVs and GMVs, we hypothesized that cyt c could be promoting changes in the elastic properties of the membrane. Therefore, we constructed biomimetic and native mitochondrial lipid monolayers, in the absence and presence of cyt c, to quantify the elasticity modulus (C s Ϫ1 ), which is an inverse measure of membrane elasticity. We first present raw surface pressure-area isotherms for both biomimetic (   2-fold. Therefore, these results demonstrate that cyt c regulates the elastic properties of biomimetic and native mitochondrial monolayers.
There is a debate in the literature whether which of these is more consequential (49). Therefore, we studied how a reduction of (18:2) 4 CL concentration and CL acyl chain composition influenced lipid microdomain organization in GUVs.

A reduction in (18:2) 4 CL concentration promotes favorable Gibbs-free energy of lipid mixing
Mechanistically, lipid domains are driven by the Gibbs-free energy of lipid mixing. Therefore, we used pressure-area isotherms to calculate the change in Gibbs-free energy between the absence and presence of cyt c. We first present the raw surface pressure-area isotherms for biomimetic mitochondrial monolayers containing (18:2) 4 CL (Fig. 6A), 50% depletion of (18:2) 4 CL concentration (Fig. 6B), or MLCL/(18:2) 4 CL (Fig. 6C) in the absence (black traces) and presence (red traces) of 3.8 M cyt c. Secondary analyses revealed that a 50% loss of (18:2) 4 CL significantly shifted the change in Gibbs-free energy of mixing upon cyt c addition toward a more favorable value by 2.5-fold ( Fig. 6D). In contrast, inclusion of MLCL had no effect on the thermodynamics of lipid mixing (Fig. 6D).

Proteolipid microdomain formation is not disrupted in GMVs synthesized using native mitochondrial phospholipids extracted from cardiac tissue of Acsl1 T؊/؊ mice
To further investigate how CL acyl chain composition regulates proteolipid domain organization, we relied on GMVs constructed from cardiac-specific acyl-CoA synthetase knockout mice (Acsl1 TϪ/Ϫ ). The rationale for this model was that CL acyl chains are notably remodeled, although total CL levels do not change (50). Specifically, previously published LC/MS analyses show that Acsl1 TϪ/Ϫ mice have increased incorporation of oleic acid (18:1) into CL at the expense of the loss of linoleic acid (18:2) (50).
Upon the addition of 3.8 M cyt c, proteolipid microdomain formation was observed in GMVs constructed from control ( Fig. 7A) and Acsl1 TϪ/Ϫ (Fig. 7B) mice. Proteolipid domains were similar in size between control and Acsl1 TϪ/Ϫ GMVs, as measured by the average area occupied by NAO (Fig. 7C). Furthermore, the diameter of the vesicles was also similar between the control and Acsl1 TϪ/Ϫ GMVs (Fig. 7D). Taken together, these results suggested that the remodeling of tetralinoleoyl-CL to other acyl chains does not impair the formation of cytochrome c-induced microdomains visualized by NAO fluorescence.
We further investigated how the accumulation of oleic acid in CL regulated proteolipid domain formation. The rationale for using the tetraoleoyl-CL species was that oleic acid is heavily accumulated in CL at the expense of linoleic acid in the murine knockout of Acsl1 TϪ/Ϫ (50). We specifically constructed biomi-

Proteolipid domains are regulated by CL concentration
metic GUVs containing tetraoleoyl-CL ((18:1) 4 CL) in the absence and presence of cyt c. In the absence of protein, NAO was homogenously distributed throughout the entire membrane of biomimetic GUVs containing (18:1) 4 CL (Fig. S3A). Upon addition of cyt c, NAO was sequestered into localized regions of the membrane (Fig. 3A). Similarly, cyt c-induced proteolipid domains were generally smaller in size, as indicated by a decreased average microdomain area and decreased vesicle diameter (Fig. 3, B and C).

Lipid microdomains form in the presence of mitochondrial creatine kinase in GUVs
We subsequently determined if other membrane-associated proteins could promote lipid microdomain formation in biomimetic mitochondrial GUVs. We tested the ability of purified recombinant ␣-synuclein (Fig. 8A) and mitochondrial creatine kinase (Fig. 8B) to induce proteolipid microdomains in GUVs. We selected ␣-synuclein (␣-syn) because its size is similar to cytochrome c, and mitochondrial creatine kinase (mtCK) because it is previously reported, in simpler model membranes, to bind to CL and promote domain formation (18, 51, 52). The addition of 3.8 M ␣-syn to biomimetic mitochondrial GUVs containing (18:2) 4 CL did not promote observable microdomains (Fig. 8A) because NAO was homogenously distributed throughout the entire GUV membrane. In contrast, mtCK promoted strong phase segregation of the NAO probe (Fig. 8B). Quantitatively, there was no change in the average area occupied by NAO for ␣-syn (Fig. 8C), whereas mtCK lowered the fluorophore area of GUVs (Fig. 8D). The diameter of the GUVs was maintained with ␣-syn (Fig. 8E) and lowered with mtCK (Fig. 8F).

(18:2) 4 CL oxidizes the heme cofactor iron-metal center in cytochrome c more effectively than MLCL
Finally, we investigated if the loss of CL concentration had a stronger effect than MLCL on the oxidation of cytochrome c. The rationale for this study was that MLCL may lack the ability to bind in the proper orientation near redox-sensitive cofactors, resulting in dramatic changes in midpoint potentials and impediments to electron transfer (53,54). By using UV-visual spectroscopy, we monitored the interaction of (18:2) 4 CL (Fig. 9A) and MLCL (Fig. 9B) with the heme cofactors of cytochrome c.

Proteolipid domains are regulated by CL concentration
Interactions between CL and the low spin, Fe(II)-heme center of the reduced protein, resulted in oxidation of the metal center to a high spin, Fe(III) center. These changes in cytochrome c's redox properties were dose-dependent as demonstrated by a blue-shift in the Soret bands (400 nm, data not shown) and decrease in ␣-heme bands (540 -560 nm, Fig. 9). Plots of the extent of cytochrome c oxidation versus lipid concentration (Fig. 9C) show that a higher concentration of MLCL (closed circles) was needed to fully oxidize cytochrome c relative to (18:2) 4 CL (open circles), and a dose-dependent effect of (18: 2) 4 CL concentration on the redox properties of cyt c.

Biochemical detergent extraction is ineffective for isolating CL-enriched domains
One major advancement from this study was data demonstrating that detergent extraction was ineffective for isolating and studying CL-enriched domains. We investigated the efficacy of three different detergents for extracting CL-enriched domains because CL domains are reported to be isolatable using nonionic detergents at cold temperatures (17). Triton X-100 was used because it is the preferred detergent for lipid domain extraction (55,56). Nonidet P-40 was selected due to its nonionic nature and digitonin was relied on given its use for isolation of mitochondrial supercomplexes (57). The analyses revealed that detergent extraction displayed no specificity for CL-enriched domains because all phospholipids and proteins were generally solubilized with increasing detergent concentrations. Thus, similar to conclusions made in the study of plasma membrane lipid rafts, detergent was not an effective tool for studying CL domains (58).

CL-cytochrome c interactions promote domain formation and membrane morphological changes
Given the limitations of detergent extraction, an imaging approach was used to study mitochondrial inner membrane domains. The novelty of our approach was in the combined analysis of native mitochondrial and biomimetic vesicles that represent or model the phospholipid composition of the inner mitochondrial membrane. The observation that proteolipid    (22). The observed morphological changes with CL were also consistent with a previous study that suggested interactions between CL and cytochrome c in raft/nonraft mixtures promoted membrane budding and ultimately formation of a folded structure (24). The positively charged NAO was likely interacting with negatively charged regions and thereby promoting the formation of cyt c multimers associated with structural modifications. Alternatively, the binding of NAO to the membranes may be simply a reflection of ion-ion interactions.
Supporting monolayer studies showed that the addition of cytochrome c was disrupting membrane packing accompanied by changes in the elasticity modulus. The morphological changes were of biological significance because CL domains are hypothesized to form in select regions of the membrane, specifically in areas of high membrane curvature (59). For instance, Renner and Weibel (16) reported that CL domains localized in

Proteolipid domains are regulated by CL concentration
regions of high curvature induced with microchambers using an Escherichia coli model. Thus, proteolipid domains in the inner mitochondrial membrane may be localized to regions of high curvature induced by strong interactions between cytochrome c and CL.
We did not observe proteolipid domain formation with the intrinsically disordered protein ␣-synuclein. The rationale for using ␣-synuclein was based on evidence that it can interact with mitochondria to diminish complex I activity (60). Furthermore, ␣-syn is known to bind acidic phospholipids although these interactions have been shown to be significantly augmented by the presence of phosphatidylethanolamine, a neutral phospholipid (61). The results with mitochondrial creatine kinase were consistent with previous literature demonstrating the ability of this protein to promote the formation of proteolipid domains (18,52). Notably, creatine kinase is also hypothesized to localize to cristae sites where CL is enriched (62,63).

A reduction in tetralinoleoyl-CL concentration but not substitution with MLCL prevents proteolipid domain formation through changes in lipid miscibility
In conditions such as Barth syndrome, cardiovascular diseases, obesity, and diabetes, CL levels are lowered or undergo changes in acyl chain composition. It is hypothesized that such changes may be detrimental to mitochondrial function. To exemplify, in cardiac ischemia-reperfusion injury, total (18: 2) 4 CL content is selectively depleted (32,48,64). Therefore, our data provide a potential mechanism by which mitochondrial function may become impaired. That is, perhaps a reduction in total (18:2) 4 CL content disrupts formation of mitochondrial proteolipid domains. Furthermore, the loss of CL in ischemia is also associated with the loss of cyt c, which triggers apoptotic signaling cascades (65,66). Thus, formation of CL-cyt c domains may be a critical mechanistic link and perhaps a therapeutic target for preventing the robust apoptosis observed with cardiac ischemia-reperfusion injury (65,66).
The result that MLCL did not perturb proteolipid domain organization does not preclude other mechanisms by which MLCL may exert its effects on mitochondrial bioenergetics. For instance, MLCL is not as tightly bound as CL to respiratory proteins, which occurs in response to CL degradation in Barth syndrome (39). Furthermore, increasing levels of MLCL are likely to influence membrane properties outside of microdomain formation such as viscosity and permeability. Thus, it is plausible that MLCL could impair respiratory enzyme activities and electron channeling. Future studies will be needed to investigate how MLCL disrupts differing aspects of membrane biophysical organization and thereby protein-lipid and proteinprotein interactions.
There are several emerging studies suggesting that CL acyl chain composition is not a major driver of mitochondrial membrane structure-function. Our results with MLCL on microdomain formation and cytochrome c oxidation status were consistent with these previous studies. For example, in a yeast model, remodeled and unremodeled CL were indistinguishable when analyzing oxidative phosphorylation activity (49). Similarly, CL content varies between differing tissues, which further argues against the notion that CL acyl chain composition is a key regulator of bioenergetics function. In addition, restoration of WT CL levels in differing disease models does not rescue mitochondrial functional responses. For instance, in aging or in a previous study with the heart-specific acyl-CoA 1 synthetase knockout, restoration of (18:2) 4 CL did not improve key mitochondrial end points (50,67).
It is important to point out that in some cases of extreme CL remodeling, the acyl chains may be major drivers of changes in mitochondrial bioenergetics. We previously demonstrated that dietary docosahexaenoic acid promoted robust CL remodeling, which lowered cardiac respiratory enzyme activities through changes in protein-CL binding and formation of membrane microdomains (68).
Although our data suggest that acyl chain composition may not be a major driver of mitochondrial microdomains, additional studies are needed to understand how CL oxidation products influence mitochondrial inner membrane physical properties. The use of the 18:1-containing CL would be a limitation given that it is not prone to generating downstream second messengers. Future studies will need to specifically investigate how oxidation products, which are known to influence bioenergetics, could regulate mitochondrial structure and thereby function (69).

Conclusion
The data demonstrate that CL-specific microdomains cannot be effectively isolated via classical biochemical detergent extraction techniques. More importantly, membrane domains were formed in GUVs and newly constructed GMVs with cytochrome c. These domains were sensitive to a reduction in CL concentration but not substitution with MLCL. At a molecular level, the impairment in domain formation with the loss of tetralinoleoyl-CL was due to changes in the Gibbs-free energy of lipid mixing. Collectively, the results have strong implications for the ongoing debate about CL concentration and acyl chain composition and their role in regulating membrane biophysical organization.
All other reagents and solvents (HPLC grade) were purchased from Sigma or Fisher Scientific. Lipid mixtures were handled under low light conditions and fresh stocks of lipid mixtures were used for each experiment to minimize oxidation.

Isolation of mitochondria from rodent cardiac tissue
Mitochondrial isolations were performed on ice and all buffers and instruments were pre-chilled to 4°C. All animal experiments were conducted in accordance with guidelines established by the Guide for Care and Use of Laboratory Animals and with prior approval by the University's Animal Care and Use Committee. Adult male C57BL/6J mice we purchased from Jackson Laboratories and were housed on a 12:12-h light:dark cycle and had unrestricted access to standard chow and water. Adult male Sprague-Dawley rats (150 -250 g) were obtained from Charles River and were housed on a 12:12-h light:dark cycle and had unrestricted access to standard chow and water. Acsl1 TϪ/Ϫ mice were maintained as previously described (50). One animal was sacrificed per isolation and cardiac tissue was collected immediately following euthanasia. Mice were anesthetized by isoflurane. Rats were anesthetized with ketamine/ xylazine (90 mg/kg of ketamine, 10 mg/kg of xylazine, i.p.) and after the absence of animal eye-blink, toe pinch, and righting reflexes, rats were sacrificed. Cardiac tissue was excised and immediately placed in pre-chilled mitochondrial isolation medium (MIM) containing 20 mM HEPES (pH 7.2), sucrose (300 mM), and EDTA (1 mM). Rodent cardiac tissue was manually minced for 5 min and subjected to homogenization (7 strokes) with a Teflon Potter glass homogenizer. The homogenate was centrifuged at 500 ϫ g for 5 min and the decanted supernatant was centrifuged again at 500 ϫ g for 5 min. The supernatant was decanted and centrifuged at 10,000 ϫ g for 15 min, and the resulting pellet was washed with MIM prior to centrifugation at 12,000 ϫ g for an additional 15 min. The resulting mitochondrial pellet(s) were resuspended in either 300 l of MIM (for rat cardiac mitochondria) or in 150 l of MIM (for murine cardiac mitochondria) and frozen at Ϫ80°C. Total mitochondrial protein concentrations were determined prior to freezing using a BCA protein assay (ThermoScientific, La Jolla, CA).

Extraction of rodent cardiac mitochondrial detergentresistant (DRM) and detergent-soluble (DSM) membranes and analyses
Experimental procedures were modified from an established protocol for isolating cardiolipin-enriched DRMs (17,70). Briefly, 1.0 mg of isolated mitochondria was incubated in a Beckham Coulter Centrifuge Tube (355635) on ice for 10 min using an extraction buffer containing 25 mM HEPES (pH 7.5), 150 mM sodium chloride, 10 g/ml of aprotinin and various concentrations of Triton X-100, digitonin, or Nonidet P-40 detergents. The solution was centrifuged in an Optima Max-XP Ultracentrifuge with a TLA 100.4 rotor at 20,000 ϫ g for 3 min at 4°C. The detergent-soluble fraction was extracted from the centrifuge tube and the detergent-resistant pellet was resuspended in a solubilization buffer containing 50 mM Tris-HCl (pH 8.8), 1% SDS, and 5 mM EDTA. DNA was sheared from the detergent-resistant fraction by passage through a 1.00-ml syringe and 20 1 ⁄ 2-gauge needle. The fractions were then transferred to a fresh glass tube for lipid extraction. TLC was conducted on DRM and DSM fractions as previously described (53). Protein levels in the fractions were also quantified using a standard BCA assay kit.

Extraction of native phospholipids from isolated rodent cardiac mitochondria
Experimental procedures were modified from the Bligh and Dyer method (71). Frozen aliquots of rodent cardiac mitochondria were thawed and transferred into a glass test tube. For every 10 mg/ml of mitochondria, samples were resuspended in 10 ml of 70/30 (v/v) CHCl 3 /MeOH (0.05% BHT) solution and vortexed for 45 s. Upon vortex mixing, 2.5 ml of 0.88% KCl was added to each sample and vortexed for an additional 45 s. The resulting mixture was centrifuged at 2000 rpm for 10 min at room temperature (23°C). Upon separation of phases, the bottom layer (organic) was aspirated and dried via a gentle stream of N 2 gas. The extract was resuspended in 2.0 ml of chloroform and a second extraction was performed. Upon recovering the lipid-containing organic layer, the sample was aspirated and dried via a gentle stream of N 2 gas prior to being resuspended in HPLC grade chloroform (final concentration 0.1-0.25 mol/25 l). Total phosphorous analysis of extracted native mitochondrial phospholipids was conducted as previously described (31).

Construction of GUVs and GMVs
GUVs were constructed by co-dissolving lipids (39.9 mol % 18:0 -22:6 PC, 30.0 mol % 16:0 -20:4 PE, 20 mol % (18:2) 4 CL, 5 mol % DOPI, 3 mol % DOPS, 2 mol % Chol), and either the fluorescent probe Texas Red DHPE or NAO (0.1 mol %) in chloroform (0.5 mg/ml). The concentration of PC, PE, CL, PI, PS, and Chol approximated ratios from shotgun lipidomic analyses (data not shown) and were consistent with the literature (7). For select GUV experiments in which CL's content and acyl chain composition was altered, (18:2) 4 CL concentration was decreased 50% by mass or replaced with MLCL. GMVs were constructed by co-dissolving extracted native mitochondrial phospholipids in HPLC-grade chloroform. Roughly 10 g of total lipid was spread onto the conductive side of an indium tin oxide-coated glass slide. The lipid-coated slide was subjected to dark vacuum for 1-2 h to remove excess solvent. Once the lipid film was dried, an electroformation chamber was assembled as described (72). GUVs and GMVs were constructed by electroformation at room temperature as previously described (26,72,73). Once electroformation was complete, vesicles were extracted from the chamber using a 20 1 ⁄ 2-gauge needle/syringe and allowed to equilibrate briefly prior to microscopy sample preparation. For select experiments, the addition of cyt c (3.8 -17 M final concentration), as well as other proteins, to GUVs and GMVs occurred immediately prior to imaging. Samples were drawn into a rectangular capillary tube (VitroTubes) and mounted onto a glass microscope slide as previously shown (68,74).

Confocal microscopy and image analysis
Imaging was conducted with an Olympus FV1000 Confocal Microscope using a ϫ60/1.35NA oil immersion objective (Olympus, Waltham, MA) and a Zeiss LSM 880 with Airyscan using a ϫ63/1.40NA oil immersion objective (Carl Zeiss

Proteolipid domains are regulated by CL concentration
Microscopy, Thornwood, NY). The Texas Red DHPE probe and Alexa 594 secondary antibody were excited by a diode laser at 559 nm, whereas the NAO probe was excited with an argon laser at 488 nm. A diode laser at 633-635 nm excited the Alexa 647 primary antibody used for cyt c immunofluorescence.
All acquired images were of GUV equatorial cross-sections. Lipid microdomain and colocalization analysis was conducted using NIH ImageJ software (74). After background subtraction, the average area occupied by the fluorophore was determined by adjusting the image threshold and analyzing particles that were Ն0.1 m 2 . We selected this threshold, because an area Ͻ0.1 m 2 resulted in particles/domains that could not be accurately measured. The average vesicle diameter was determined by measuring the horizontal distance between the membrane edges. For colocalization measurements, we determined the fraction of cytochrome c colocalized with NAO using the Coloc 2 and JACoP plugins. Colocalization was quantified after background subtraction using Pearson's correlation coefficient.

Construction and analysis of biomimetic and native mitochondrial monolayers
Biomimetic mitochondrial monolayers were generated by co-dissolving lipids (40 mol % 18:0 -22:6 PC, 30.0 mol % 16:0 -20:4 PE, 20 mol % (18:2) 4 CL, 5 mol % DOPI, 3 mol % DOPS, 2 mol % Chol) in chloroform (10 g/l). For select lipid monolayer experiments in which CL's content and acyl chain composition was altered, (18:2) 4 CL concentration was decreased 50% by mass or replaced with MLCL. Native mitochondrial monolayers were initially generated by aliquoting native mitochondrial phospholipids (0.25 mol/25 l) extracted from murine (C57BL6/J) cardiac tissue. Lipid monolayers were constructed by spotting either 9.0 nmol (biomimetic mitochondrial monolayers) or 6.0 nmol of total lipid (native mitochondrial monolayers) on a subphase of 10 mM sodium phosphate buffer (pH 7.4). Biomimetic and native mitochondrial lipid monolayers were analyzed in the presence and absence of cytochrome c (cyt c; 3.8 -17 M final concentration) by gently injecting protein underneath the subphase immediately after spotting lipid. Excess chloroform was allowed to evaporate for 10 min prior to monolayer compression. Pressure-area isotherms were analyzed at physiologically relevant surface pressures (i.e. 30 mN/m) as previously described (26,68). Raw isotherms were used to calculate the surface elasticity modulus (C s Ϫ1 ), A represents the mean molecular area of the lipid mixture of interest at the indicated surface pressure () (75). The Gibbsfree energy of mixing (⌬G mix ) was calculated by the following relationship to quantify lipid-lipid mixing, ⌬G ex represents the excess Gibbs-free energy of mixing and ⌬G ideal represents the ideal Gibbs-free energy of mixing, which were calculated by the following expressions, A n and X n are the mean molecular area and mol fraction of each component, respectively, at a given surface pressure () (76). ⌬G ideal was determined by, R represents the ideal gas constant and T is temperature in Kelvin (76). All lipid mixtures were acquired multiple times on differing days to ensure reproducibility.
Overexpression and purification of recombinant ␣-synuclein Electrocompetent E. coli BL21 STAR (dE3) cells were transformed with expression plasmid pReceiver-B31 (GeneCopeia, Rockville, MD) encoding for the N-terminally His-tagged ␣-synuclein. Protein was expressed in 20-liter batch cultures of LB media, supplemented with 200 g/ml of ampicillin for plasmid maintenance, and inoculated with a 1-liter overnight culture of transformed E. coli. Large-scale fermentations were carried out in a 30-liter carboy equipped with an air inlet hose connected to, and fitted with, a gas diffusion stone for proper aeration and agitation of the media during growth and induction. Batch cultures were grown at 37°C to an A 600 ϭ 0.9 -1.2. The culture was then chilled on ice for 20 min prior the addition of isopropyl 1-thio-␤-D-galactopyranoside (final concentration 1 mM). Protein expression was induced at 16°C for ϳ20 -24 h. Cells were harvested via centrifugation, yielding 100 -200 ϫ g of cell paste. Cell paste was flash frozen in liquid N 2 and stored at Ϫ80°C until needed.

Overexpression and purification of human mtCK
The human sarcomeric mtCK localized in skeletal and heart muscle was expressed in bacteria as a tag-free protein and was Proteolipid domains are regulated by CL concentration purified in a two-dimensional HPLC protocol as previously described (54). Purified protein was aliquoted and stored at Ϫ20°C.

UV-visible spectroscopy and oxidation of cytochrome c
UV-visible spectra of cytochrome c were recorded with a Shimadzu 1800 spectrophotometer fitted with a thermostated cell holder. Stock solutions of proteins, lipids, and buffers/substrates were made fresh prior to use. The 3-ml reactions contained HEPES (30 mM, pH 7.4), catalase (10 g/ml), glucose (50 M), glucose oxidase (50 g/ml), and cytochrome c (5 M). Initial UV-visible spectra were recorded (200 -800 nm) and ascorbate (final concentration 10 M) was added to completely reduce the cytochrome c. Spectra were recorded every minute for 5 min after the addition of ascorbate to ensure all cytochrome c was reduced. 2-3 l of 1 mg/ml of solubilized lipid (DOPC, (18:2) 4 CL, and MLCL) was subsequently titrated into the reaction mixture, and after a 3-min equilibration period, spectra were recorded. After all titrations were complete, K 3 [FeCN6] (55 M, final concentration) was added, and after a 3-min equilibration period, the final spectra of fully oxidized cytochrome c were recorded.

Statistics
Data were analyzed with GraphPad Prism version 7.0. Data were ensured to display normalized distributions. Statistical analyses were conducted using unpaired two-tailed t-tests or one-way analysis of variance followed by a post hoc Bonferroni test. p values less than 0.05 were considered significant. Results are plotted as mean Ϯ S.D. from multiple independent experiments.