In parallel with these findings in NPC, there is also evidence of autophagy defects in NPA/B diseases (Fucho et al., 2014; Baulies et al., 2015; Canonico et al., 2016). For instance, hepatocytes from ASMase−/− mice exhibit impaired autophagy flux determined by the combination of rapamycin with or without chloroquine, an effect that was accompanied by increased LC3BII and p62 levels. In addition, ASMase−/− hepatocytes displayed impaired fusion of autophagosome-containing mitochondria with lysosomes in response to acetaminophen, which translated in increased susceptibility to acetaminophen-mediated liver injury by sustaining mitochondrial damage (Baulies et al., 2015). Although ASMase deficiency results primarily in increased lysosomal SM content, it also causes lysosomal cholesterol accumulation, which accounts for the sensitization to acetaminopheninduced hepatocellular injury. In line with these findings, U18666A, an amphiphilic amino-steroid that induces lysosomal cholesterol accumulation sensitized primary hepatocytes to acetaminophen-induced cell death. Treatment with oxysterol 25-hydroxycholesterol, a ligand for liver X receptors (LXR) that suppresses sterol synthesis, decreases cholesterol accumulation and protects ASMase−/− mice from acetaminophen-mediated liver injury. Moreover, human B lymphocytes from patients with NPB disease exhibit alterations in the rate of autophagic vacuole accumulation, mitochondrial fragmentation and mitophagy induction, indicating impaired clearance of damaged mitochondria (Canonico et al., 2016). Thus, taken together these findings indicate that defective clearance of dysfunctional mitochondria due to impaired lysosomal function or defective fusion of lysosomes with autophagosomes is a common feature of both NPA/B and NPC diseases.

### MITOCHONDRIAL CHOLESTEROL ACCUMULATION: A DIFFERENTIAL FEATURE BETWEEN NPA/B AND NPC DISEASES

In contrast to defective autophagy, which is common to NPA and NPC, intracellular cholesterol trafficking and accumulation in mitochondria is a differential feature between NPA/B and NPC diseases (**Figure 2**). Although the primary feature of NPA is the accumulation of SM due to the deficiency of ASMase, cholesterol levels also increase through the regulation of SREBP2. However, the bulk of the intracellular cholesterol accumulation in NPA is sequestered in lysosomes as shown in hepatocytes from ASMase null mice stained with filipin and lysotracker (Fucho et al., 2014). In contrast to this event, there is growing evidence of increased mitochondrial cholesterol loading in both hepatocytes and neurons from Npc1−/− mice (**Figure 2**), and this process mediates, in part, the mitochondrial dysfunction reported in NPC disease (Yu et al., 2005; Marí et al., 2006; Charman et al., 2010; Balboa et al., 2017; Torres et al., 2017). Moreover, while alcohol induced the trafficking and accumulation of cholesterol in hepatic mitochondria, this event was defective in hepatocytes deficient in ASMase, which paralleled the lack of induction of StARD1 in ASMase null mice in response to alcohol feeding (Fernandez et al., 2013). The mechanism underlying the differential increase in mitochondrial cholesterol and expression of StARD1 between NPA and NPC

is not currently understood (Torres et al., 2016). As acid ceramidase (ACDase) has been shown to repress steroidogenic factor-1-dependent expression of StARD1 (Lucki et al., 2012), it remains to be established whether decreased ACDase in NPC disease contributes to the upregulation of StARD1 and subsequent mitochondrial cholesterol loading (**Figure 2**), which is currently under investigation. Moreover, as mentioned above, NPC2 has been shown to contribute to the transport of endosomal cholesterol to mitochondria independently of NPC1 (Kennedy et al., 2012). As the loss of function of NPC2 accounts for a minor fraction of NPC cases, it is conceivable that NPC2 may play a role in the trafficking of endosomal cholesterol to mitochondria in most NPC patients (Kennedy et al., 2012).

Increased mitochondrial cholesterol content in NPC cells can lead to important functional consequences, such as decreased mitochondrial membrane fluidity (Colell et al., 2003), reduced ATP generation (Echegoyen et al., 1993; Yu et al., 2005), and decreased mitochondrial GSH (mGSH) import (Marí et al., 2006; Garcia-Ruiz et al., 2009). In addition, mitochondrial membrane potential and the activity of ATP synthase are markedly decreased in NPC1 mouse brains and neurons (Yu et al., 2005; Balboa et al., 2017; Torres et al., 2017) and the inhibition of ATP synthase is reversible upon mitochondrial cholesterol extraction by cyclodextrin (Yu et al., 2005). In line with these findings, increased mitochondrial cholesterol loading triggers metabolic adaptations in NPC models, characterized mainly by enhanced glycolysis (Kennedy et al., 2014). The depletion of mGSH levels has a severe impact in NPC disease, as inferred by its replenishment with GSH ethyl ester (GSH-EE), which restored the mGSH pool in liver and brain of Npc1−/− mice and in fibroblasts from NPC patients, leading to increased median survival and maximal life span of Npc1−/− mice, protection against oxidative stress and oxidant-induced cell death and restoration of calbindin levels in cerebellar Purkinje cells, which improved locomotor impairment in Npc1−/− mice. In addition, high-resolution respirometry analyses showed that GSH-EE treatment improved oxidative phosphorylation, coupled respiration and maximal electron transfer in cerebellum of Npc1−/− mice (Torres et al., 2017). In contrast, the antioxidant and cytosolic GSH precursor N-acetylcysteine (NAC), which increased total GSH levels in liver and brain homogenates from Npc1−/− mice (Fu et al., 2013), failed to restore the mGSH pool (Torres et al., 2017). These results are in agreement with the notion that increased cholesterol content in NPC mitochondria disrupts membrane dynamics and impairs the transport of GSH from the cytosol into mitochondria (Marí et al., 2009), contributing to the activation of apoptotic pathways seen in NPC mutant cells (Klein et al., 2011). In line with the role of oxidative stress, it has been shown that vitamin E supplementation in the diet delayed weight loss, improved coordination and locomotor function and increased the survival of Npc1−/− mice (Marín et al., 2014). Moreover, vitamin E supplementation preserved Purkinje neurons and reduced levels of astrogliosis, nitrotyrosine and apoptotic signaling mediated by the c-Abl/p73 pathway (Marín et al., 2014). Part of this beneficial effect could be related to increased levels of α-tocopherol in mitochondria, which has been shown to quench ROS production, especially in hepatic mitochondria (Chow et al., 1999), thus preventing GSH depletion (Stojkovski et al., 2013).

In addition to these events, mitochondrial cholesterol loading may impair mitochondrial dynamics reflected in the balance between fusion and fission events. Disruption of appropriate mitochondrial fluidity following cholesterol accumulation can prevent the fusion of mitochondria with adjacent healthy mitochondria (Baker et al., 2014), leading to increased predominance of fragmented mitochondria. In line with these events, we have recently reported that mitochondria from NPC cells exhibited a more rounded and smaller morphology compared to wild type cells, paralleling the decrease in mitochondrial membrane potential and increased mitochondrial superoxide production (Balboa et al., 2017). Finally, disruption of lysosomal-mitochondrial interplay can also have an impact in the Ca2+ buffering capacity of mitochondria (Kiselyov and Muallem, 2008) in LSDs, in line with the effect of suppressing lysosomal function and autophagy in mitochondrial Ca2+ homeostasis (Jennings et al., 2006). As mentioned above, it has been recently shown that NPC disease is characterized by the disruption of lysosomal Ca2+ homeostasis preceded by the accumulation of sphingosine that impairs the endocytic pathway (Lloyd-Evans et al., 2008). Whether this event is linked to altered mitochondrial Ca2+ content and function remains to be established. In addition, further work is needed to demonstrate a causal role for the disruption of Ca2+ homeostasis in the death of the Purkinje neurons in NPC disease, as shown in other related diseases (Girard et al., 2012; Kasumu and Bezprozvanny, 2012).

### THERAPEUTIC APPROACHES AND CONCLUDING REMARKS

Although the genetic causes of NPA and NPC disease are well understood, the consequences at the level of disruption of intracelular trafficking and interorganelle communication is still incomplete, thus limiting the availability of effective therapy. Enzyme replacement therapy for ASMase and NPC1 deficiency is expected to successfully treat peripheral non-neuronal symptoms of both diseases (Schuchman and Desnick, 2017). However, as this approach may be inefficient to replenish the defective enzymes in the brain due to the blood brain barrier, the expectations for the correction of the neurological symptoms may be disapointing. Therefore, there is the urgent need for alternative possibilites to target both diseases exploiting the fact that several intracelular compartments are involved in the pathogenesis of NPA and NPC diseases (Saffari et al., 2017). In the

### REFERENCES

case of NPC, in which mitochondrial dysfunction and cholesterol accumulation have been well established, strategies targeting mitochondrial cholesterol accumulation and/or downstream consequences may be of potential relevance. In this scenario, recent data indicated that the replenishment of mGSH levels by permeable GSH prodrugs have shown promising results in fibroblasts from NPC patients, correcting the mitochondrial dysfunction and ameliorating oxidative stress, while extending the survival of NPC1 null mice (Torres et al., 2017). Quite intriguingly, the efficacy of mGSH replenishment by GSH-EE was not potentiated by parallel treatment with cholesterol extraction with cyclodextrin, suggesting that both processes are related to each other. This outcome raises the possibility that the efficacy of mGSH restoration by GSH-EE could be explored in combination with other therapies targeting differential pathways. For instance, since curcumin has been shown promising results by restoring intracellular sphingosine balance and cytosolic calcium homeostasis (Lloyd-Evans et al., 2008), it may be worth investigating the combination of curcumin with GSH-EE in NPC disease. Moreover, recent evidence has shown a beneficial role for histone deacetylase (HDAC) inhibitors (i.e., vorinostat or valproic acid) as a novel line of therapy for NPC disease, resulting in the reduction of cholesterol accumulation, improvement of neurodegenerative symptoms and visceral alterations, which culminate in the extension of mouse life span (Kim et al., 2007; Pipalia et al., 2011; Alam et al., 2016; Contreras et al., 2016; Munkacsi et al., 2017). As HDAC can regulate mitochondrial function, it may be worth to test whether combined treatment with HDAC inhibitors and GSHEE would exhibit greater therapeutic benefit in NPC1 mutant mice with respect to either agent alone, supportingh this approach as a promising combination therapy for NPC.

### AUTHOR CONTRIBUTIONS

ST, EB, SZ, CE, CG-R, and JF-C discussed findings, analyzed literature and wrote the manuscript. ST, CG-R, and JF-C designed the schematic Figures.

### ACKNOWLEDGMENTS

The work was supported by grants SAF-2014-57674-R, SAF-2015-69944-R from Plan Nacional de I+D, Spain, Fundació Marató de TV3, La Mutua Madrileña, PI11/0325 (META) grant from the Instituto Salud Carlos III, and by the support of CIBEREHD; the center grant P50-AA-11999 Research Center for Liver and Pancretic Diseases funded by NIAAA/NIH; and support from AGAUR of the Generalitat de Catalunya 2014- SGR785.

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**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2017 Torres, Balboa, Zanlungo, Enrich, Garcia-Ruiz and Fernandez-Checa. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Interorganelle Communication between Mitochondria and the Endolysosomal System

#### Gonzalo Soto-Heredero1 , Francesc Baixauli 2 and María Mittelbrunn1, 3 \*

1 Centro de Biología Molecular Severo Ochoa (CSIC-UAM), Madrid, Spain, 2 Max Planck Institute of Immunobiology and Epigenetics, Freiburg, Germany, 3 Instituto de Investigación Hospital 12 de Octubre, Madrid, Spain

The function of mitochondria and lysosomes has classically been studied separately. However, evidence has now emerged of intense crosstalk between these two organelles, such that the activity or stress status of one organelle may affect the other. Direct physical contacts between mitochondria and the endolysosomal compartment have been reported as a rapid means of interorganelle communication, mediating lipid or other metabolite exchange. Moreover, mitochondrial derived vesicles can traffic obsolete mitochondrial proteins into the endolysosomal system for their degradation or secretion to the extracellular milieu as exosomes, representing an additional mitochondrial quality control mechanism that connects mitochondria and lysosomes independently of autophagosome formation. Here, we present what is currently known about the functional and physical communication between mitochondria and lysosomes or lysosome-related organelles, and their role in sustaining cellular homeostasis.

#### Edited by:

Nuno Raimundo, Universitätsmedizin Göttingen, Germany

#### Reviewed by:

Laura Elizabeth Swan, University of Liverpool, United Kingdom Nina Kaludercic, National Research Council of Italy (CNR), Italy

\*Correspondence:

María Mittelbrunn<EMAIL_ADDRESS>
#### Specialty section:

This article was submitted to Mitochondrial Research, a section of the journal Frontiers in Cell and Developmental Biology

Received: 25 September 2017 Accepted: 23 October 2017 Published: 07 November 2017

#### Citation:

Soto-Heredero G, Baixauli F and Mittelbrunn M (2017) Interorganelle Communication between Mitochondria and the Endolysosomal System. Front. Cell Dev. Biol. 5:95. doi: 10.3389/fcell.2017.00095 Keywords: lysosome, exosomes, autophagy, quality control, proteostasis, aging

### INTRODUCTION

Eukaryotic cells contain membrane-confined organelles that enable them to compartmentalize specialized biochemical reactions like ATP production, lipid breakdown or protein degradation, performing them at specific cellular locations. To fully benefit from the clear advantages of such specialization, organelles within a cell must communicate, and exchange information and metabolites. Indeed, it is now well accepted that such interactions are necessary to maintain cell homeostasis. Communication between organelles does not necessarily require physical contacts, as information between organelles can be transmitted through transfer of specific metabolites. However, membrane contact sites (MCSs) represent a fast and efficient way to communicate. Such interorganelle physical contacts are currently the focus of much study and they have been associated to many cellular processes, such as signaling, the regulation of calcium homeostasis, lipid metabolism, and organelle localization and dynamics (Helle et al., 2013; Daniele and Schiaffino, 2014). Interactions between organelles are also very important for protein quality control, since misfolded, oxidized and obsolete proteins are dangerous to the cell, and they must be handled suitably to maintain correct proteostasis. This problem is resolved in the cell by organelles working together in a complex and coordinated network (Baixauli et al., 2014; Gottschling and Nyström, 2017).

Mitochondria and lysosomes are at the center of devastating genetic diseases. Research into mitochondrial and lysosome-originated diseases has focused on trying to understand the molecular pathology of the affected organelle. However, it was recently discovered that genetic defects in one type of organelle may lead to an impairment of the other. Specifically, genetic disruption of mitochondria produces a secondary impairment of lysosomes, and genetic defects in lysosomes can lead to mitochondrial dysfunction, highlighting the functional connection between these two organelles.

Mitochondria are fundamental metabolic organelles in cells, but they also participate in iron and calcium homeostasis, as well as in the regulation of apoptosis, and they are increasingly recognized as key signaling platforms (Friedman and Nunnari, 2014; Chandel, 2015). Like the rest of organelles, mitochondria need to interact with other organelles to carry out their functions, exchanging material and transmitting signals responsible for regulating metabolism, intracellular signaling and cell maintenance (Murley and Nunnari, 2016). This communication can be established in different ways, such as vesicular transport (as initially revealed for organelles within the secretory pathway), the exchange of metabolites or signaling molecules by diffusion, or via direct physical contacts.

The best characterized MSCs are the physical contacts established between the mitochondria and endoplasmic reticulum (ER). These contacts are named mitochondrialassociated membranes (MAMs) and they are critical for many intracellular processes, particularly the regulation of calcium homeostasis, lipid metabolism, mitochondrial fission or regulation of mitochondrial DNA synthesis and division (Friedman et al., 2011; Bravo-Sagua et al., 2014; Lewis et al., 2016). These contacts allow the flux of calcium between the two organelles, coordinating ATP production and facilitating mitochondrial calcium buffering. Under specific conditions, the distance between the mitochondria and ER may vary, affecting their communication and organelle function. For example, tight junctions between mitochondria and ER can cause mitochondrial calcium overload, and mitochondrial membrane permeabilization, driving the cell into an apoptotic programme (Csordás et al., 2006).

Interestingly, defects in MAMs, or in their homeostasis, have been associated with certain diseases, highlighting the importance of the contacts between organelles for controlling both cell and organism homeostasis. Obesity induces an increase of MAMs in the liver, which compromises mitochondrial activity and drives oxidative stress (Arruda et al., 2014; Ma et al., 2017). In obese animals, experimental induction of MAMs deteriorates metabolic homeostasis, while their disruption improves the oxidative capacity of mitochondria (Arruda et al., 2014). MAMs have also been linked to inflammation and there is evidence suggesting that MAMs are involved in triggering an inflammatory response to oxidative stress. Inducers of reactive oxygen species (ROS) activate the inflammasome, driving its localization to MAMs (Zhou et al., 2011). MAMs have been related to neurodegenerative diseases (Manfredi and Kawamata, 2016; Paillusson et al., 2016), such that their frequency and activity are enhanced in both familial and sporadic Alzheimer's disease patients (Area-Gomez et al., 2012). Reduced numbers of ER–mitochondria associations and alteration in Ca2+ exchange between these two organelles have been observed in two different amyotrophic lateral sclerosis (ALS) mouse models (Paillusson et al., 2016). The loss of MAM homeostasis has also been linked to cancer, diabetes and pulmonary arterial hypertension (Sutendra et al., 2011; Giorgi et al., 2015; Ma et al., 2017; Rodríguez-Arribas et al., 2017).

The endolysosomal system is a highly dynamic membrane compartment that is critical in the maintenance of cellular homeostasis. The endolysosomal system is made up of several membrane-bound organelles, such as early endosomes, multivesicular bodies (MVBs), lysosomes, lysosome-related organelles (LROs) and other specialized organelles (Klumperman and Raposo, 2014). During endosome maturation, inward budding of the limiting membranes forms small intraluminal vesicles (ILVs) in organelles referred to as MVBs. Endosomes, MVBs and lysosomes regulate signaling, secretion, and the degradation of receptors and other cellular components. MVBs can fuse with the plasma membrane and release their ILVs into the extracellular environment as exosomes, or alternatively, they can fuse with lysosomes for their content to be degraded. Besides degradation, lysosomes carry out many other critical functions, acting as acidic calcium stores that release calcium in response to different stimuli, secreting their content through exocytosis, or repairing the plasma membrane (Perera and Zoncu, 2016). Furthermore, lysosomes can sense the metabolic state of the cell, as it occurs in conditions of starvation or stress, in which mTORC1 is activated and released from the lysosomal membrane, triggering a transcriptional response to increase lysosome biogenesis and activity (Roczniak-Ferguson et al., 2012; Settembre et al., 2013; Ballabio, 2016).

This review will explore some of the more recent findings regarding the communication between mitochondria and the endolysosomal system. Physical contacts between lysosomes and mitochondria are a recent discovery due to the advances in microscopy techniques and molecular biology, and they have mainly been studied in yeast. Beside these physical contacts, mitochondrial derived vesicles (MDVs) represent another means of achieving molecular transfer between mitochondria and the endolysosomal system. Although different modes of communication between these organelles have been identified, their biological meaning is often not yet clear.

### MITOCHONDRIA AND VACUOLE INTERACTIONS IN YEAST

Yeast has been a powerful model to study the connections between organelles. The vacuole is the equivalent of the lysosome in yeast, and the first description of a functional connection between the vacuole and mitochondria supported a role in lipid metabolism. Indeed, altered cardiolipin synthesis caused vacuolar damage (Chen et al., 2008). Cardiolipin is an anionic phospholipid predominantly found in the mitochondrial inner membrane and its dissociation from mitochondria leads to mitochondrial dysfunction. Deleting cardiolipin synthase, the enzyme responsible for cardiolipin synthesis in mitochondria, causes several defects in vacuoles and it affects cell survival (Chen et al., 2008). Yeast carrying a mutation in cardiolipin synthase have defective vacuolar morphology, decreased v-ATPase activity and vacuolar basification. Hence, cardiolipin plays a key role in the functional crosstalk between mitochondria and vacuoles. Although the mechanism underlying this crosstalk is not clear, it seems to involve a perturbation of ion homeostasis since deletion of the Na+/H+ exchanger ameliorated the vacuolar defects.

Another functional correlation between the vacuole and mitochondria was discovered during yeast aging. Aged yeast exhibit mitochondrial fragmentation, loss of mitochondrial DNA in their progeny and increased levels of mitochondrial ROS (Lam et al., 2011; McFaline-Figueroa et al., 2011). Two genes have been identified that are related to aging: VMA1, encoding a subunit of the v-ATPase, and VPH2, an integral membrane protein of the ER required for v-ATPase assembly. Overexpression of VMA1 and VPH2 decreases the vacuolar pH, delaying the mitochondrial dysfunction associated with aging. This pro-longevity effect is related to the capacity of the vacuole to store neutral amino acids rather than to its degradative function (Hughes and Gottschling, 2012). Reduced vacuolar acidity in aged yeast also remodels the mitochondrial proteome (Hughes and Gottschling, 2012). Upon loss of vacuolar acidity, specific membrane proteins from mitochondrial membranes are sorted into a mitochondrial-derived compartment (MDC) (**Figure 1**). This membrane compartment is then released by fission and transported to the vacuole for protein degradation by autophagy. Proteins are selectively incorporated into the MDC and this requires the mitochondrial import receptors Tom70 and Tom71. This mechanism represents a protective pathway to preserve mitochondrial integrity in times of stress (Hughes et al., 2016).

Two parallel studies demonstrated that physical contacts exist between mitochondria and the yeast vacuole, known as the vacuole and mitochondria patch (vCLAMP) (**Figure 1**). By screening yeast mutants, two genes were found that affect the formation of the physical contact between the ER and mitochondria, the endoplasmic reticulum-mitochondria encounter structure (ERMES) (Elbaz-Alon et al., 2014). Deletion of the two proteins encoded by these genes, Vps39 and Vam7, that are involved in fusion at the vacuole (Ungermann, 2015), causes an increase in the number of ERMES, although they do not act directly on these structures. Vps39 was previously identified as part of the homotypic fusion and protein sorting (HOPS) tethering complex and it was localized to contact sites between the vacuole and mitochondria (Elbaz-Alon et al., 2014). In a parallel study, physical contacts between mitochondria and vacuoles were discovered by electron microscopy and while Vps39 was implicated in this interaction, Ypt7 was identified in the vacuolar part of the contact (Hönscher et al., 2014) (**Figure 1**). In both cases, a relationship between ERMES and vCLAMP was seen to exist, as confirmed by an increase in the number of ERMES when the vCLAMP is impaired. Conversely, there are more vCLAMPs when ERMES are disrupted. Notably, yeast mutants for both mitochondrial contacts are lethal, demonstrating that both vacuolar and ER contacts with mitochondria contribute to mitochondrial function, possibly by carrying out partially redundant functions (Elbaz-Alon et al., 2014; Hönscher et al., 2014). These connections are necessary to mediate the transport of phospholipids between the mitochondria and endomembrane system during phospholipid

synthesis, a process that requires both organelles. Moreover, vCLAMPs are also enriched for specific ion and amino acid transporters, such that the activity of vCLAMPs may extend beyond lipid transport to nutrient sensing and utilization (Elbaz-Alon et al., 2014). Interestingly, the formation of these contacts depends on the metabolism of the cell. Yeast grown on glucose have more contact sites between mitochondria and vacuoles, and this number is reduced when they are grown in glycerol (Hönscher et al., 2014). Hence, ERMES and vCLAMPs are regulated in response to the metabolic status of cells, suggesting

derived compartment; vCLAMP, vacuole and mitochondria patch; ERMES, endoplasmic reticulum-mitochondria encounter structure; (?), unidentified

mitochondrial component of vCLAMP.

that both contacts are implicated in similar activities but under different conditions, i.e., respiratory vs. fermentative metabolism.

The coordination of these different contact sites is further revealed by studying Lam6, a novel protein that regulates both contact sites in yeast (Elbaz-Alon et al., 2015). Lam6 is present at both ERMES and vCLAMP contact sites (**Figure 1**), although it is not essential for the formation of these contacts and its absence does not affect their integrity. Rather, Lam6 can extend the contacts between organelles and indeed, vCLAMP is extended in an ERMES mutant but this enlargement does not occur when Lam6 is deleted. Likewise, there are no more ERMES when vCLAMP is impaired in the absence of Lam6, proof of that Lam6 mediates the crosstalk between different organelle contacts in yeast, and in particular that of ERMES and vCLAMP (Elbaz-Alon et al., 2015).

### MITOCHONDRIA-ENDOLYSOSOMAL SYSTEM CROSSTALK IN MAMMALS

The molecular mechanisms mediating the communication between mitochondria and lysosomes in mammalian cells remain unclear. An initial mode of communication was discovered when mitochondria were shown to form specific vesicles targeted to the endolysomal system (Soubannier et al., 2012). The protein cargo of MDVs is selectively incorporated, and it may be limited to elements at the outer membrane or it might also include inner membrane and matrix components (**Figure 2**). The molecular mechanism that drives MDVs formation is different from mitochondrial fission, since MDVs are formed in Drp1-silenced cells, Drp1 being a key protein in mitochondrial fission (Soubannier et al., 2012). Parkin induces the PINK1-dependent formation of MDVs under conditions of mitochondrial stress, which are enriched for mitochondrial specific proteins like Tom20 that are targeted to endolysosomal system for degradation (McLelland et al., 2014). These MDVs are originally targeted to the MVB and they finally fuse with lysosome. Such vesicle formation may be a rapid response to mitochondrial stress, shuttling oxidized cargo to lysosomes in order to preserve the integrity of the organelle (McLelland et al., 2014; Sugiura et al., 2014). MDVs have also been reported to be involved in mitochondrial quality control upon acute stress in the cardiac system (Cadete et al., 2016).

Autophagy is the process of removing cellular components through their encapsulation in a double-membrane structure followed by degradation in lysosomes. In particular, the selective removal of damaged mitochondria by autophagy is known as mitophagy. Mitophagy, like MDVs formation, is parkin and PINK1 dependent (Youle and Narendra, 2011; Roberts et al., 2016). However, MDVs are also formed when essential components of autophagy are silenced like Atg5, Beclin-1 or Rab9. Moreover, MDVs do not co-localize with LC3, an autophagosome marker, demonstrating that they are independent of a canonical autophagy process (Soubannier et al., 2012; McLelland et al., 2014). Both MDVs and mitophagy are quality control mechanisms to handle damaged mitochondria. Nevertheless, given that MDVs formation is a faster and less drastic process and it is independent of the classical mitophagy machinery, it could be the first response to mitochondrial stress preceding mitophagy (McLelland et al., 2014; Sugiura et al., 2014).

Mitochondrial dysfunction provokes lysosome impairment, representing a functional connection between the mitochondria and lysosome (**Figure 2**). Deletion of mitochondrial transcription factor A (TFAM) has been used as a model of mitochondrial dysfunction (Vernochet et al., 2012; Viader et al., 2013), as TFAM is essential for the replication and transcription of mitochondrial DNA (mtDNA) (Larsson et al., 1998). In the absence of TFAM, there is less mtDNA, fewer transcripts encoding mtDNA genes and an impairment of the electron transport chain (Gustafsson et al., 2016). Upon deletion of TFAM in T cells, the impairment of mitochondria augments the number of lysosomes. However, lysosomal activity is profoundly impaired, provoking an accumulation of sphingomyelin and autophagy intermediates, and triggering inflammatory responses (Baixauli et al., 2015). The increased lysosomal mass was caused by the activation of the transcription factor TFEB (Transcription Factor EB), a master transcription factor controlling lysosome biogenesis. Increasing the cellular levels of NAD+ by addition of a NAD+ precursor improved lysosomal function in this model, evidence that the reduced NAD+ levels in cells with impaired mitochondrial function causes lysosome dysfunction. Indeed, disturbing mitochondrial function in neurons by genetic deletion of AIF, OPA1 or PINK1, or with specific mitochondrial inhibitors, provokes morphological alterations to lysosomes (Demers-Lamarche et al., 2016). Upon such mitochondrial alterations, lysosomes become larger and they accumulate lysosomal vacuoles, although neurons appear healthy and viable. A higher lysosomal pH, dampened activity of lysosomal enzymes and an accumulation of substrates of autophagy were also registered in these conditions. Antioxidant treatment partially rescued these lysosomal defects, suggesting that excess of mitochondrial ROS triggers lysosome dysfunction (Demers-Lamarche et al., 2016).

More recently, another functional connection between mitochondria and lysosome was discovered. Pharmacologically impaired mitochondria affect lysosome biogenesis, enhancing the expression of certain lysosomal proteins: Lamp1, a lysosomal membrane protein; and lysosomal enzymes for degradation, such as acid alpha-glucosidase and cathepsins. This effect on lysosome activity is dependent on the duration of the mitochondrial damage. Accordingly, some lysosomal genes are upregulated after 1 week of treatment but their expression is downregulated after 4 weeks of treatment, a pattern that is also observed in fibroblasts obtained from patients with complex I deficiency and in mouse embryonic fibroblasts from a mouse model of mitochondrial malfunction (Fernández-Mosquera et al., 2017). The induction of lysosomal biogenesis in early mitochondrial damage is TFEBdependent and it requires AMPK signaling, representing a response to mitochondrial stress, increased autophagy flux and the capacity to remove damaged mitochondria (Fernández-Mosquera et al., 2017).

Besides this functional and vesicular connection, direct interorganelle contacts between mitochondria and the

the mitochondria-melanosome physical connection. MVB, Multivesicular body; MDVs, Mitochondrial-derived vesicles; PINK1, PTEN-induced kinase 1; VDAC1, mitochondrial voltage-dependent anion channel isoform 1; Mfn2, Mitofusin2; TFEB, Transcription Factor EB.

endolysosomal compartment have been identified in mammals, specifically in hypoxic cells and erythrocytes (Sheftel et al., 2007; Hamdi et al., 2016). In conditions of hypoxia, mitochondria are hyperfused and they establish physical contacts with late endolysosomes (**Figure 2**). Such interorganelle contact may be important for the cleavage of VDAC1 by a cysteine protease mainly located in endolysosomes. Cleaved VDCA1 has a different conformation, which protects the mitochondria from autophagy and increases metabolic efficiency (Brahimi-Horn et al., 2015). In erythrocytes, interactions between the mitochondria and late endosomes facilitate the transport of iron between these two organelles (Sheftel et al., 2007; Hamdi et al., 2016).

## PHYSICAL CONNECTIONS BETWEEN MITOCHONDRIA AND MELANOSOMES

Lysosome related organelles (LROs) represent a heterogeneous set of organelles that possess some features common to lysosomes but that are cell type-specific. Consistent with their distinct functions, LROs vary in composition and morphology, and they can be distinguished by the primary source of their membrane and their intralumenal content. Some of the LRO's contents are derived from the endolysosomal system, and most of them contain lysosomal proteins and have a low lumenal pH (Raposo et al., 2007).

Melanosomes are LROs of pigmented cells devoted to the synthesis, storage and transport of melanins. Melanosomes are considered biological models of organelle biogenesis and motility. They originate from endosomal precursors and subsequently, they mature through the progressive deposition of melanin and they are transported toward the cell periphery (Raposo et al., 2001). Contacts between mitochondria and melanosomes are visible by electron microscopy and tomography, and these connections affect ca. 1% of melanosomes (Daniele et al., 2014). The formation of these contacts is not based on membrane fusion but on the establishment of fibrillar bridges between the two organelles (**Figure 2**). Mitofusin 2 is thought to play an essential role in these contacts, a surprising new function of this protein that was originally identified as part of MAMs (de Brito and Scorrano, 2008). Mitochondriamelanosome contacts are more abundant in the perinuclear area where new melanosomes are generated and, indeed, stimulating melanosome biogenesis was found to enhance the presence of these contacts. Moreover, these contacts could mediate the ATP synthesis necessary for the maturation and acidification of melanosomes, as well as influencing other events in melanosomes, like melanin synthesis and the exchange of small molecules. Melanosomes are considered acidic calcium stores, indicating that they may also be important in calcium signaling. Due to the antioxidant and free radical scavenging roles of melanin, this connection might be involved in controlling the redox-status of melanocytes.

### MITOCHONDRIAL MOLECULES ARE SECRETED TO THE EXTRACELLULAR MEDIA BY EXOSOMES

Exosomes are extracellular vesicles that are secreted by all cell types and that are derived from MVBs upon their fusion with the plasma membrane. The presence of mitochondrial molecules in exosomes represents further indirect evidence of the crosstalk between mitochondria and the endolysosomal system (Torralba et al., 2016). Before being exported to the extracellular milieu, toxic, obsolete or damaged mitochondrial material is loaded into endolysosomes for degradation or extracellular export. Exosomes contain genetic material, mostly non-coding RNA, but also mtDNA (Guescini et al., 2010a,b) (**Figure 2**). Moreover, mesenchymal stem cells and astrocytes can produce even larger vesicles that contain entire mitochondrial particles and mtDNA (Phinney et al., 2015; Hayakawa et al., 2016). The physiological function behind the transport of mitochondrial DNA and proteins out of the cell remains unclear, as do the molecular events responsible for their loading into vesicles. MDVs or direct interorganelle contacts could represent a rapid mechanism to

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## CONCLUDING REMARKS

The study of all facets of interorganelle connections is possible due to new super-resolution imaging and molecular biology techniques, approaches that have shed more light on this emerging and promising field of research. The crosstalk and dependency between organelles could be fundamental to better relate currently misunderstood cell processes. Indeed, secondary effects on related organelles are important in disease pathophysiology and they may represent a starting point to identify novel therapeutic targets to treat human disease. This is especially critical in the case of mitochondrial and lysosomal diseases, devastating pathologies that mostly represent unmet medical needs. Mitochondria and lysosomes (or LROs) interact at different levels and in distinct ways to control cell homeostasis, and these connections are essential for the correct functioning of both organelles. In mammals, communication between organelles has mainly been described at a functional level, observed as secondary damage to lysosomes or LROs when mitochondria are altered, particularly when cells are under stress. Due to the role of lysosomes in degradation, this communication could be a rapid way to firstly reduce mitochondrial stress and preserve mitochondrial integrity. Although physical contact between mitochondria and the endolysosomal compartment has been mainly studied in yeast, recent evidence suggests that this is preserved in mammals. The connection between these organelles could coordinate processes that require the participation of both organelles, such as metabolic adaptation, phospholipid metabolism, regulation of calcium signaling and the control of cellular homeostasis.

## AUTHOR CONTRIBUTIONS

All authors listed have made a substantial, direct and intellectual contribution to the work, and approved it for publication.

## ACKNOWLEDGMENTS

We thank to G. Desdín-Micó and Dr. E. Carrasco for helpful ideas and comments on the manuscript. This work is supported by the research grant CP 14/00219 from Fondo de Investigación Sanitaria del Instituto de Salud Carlos III, Fondo Europeo de Desarrollo Regional (FEDER), the European Research Council (ERC-2016-StG 715322-EndoMitTalk) and the Instituto de Salud Carlos III (FIS16/188).

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**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2017 Soto-Heredero, Baixauli and Mittelbrunn. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

#### Edited by:

Nuno Raimundo, Universitätsmedizin Göttingen, Germany

#### Reviewed by:

Giovanna Cenini, University of Bonn, Germany Luke Esposito, Allen Institute for Brain Science, United States

#### \*Correspondence:

Serena Stanga<EMAIL_ADDRESS>Pascal Kienlen-Campard<EMAIL_ADDRESS>
#### † Present Address:

Paolo E. Porporato, Department of Molecular Biotechnology and Health Sciences, Molecular Biotechnology Center, University of Torino, Torino, Italy; Matthew Bird, Department of Clinical and Experimental Medicine, Hepatology, KU Leuven, Leuven, Belgium; Claudia Marinangeli, UMR-S 1172-JPArc-Centre de Recherche Jean-Pierre AUBERT, CHU Lille, Institut National de la Santé et de la Recherche Médicale, Université de Lille, Lille, France

> ‡ These authors have contributed equally to this work.

#### Specialty section:

This article was submitted to Mitochondrial Research, a section of the journal Frontiers in Physiology

Received: 26 July 2017 Accepted: 28 September 2017 Published: 12 October 2017

#### Citation:

Contino S, Porporato PE, Bird M, Marinangeli C, Opsomer R, Sonveaux P, Bontemps F, Dewachter I, Octave J-N, Bertrand L, Stanga S and Kienlen-Campard P (2017) Presenilin 2-Dependent Maintenance of Mitochondrial Oxidative Capacity and Morphology. Front. Physiol. 8:796. doi: 10.3389/fphys.2017.00796

## Presenilin 2-Dependent Maintenance of Mitochondrial Oxidative Capacity and Morphology

Sabrina Contino1 , Paolo E. Porporato2†, Matthew Bird1†, Claudia Marinangeli 1† , Rémi Opsomer 1 , Pierre Sonveaux 2 , Françoise Bontemps 3 , Ilse Dewachter 1 , Jean-Noël Octave1 , Luc Bertrand4 , Serena Stanga1 \* ‡ and Pascal Kienlen-Campard1 \* ‡

1 Alzheimer Research Group, Institute of Neuroscience, Université catholique de Louvain, Brussels, Belgium, 2 Pharmacology and Therapeutics, Institute of Experimental and Clinical Research, Université catholique de Louvain, Brussels, Belgium, 3 Metabolic Research Group, de Duve Institute, Université catholique de Louvain, Brussels, Belgium, 4 Pole of Cardiovascular Research, Institute of Experimental and clinical Research, Université catholique de Louvain, Brussels, Belgium

Mitochondrial dysfunction plays a pivotal role in the progression of Alzheimer's disease (AD), and yet the mechanisms underlying the impairment of mitochondrial function in AD remain elusive. Recent evidence suggested a role for Presenilins (PS1 or PS2) in mitochondrial function. Mutations of PSs, the catalytic subunits of the γ-secretase complex, are responsible for the majority of inherited AD cases (FAD). PSs were shown to be present in mitochondria and particularly enriched in mitochondria-associated membranes (MAM), where PS2 is involved in the calcium shuttling between mitochondria and the endoplasmic reticulum (ER). We investigated the precise contribution of PS1 and PS2 to the bioenergetics of the cell and to mitochondrial morphology in cell lines derived from wild type (PS+/+), PS1/2 double knock-out (PSdKO), PS2KO and PS1KO embryos. Our results showed a significant impairment in the respiratory capacity of PSdKO and PS2KO cells with reduction of basal oxygen consumption, oxygen utilization dedicated to ATP production and spare respiratory capacity. In line with these functional defects, we found a decrease in the expression of subunits responsible for mitochondrial oxidative phosphorylation (OXPHOS) associated with an altered morphology of the mitochondrial cristae. This OXPHOS disruption was accompanied by a reduction of the NAD+/NADH ratio. Still, neither ADP/ATP ratio nor mitochondrial membrane potential (19) were affected, suggesting the existence of a compensatory mechanism for energetic balance. We observed indeed an increase in glycolytic flux in PSdKO and PS2KO cells. All these effects were truly dependent on PS2 since its stable re-expression in a PS2KO background led to a complete restoration of the parameters impaired in the absence of PS2. Our data clearly demonstrate here the crucial role of PS2 in mitochondrial function and cellular bioenergetics, pointing toward its peculiar role in the formation and integrity of the electron transport chain.

Keywords: presenilin, mitochondria, glycolysis, oxidative phosphorylation, cellular bioenergetics, Alzheimer's disease

## INTRODUCTION

Metabolic dysfunction is central in Alzheimer's disease (AD) since it appears at very early stage of the disorder, even before clinical symptoms (Chen and Zhong, 2013). It is evidenced in patients by a decrease in glucose utilization in temporoparietal association areas, together with cognitive decline and a severe failure in mitochondria oxidative metabolism (Herholz, 2012). Mitochondria are known as the powerhouse of the cell for their capacity to supply energy, but they are also critical in other cellular processes such as apoptosis, reactive oxygen species (ROS) production (Paradies et al., 2014), and calcium homeostasis (Osellame et al., 2012). All these processes turn out to be affected in AD pathology (Hroudova et al., 2014). Studies carried out either on AD patients' tissue samples or AD transgenic mice models reported an array of mitochondrial dysfunctions, including a morphological shift toward fission (Wang et al., 2009), a disrupted motility (Wang et al., 2009; Xie et al., 2013), an impairment of the electron transport chain (ETC) (Bosetti et al., 2002), and an increase in ROS production with deleterious effects on mitochondrial DNA integrity (Onyango et al., 2006). In cytoplasmic hybrid (cybrid) cell lines, generated by insertion of platelets' mitochondria collected from sporadic AD (SAD) patients into human neuroblastoma cells (SHSY5Y) depleted of mitochondria, bioenergetics dysfunctions such as oxidative phosphorylation (OXPHOS) and glucose utilization defects have been observed (Silva et al., 2013). These metabolic perturbations found in AD raised a chicken and the egg issue, namely to define whether mitochondrial dysfunction is a cause or a consequence in this pathology.

Recent studies suggested that Presenilins are involved in the control of mitochondrial functions (Behbahani et al., 2006; Filadi et al., 2016). PS1 and PS2 are two homologous polytopic aspartyl proteases identified as the catalytic subunits of the γ-secretase complex. PSs are directly involved in AD since the production of Aβ is generated after the sequential cleavage of the Amyloid Precursor Protein (APP) by the β and the γ-secretase (Zheng and Koo, 2011). Aβ is the major component of senile plaques found in the brain of AD patients (Miller et al., 1993). Beside their key role in γ-secretase activity and Aβ production, mounting evidence indicate additional roles for PSs in cell physiology, for which their catalytic activity seems less evident. PSs are involved in calcium homeostasis (Zhang et al., 2013), autophagy (Lee et al., 2010), neurotransmitters release (Zhang et al., 2009), synaptic plasticity and memory (Saura et al., 2004). Moreover, although PSs have different localization in the cell with PS1 mainly located at the plasma membrane and PS2 in the trans-Golgi network and endosomal compartments (Meckler and Checler, 2016; Sannerud et al., 2016), they are both enriched in the mitochondriaassociated membranes (MAM) (Area-Gomez et al., 2009). MAM are specific membrane domains connecting endoplasmic reticulum (ER) and mitochondria. They are involved in lipid metabolism, calcium and cholesterol homeostasis (van Vliet et al., 2014; Filadi et al., 2017). PS2 has been reported to regulate the formation of ER/mitochondria contacts (Filadi et al., 2016) and calcium cross-talk between these two organelles (Zampese et al., 2011a). It is hence of particular interest to define (i) the precise contribution of PSs to the multiple aspects of mitochondrial function and (ii) to understand the respective contribution of PS1 or PS2 to these processes. Unraveling the role of PSs in cell metabolism is crucial to get insight in their physiological function and to understand how PS gain- or loss-of-function can create a pathological context, related for instance to AD. We performed an array of metabolic measurements on Mouse Embryonic Fibroblasts cell lines (MEFs) derived either from wild type (PS+/+), PS1/2 double knock-out (PSdKO), PS2KO, and PS1KO mice embryos. We found a key role for PS2 (and not PS1) in cell metabolism and especially in OXPHOS and glycolysis. Indeed, the absence of PS2 altered the OXPHOS capacity and integrity but increased glycolytic flux to support energy needs. These pathways are crucial for energy-intensive consumer cells, like brain cells. Such defects might set the basis for further investigation of the metabolic impairments observed in AD.

## MATERIALS AND METHODS

### Cell Lines and Cell Culture

Mouse Embryonic Fibroblasts cell lines (MEFs) derived from wild type (PS+/+), PS1/2 double knock-out (PSdKO), PS2KO and PS1KO mice embryos were previously described (Hebert et al., 2006; Marinangeli et al., 2015).  Cells were maintained in DMEM low glucose (5.5 mM) (Sigma-Aldrich, St Louis, USA) supplemented with penicillin/streptomycin solution (10 units–10 µg) and 10% fetal bovine serum (FBS) (Thermo Scientific, Rockford, USA). MEF 1R1 and 2R2 cell lines were generated by subcloning the human PSs cDNA sequences in the lentiviral backbone vectors plentiCMV/TOpuro and pTMtm898neo, respectively. Lentiviruses were produced in human embryonic kidney 293T cells. Supernatants containing the lentiviruses were concentrated with the Lenti-X Concentrator kit according to the manufacturer's instructions (Clontech Laboratories; California, USA). MEFs PS1KO and PS2KO were transduced by lentiviral particles followed by puromycin (2.5µg/ml) selection (1R1) and G418 (500µg/ml) selection (2R2), respectively. Stable expression of human PSs was monitored by western blotting (WB).

### Western Blotting

WB was performed on cell lysates as previously described (Stanga et al., 2016). Primary antibodies used were Anti-OXPHOS

**Abbreviations:** 19, mitochondrial membrane potential; 1R1, PS1 knockout rescued PS1; 2R2, PS2 knockout rescued PS2; AD, Alzheimer disease; APP, amyloid precursor protein; CI-CV, complexes I-V; DTAB, dodecyltrimethylammonium bromide; ETC, electron transport chain; ER, endoplasmic reticulum; FAD, familial AD; FC, flow cytometry; FCCP, carbonyl cyanide-4-(trifluoromethoxy) phenylhydrazone; HPLC, high pressure liquid chromatography; MAM, mitochondria-associated membranes; MEF, mouse embryonic fibroblast; OCR, oxygen consumption rate; OXPHOS, oxidative phosphorylation; PS, presenilin; ROS, reactive oxygen species; SAD, sporadic AD; TEM, transmission electron microscopy; TMRM, tetramethylrhodamine methyl ester; WB, western blotting.

Cocktail (1:1,000; Abcam, Cambridge, United Kingdom); Anti-TOM20 (1:1,000; Proteintech, Rosemont, USA); anti-Presenilin (1:1,000; Cell Signaling, Danvers, USA); anti-Actin (1:3,000; Abcam, Cambridge, United Kingdom). Secondary antibodies used were anti-mouse (1:10,000; GE Healthcare, Little Chalfont, United Kingdom) and anti-rabbit (1:10,000; GE Healthcare, Little Chalfont, United Kingdom).

### Metabolic Measurements

Oxygen consumption rate (OCR) measurements were performed with a Seahorse XF96 bioenergetic analyzer (Seahorse Bioscience; Massachusetts, USA). Cells were plated at 80% cell confluence onto a Seahorse 96 well plates 24 h before the assay. According to manufacturer's instructions, cell medium was replaced by the conditional medium (culture medium without FBS and sodium bicarbonate) and incubated at 37◦C without CO2 for 1 h before completion of probe cartridge calibration. Inhibitors targeting the different mitochondrial complexes (Cell Mito Stress Test kit, Seahorse Bioscience) have been added sequentially to the cells during the experiment to measure the basal respiration, the coupling and the spare respiratory capacity: Oligomycin (1 µM); FCCP (1µM); Rotenone and antimycin A (0.5µM). Results were normalized to the total amount of protein measured by the Bradford assay kit (Bio-Rad Laboratories, California, USA).

### Mitochondrial Membrane Potential (1ψ)

Fluorescent cationic probe tetramethylrhodamine methyl ester (TMRM) (Sigma-Aldrich, St Louis, USA) was used to evaluate the 19. The uncoupling agent Carbonyl cyanide-4- (trifluoromethoxy) phenylhydrazone (FCCP) served as control (Sigma-Aldrich, St Louis, USA). Cyclosporin H (2µM; Abcam, Cambridge, United Kingdom), an inhibitor of multidrug resistance pump activity was used to limit the efflux of TMRM. MEFs were incubated for 30 min at 37◦C with TMRM (30 nM), cyclosporine H with or without FCCP (10µM) diluted in KREBS medium. For the flow cytometry (FC) analysis we trypsinized cells seeded in 6 well plates and harvested them in PBS/5% FBS. After centrifugation at 300 g for 5 min, cells were resuspended in KREBS medium for the FC analysis performed on a BD FACSCantoTM flow cytometer (Biosciences; San Jose, USA). Data were analyzed with the FlowJo software (FlowJo,LLC; Oregon, USA).

### Adenine Nucleotides Analysis

24 h after seeding in 10 cm culture dishes, cells were harvested and lysed for 30 min on ice with HCLO4 1N (Sigma-Aldrich, St Louis, USA) and then centrifuged for 20 min at 10,000 g at 4 ◦C. The pellets were re-suspended in NaOH 0.5 M for protein quantification and the supernatants were adjusted to pH 6–8 using a solution of KOH/KHCO3 3 M (Sigma-Aldrich, St Louis, USA). Precipitated salts were separated from the liquid phase by centrifugation at 10,000 g at 4◦C for 20 min. Samples were stored at −80◦C. Nucleotides were separated by high-performance liquid chromatography (HPLC) on 125 × 4.6-mm PartiSphere 5 SAX anion-exchange column (Whatman, Maidstone, UK). Nucleotides were separated with a gradient from 100% buffer A (0.01 M NH4H2PO4, pH 3.7) to 100% buffer B (0.48 M NH4H2PO4, pH 3.7) over 27 min at a flow rate of 2 ml/min according to the method of Hartwick and Brown (1975). UV detection of nucleotides was performed at 254 nm. Quantification of nucleotides was achieved by peak integration of the area under the curve, validated by the use of external standards (ADP and ATP).

## NAD+/NADH Ratio

Cells were seeded in 96 well plates at 80% confluence 24 h before the assay. NAD+/NADH ratio was measured by using the bioluminescent NAD+/NADH-GloTM assay kit (Promega, Wisconsin, USA) according to the manufacturer's instructions. Briefly, total NAD+ and NADH were extracted from cell pellets with the basic solution 1% dodecyltrimethylammonium bromide (DTAB). Samples were divided in two for both acid and basic treatments and heated at 60◦C for 15 min. The oxidized form is selectively decomposed in the basic solution while the reduced form is decomposed in the acidic solution. For the luminescent reaction, samples were mixed with 100 µl of NAD+/NADH-GloTM detection reagent and incubated for 45 min before reading on the GloMax R 96-well plate luminometer (Promega, Wisconsin, USA).

### Glucose and Lactate Measurements

Cells were seeded at 80% confluence in 6 well plates. 24 h after, 500 µl of medium per well was collected and deproteinized. Glucose and lactate concentrations were measured using specific enzymatic assays on a CMA600 microdialysis analyzer (CMA Microdialysis AB, Solna, Sweden). Cells were collected for protein quantification and data were normalized to the amount of protein measured by BCA assay (Thermo Scientific, Rockford, USA).

### Glycolytic Flux Measurement

Glycolytic rate was evaluated by measurement of the detritiation rate of [3-3H] glucose. Briefly, tritiated glucose (0.2 µCi/ml; Perkin-Elmer; Massachussets, USA) was added to the medium (including 5.5 mM glucose) for 30 min. After medium removal, the tritiated water resulting from detritiated glucose was separated from the non-transported tritiated glucose by column chromatography and measured with the Tri Carb 2,810 liquid scintillation analyzer (Perkin Elmer; Massachussets, USA) as described previously (Marsin et al., 2002). Data were normalized to the amount of protein measured by BCA assay (Thermo Scientific, Rockford, USA).

### Complex I (CI) Enzyme Activity Assay

The activity of complex I of the mitochondrial respiratory chain was evaluated with the Complex I Enzyme Activity Dipstick Assay kit (Abcam, Cambridge, United Kingdom). Cells were scraped in PBS and centrifuged for 5 min at 4◦C at 500 g. 10 volumes of extraction buffer were added to the pellets prior to 20 min of incubation on ice. After centrifugation, pellets were discarded and supernatants were used to determine protein concentration (BCA assay). Samples (corresponding to 30 µg of proteins) were added to a microplate with blocking buffer. The dipsticks (containing an antibody capturing the CI) were immersed in the samples to capture the CI. The NADH and NBT were added to allow the oxidation of NADH by the complex I, which in turn reduce NBT to form a purple precipitate. Colorimetric signals were quantified with the Gel Doc 2,000 coupled with Quantity One software (Bio-Rad; California, USA).

### Transmission Electron Microscopy (TEM)

Cells were fixed with 2.5% glutaraldehyde in phosphate buffer and kept in the fixative during 1 h at room temperature. They were washed and postfixed with 1% osmium tetroxide in the same buffer containing 0.8% potassium ferricyanide at 4◦C. The samples were dehydrated in acetone, infiltrated with Epon resin during 2 days, embedded in the same resin and polymerised at 60◦C during 48 h. Ultrathin sections were obtained using a Leica Ultracut UC6 ultramicrotome (Leica Microsystems, Vienna) and mounted on Formvar-coated copper grids. They were stained with 2% uranyl acetate in water and lead citrate. Sections were observed in a Tecnai Spirit electron microscope equipped with an Eagle CCD camera (FEI, Eindhoven, The Netherlands).

### Statistical Analysis

The number of experiments for each experimental condition is indicated in the figure legends. Data were analyzed using GraphPad Prism software (GraphPad Software, La Jolla, CA, USA) by ANOVA followed by Bonferroni's multiple comparison tests. ∗p < 0.05, ∗∗p < 0.01, ∗∗∗p < 0.001.

## RESULTS

### Absence of PS2 Results in Decreased OXPHOS without Alteration of the 19

PSs expression profile was measured by WB in MEF cell lines with antibodies directed against PS1 and PS2 CTFs (**Figures 1A,B**). The expression profile indicated the restoration of PSs expression in 2R2 (PS2 KO stably expressing human PS2) and 1R1 (PS1KO stably expressing human PS1). We evaluated the OXPHOS capacity by measurement of the OCR. The overall profile of OCR and the parameters related to the OXPHOS activity measured before or after drug addition, that are basal respiration, coupling (oxygen consumption devoted to ATP synthesis under resting conditions) and spare respiratory capacity (maximal uncoupled rate of respiration minus the basal rate) were all impaired in PSdKO and PS2KO cells (**Figures 1C–F**). Stable re-expression of PS2 in the PS2KO background (2R2) restored all these parameters to the levels measured in control cells (PS+/+). Importantly, the absence of PS1 did not affect mitochondrial respiration. We next measured the ∆9 with the TMRM probe by FC (**Figure 2A**) and by fluorescence measurement in microplates (Figure S1A). FC analyzes were performed on gated homogenous populations that accounted for more than 90% of the total population (Figure S1B). FCCP, an uncoupling agent abolishing 19 was used as a control. In both approaches (FC and direct fluorescence measurements), no significant differences in 19 were observed between the different populations of MEFs, although oxygen consumption was altered in PS2KO and PSdKO cells.

### Increased Anaerobic Glycolysis Sustains the Energy Production in PSdKO and PS2KO MEFs

The observed PS2-dependent OXPHOS capacity decrease without impairment of 19 led us to evaluate the total ADP and ATP cellular levels. This was achieved by HPLC. No differences were observed in the ADP/ATP ratio between the cell lines (**Figure 2B**). Considering that we detected a significant decrease in coupling in PSdKO and PS2KO cells, we suspected that ATP production in PSdKO and PS2KO cells could result from an increased glycolysis (**Figures 2C,D**) that would counteract the defects observed in OXPHOS. We measured the levels of lactate secreted and glucose consumed by cells, as marker of glycolysis (**Figure 2C**). Both were indeed significantly higher in PSdKO and PS2KO cells. No changes were observed in PS1KO cells. Lactate secreted and glucose consumed measured in 2R2 cells were comparable to those measured in PS+/+ cells. These data were in line with the significant increase of the glycolytic flux (measured by detritiation of [3- 3H] glucose) observed in PSdKO and PS2KO cells, that was also restored in 2R2 to the levels measured in PS+/+ cells (**Figure 2D**).

### Absence of PS2 Impairs the ETC

So far, our results showed that mitochondrial oxidative capacity was impaired in the absence of PS2. This defect can be compensated by the increased glycolytic flux we observed. We further investigated the mitochondrial defects by profiling the expression of the five mitochondrial complexes. This was performed by WB with a cocktail of antibodies targeting representative subunits of the five mitochondrial complexes. Results showed that the expression of some complexes of the ETC was perturbed in PSdKO and PS2KO cells: CI and CII subunits were significantly decreased in both cell lines, and CIV was more specifically decreased in PS2KO cells. The expression profile of CI, CII, and CIV was restored in 2R2 cells (**Figures 3A,B**). To confirm these observations, we measured by an enzymatic assay the activity of the CI, which was significantly decreased in PSdKO and PS2KO cells but totally restored in 2R2 cells (**Figure 3C**). Since CI uses NADH as an electron donor, we assessed whether the redox state was also affected by measuring the NAD+/NADH ratio. We measured the NAD+/NADH ratio using a luminescence assay. Consistent with the measurement of CI subunits by WB, NAD+/NADH ratio was decreased in PSdKO and PS2KO cells and restored in 2R2 cells (**Figure 3D**).

### Mitochondrial Morphology in PSs-Deficient Cells

We measured by WB in cell lysates the expression level of the mitochondrial import receptor subunit TOM20 (**Figures 4A,B**) as a marker of mitochondrial mass (Whitaker-Menezes et al., 2011). No significant difference in TOM20 levels was observed between the different cell types suggesting a comparable mass of mitochondria in all the cell lines tested, including PSdKO and PS2KO cells. Mitochondrial mass is

a gross indicator that could be only hardly related to the changes in mitochondrial function we observed. Thus, general mitochondrial morphology was evaluated by immunofluorescent staining targeting TOM20 (Figure S2), and no significant changes were observed about the shape and distribution of the mitochondrial network. Morphology and structure of single mitochondria were evaluated by TEM. Strikingly, we observed defective cristae (**Figure 4C**) in PSdKO and PS2KO cells when compared to PS+/+ cells. Cristae were less defined and less numerous in MEFs PSdKO and PS2KO and, importantly, these morphological defects were not observed in PS1KO cells.

### DISCUSSION

The physiological function of the Presenilins 1 and 2 is far from being fully elucidated. Over the two last decades, the seminal observation identifying PS1 as the catalytic core of the γ-secretase (De Strooper et al., 1998; Kimberly et al., 2003) launched a considerable research effort aimed at understanding the exact role of PSs in γ-secretase activity and Aβ production. However, an increasing number of studies focused on γ-secretase independent PSs functions such as regulation of calcium homeostasis (Zhang et al., 2013) or neurotransmitter (glutamate) release (Zhang et al., 2009). Since PSs were shown to be enriched

experiments). (D). Glycolysis rate was determined by the detritation rate of [3-3H] glucose after a 30 min incubation. Data were normalized to protein content. \*p < 0.05, \*\*p < 0.01, \*\*\*p < 0.001 (n = 12 from 6 independent experiments).

(n = 6 from 3 independent experiments). (C). Glucose consumption (negative columns) and lactate release (positive columns) were measured in supernatant from fresh medium after 24 h of culture. Data were normalized to protein content and expressed as µmol/day/µg prot. \*p < 0.05, \*\*p < 0.01, (n = 16 from 6 independent

control. TOM20 quantification (B) (means ± sem) is given as the percentage of the signal measured in the PS+/+ cells (n = 6 independent experiments). (C) TEM's of cell section. Higher magnification regions (right columns) are boxed in white (left column). Red stars are indicating the position of the mitochondria on the micrograph. Scale bars are indicated on the bottom of the pictures.

in MAM, a domain involved in different pathways related to ER and mitochondria functions (van Vliet et al., 2014; Filadi et al., 2017), we investigated the potential role of PSs in the control of cellular metabolism by measuring an array of parameters related to mitochondrial activity in PSs-deficient MEF cell lines. We observed in the absence of PS2, alterations in OXPHOS capacity and integrity associated with a decrease NAD+/NADH ratio. Along with these functional defects, the integrity of the mitochondrial cristae was also affected. These defects occurring in absence of PS2 were compensated by an increased glycolysis. Mitochondrial function was fully restored by stable re-expression of PS2 in our model.

### PS2 in OXPHOS and 19

We measured the OCR as an indirect measurement of the OXPHOS. Our results indicate a significant decrease in PSdKO and PS2KO of the OCR. All the parameters we measured (basal respiration, coupling and spare respiratory capacity) were decreased in PSdKO and PS2KO cells, supporting a general defective OXPHOS capacity. One could have expected here an impact on the 19 and ATP production, since the ATP synthase uses the electrochemical gradient formed by pumping protons through the inner membrane to produce ATP. With the TMRM probe, we observed that the 19 measured is stable in all the cell types. Previous studies reported a 19 decrease in PSdKO and PS2 KO cells (Behbahani et al., 2006). This discrepancy could be due to different procedures as for instance the use of a different probe (JC-1) at high concentration (Perry et al., 2011). Behbahani et al. used like us FC analysis as a readout for 19, but it should be noted that in their case the use of the JC-1 probe to measure 19 changes implies a shift in the emission wavelength of the probe (with overlapping spectra) that renders the analysis by FC more difficult to interpret. With the TMRM probe, by using FC or direct fluorescent measurement in microplates (see Figure S1) as a readout we observed that the 19 is stable in all the cell types analyzed. 19 is also known to be one of the most stable parameter in the cell; even in the case of a deficit, the ATP synthase will work in reverse to keep it stable (Uechi et al., 2006).

To note, the main source of ROS production is the ETC, and their implication in AD could rise the question about their production in our model. We could expect a decrease of ROS production in PSdKO and PS2KO regarding of the OXPHOS deficit. A previous study showed a ROS increase in PSdKO compared to PS+/+, in serum deprivation conditions (Boo et al., 2009) that we do not modelize here. We performed few pilot ROS measurement (data not shown) in basal state but no significant message came up. It would be of interest, given the implication of ROS in AD, to further study the PSs-dependent and ROS production in relevant models.

### PS2 and ATP Production

Surprisingly, the ADP/ATP ratio we measured by HPLC was not impaired while the coupling (oxygen dedicated to ATP production) was significantly decreased in PSdKO and PS2KO cells. This is consistent with the increase of glycolytic flux and lactate production we observed in PSdKO and PS2KO cells. We used two distinct approaches to estimate glycolysis in our model: (i) we measured the levels of glucose consumed and lactate released in cell media; (ii) we measured the glycolysis rate by detritiation of [3-3H]glucose. These combined approaches showed that the glycolysis is increased significantly in PSdKO and PS2KO, providing the fuel to compensate their OXPHOS deficit. It should be noted that, similarly to undifferentiated stem cells (de Meester et al., 2014), MEF cell lines use anaerobic glycolysis even in the presence of oxygen to produce their energy. It would be of prime interest to analyze if such compensatory capacity exists in non-proliferative and oxidative cells like primary neurons. The metabolic phenotype resulting from altered PS2 function could be more drastic in neuronal cells. This would be very important in the context of the Alzheimer pathology, in which PSs play a central role. Zampese and collaborators highlighted in SH-SY5Y neuronal cells overexpressing a FAD PS2 mutant an enhanced Ca2+ transfer between ER and mitochondria (Zampese et al., 2011b). On the other hand, SH-SY5Y treated with siRNA specific for PS2 showed a decreased contact between mitochondria and ER, which induced a decrease in calcium flux. The calcium crosstalk is known to be an important messenger for Krebs cycle enzymes stimulation and kinetics of the mitochondrial complexes. The mitochondrial calcium sequestration capacity is also important in the excitotoxicity and mitochondrial oxidative stress related to AD. It is therefore possible that a lack of calcium flux could be one cause of the OXPHOS defect observed in our PS2KO cells.

### PS2 in ETC and Mitochondrial Network and Morphology

We found that the bioenergetics defects observed were neither due to a decrease in the mitochondrial mass nor associated with a compensatory induction in mitochondrial mass. This favors the hypothesis of a direct disruption of the ETC activity in absence of PS2 rather than an overall defect in mitochondria biogenesis. This would be interesting to further address in postmitotic cells, such as neurons, where one might expect a proliferation of mitochondria as an attempt to compensate for the OXPHOS deficits. Such a compensatory induction was seen in heart and muscle (and to a lesser extent in brain) of mice deficient in the adenine nucleotide translocator-1 (ANT1), on which OXPHOS depends since it provides the ADP substrate (Esposito et al., 1999)

In order to understand the origin of the OXPHOS defect, we assessed the expression profile of the 5 mitochondrial complexes (CI-CV) of the ETC with a cocktail of antibodies. Even if we did not evaluate the expression of the entire complexes by this approach, we found the expression of several complexes of the ETC (CI, CII, and CIV) to be decreased in PSdKO and PS2KO cells. This was consistent with a functional analysis indicating a decreased CI activity in the same cells. The most important electron donor produced by the Krebs cycle and used by the CI is NADH and NAD+ is an important co-factor regulating metabolic homeostasis (Canto et al., 2015). We therefore assessed their redox state by measuring the NAD+/NADH ratio and observed a significant decrease of the ratio in PSdKO and PS2KO cells. This can result either from an accumulation of NADH not used by the dysfunctional CI or from a decrease in NAD+

associated to metabolic pathways. NAD+ and NADH could also both be affected and synergistically influence pathways regulating metabolism homeostasis (Canto et al., 2015).

The perturbation of the integrity of the ETC in PSdKO and PS2KO correlates with an altered structure of their cristae as evidenced by TEM. Cristae are the seat of the ETC, it is thus logical that the OXPHOS capacity and complexes expression are affected when their structure is altered. It has been demonstrated that cristae shape could regulate respiratory chain supercomplexes assembly and stability, impacting the respiratory efficiency (Cogliati et al., 2013). One of the key regulators of cristae shape is OPA1 and it would be interesting to further study the relationship between PS2 and OPA1. Alternatively, considering MAM as an important compartment for the homeostasis of lipids such as for example cardiolipin, one of the most prominent lipids in cristae (Paradies et al., 2014; Ikon and Ryan, 2017), we suggest that the morphological deficit of the cristae could be related to a dysregulation of MAM function in absence of PS2. To note, the MAM hypothesis has been proposed in AD since all the pathways (cholesterol, lipid and calcium homeostasis, mitochondrial OXPHOS stimulation) controlled by this compartment are altered in the pathology (Area-Gomez and Schon, 2017).

In conclusion, we reported here a specific physiological role of PS2 in cellular metabolism. Indeed, absence of PS2 results in defective mitochondrial cristae correlating with an impaired OXPHOS capacity and a modified redox state (NAD+/NADH ratio). MEF cell lines, which are glycolytic cells, increase this glycolytic capacity to sustain their energy need in absence of PS2. This implies that impairment of these PS2-dependent processes could be involved in the progression of pathologies like AD in which PS2 play a key role. Investigation in an oxidative model and particularly in neurons are required to further explore this hypothesis.

### REFERENCES

### AUTHOR CONTRIBUTIONS

SC, SS, and PK-C designed the research study; SC and SS conducted experiments with fundamental input of PEP, MB, and LB; all the authors analyzed data. SC, SS, and PK-C wrote the paper; all the authors have read and approved the final manuscript.

### FUNDING

This work was supported by a grant of the Foundation for Research on Alzheimer's disease (PK-C), by the Interuniversity Attraction Pole Programme-Belgian Sate-Belgian Science Policy (IAP-P7/16 and IAP-P7/13) to J-NO, ID, and PK-C, by the Fondation Médicale Reine Elisabeth (FMRE) to J-NO, ID, and PK-C, by the Action de Recherche Concertée (ARC 14/19-059) to PK-C.

### ACKNOWLEDGMENTS

We are grateful to Bart de Strooper (KU Leuven, Belgium) for the kind gift of MEF cell lines, to Bernadette Tasiaux (Alzheimer Research Group, UC Louvain, Belgium) for the helpful technical assistance, to Carmen López Iglesias for her kind assistance and to the Microscopy Core Lab of the Maastricht Multimodal Molecular Imaging Institute (M4I), FHML, University of Maastricht. LB; ID and PS are Senior Research Associates of Fonds National de la Recherche Scientifique (Belgium).

### SUPPLEMENTARY MATERIAL

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fphys. 2017.00796/full#supplementary-material

by tuning the antagonistic effect of mitofusin 2. Cell Rep. 15, 2226–2238. doi: 10.1016/j.celrep.2016.05.013

memory and synaptic plasticity followed by age-dependent neurodegeneration. Neuron 42, 23–36. doi: 10.1016/S0896-6273(04)00182-5

**Conflict of Interest Statement:** The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.

Copyright © 2017 Contino, Porporato, Bird, Marinangeli, Opsomer, Sonveaux, Bontemps, Dewachter, Octave, Bertrand, Stanga and Kienlen-Campard. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) or licensor are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.

# Role of Cardiolipin in Mitochondrial Signaling Pathways

### Jan Dudek\*

Department of Cellular Biochemistry, University Medical Center Göttingen, Göttingen, Germany

The phospholipid cardiolipin (CL) is an essential constituent of mitochondrial membranes and plays a role in many mitochondrial processes, including respiration and energy conversion. Pathological changes in CL amount or species composition can have deleterious consequences for mitochondrial function and trigger the production of reactive oxygen species. Signaling networks monitor mitochondrial function and trigger an adequate cellular response. Here, we summarize the role of CL in cellular signaling pathways and focus on tissues with high-energy demand, like the heart. CL itself was recently identified as a precursor for the formation of lipid mediators. We highlight the concept of CL as a signaling platform. CL is exposed to the outer mitochondrial membrane upon mitochondrial stress and CL domains serve as a binding site in many cellular signaling events. During mitophagy, CL interacts with essential players of mitophagy like Beclin 1 and recruits the autophagic machinery by its interaction with LC3. Apoptotic signaling pathways require CL as a binding platform to recruit apoptotic factors such as tBid, Bax, caspase-8. CL required for the activation of the inflammasome and plays a role in inflammatory signaling. As changes in CL species composition has been observed in many diseases, the signaling pathways described here may play a general role in pathology.

### Edited by:

Anita Krisko, Mediterranean Institute for Life Sciences, Croatia

### Reviewed by:

Abdussalam Azem, Tel Aviv University, Israel Carsten Merkwirth, Ferring Research Institute, Inc., United States

\*Correspondence: Jan Dudek<EMAIL_ADDRESS>
### Specialty section:

This article was submitted to Mitochondrial Research, a section of the journal Frontiers in Cell and Developmental Biology

Received: 26 July 2017 Accepted: 21 September 2017 Published: 29 September 2017

#### Citation:

Dudek J (2017) Role of Cardiolipin in Mitochondrial Signaling Pathways. Front. Cell Dev. Biol. 5:90. doi: 10.3389/fcell.2017.00090 Keywords: Barth-Syndrome, cardiolipin, mitochondria, respiratory chain

### INTRODUCTION

The adult human body hydrolyses 64 kg adenosine triphosphate (ATP) per day. Mitochondria are the primary source of the energy in most tissues and their contribution to energy production is particularly important for tissues with high-energy demand, like neuronal or cardiac tissue. Besides their role in energy conversion, mitochondria have multiple functions in the metabolism, like the citric acid cycle, the urea cycle, the metabolism of amino acids and lipids. Mitochondria also play a role in the biogenesis of heme and iron-sulfur clusters (DiMauro and Schon, 2003). Dysfunctional mitochondria, which cannot provide the required energy particularly affect neuronal tissue and the heart (Wallace, 1999). Cardiomyopathies are frequently associated with defects in respiratory chain subunits and their assembly factors, but also with defects in mitochondrial translation and mtDNA maintenance (Schwarz et al., 2014).

 The inner membrane (IM) separates two compartments, the intermembrane space (IMS) and the matrix. The inner boundary membrane and the cristae structures are distinct functional areas of the IM. The inner boundary membrane is formed by segments of the IM, which approximate the OM in close apposition. The cristae structures are invaginations into the matrix and harbor the respiratory chain. The phospholipid cardiolipin (CL) is a hallmark lipid of mitochondria and almost exclusively found in mitochondrial membranes (Pangborn, 1945). CL is predominantly located in the inner membrane and associated to many mitochondrial functions (see below). Mitochondrial dysfunctions caused by changes in the CL pool are associated to a large number of cardiac diseases (Paradies et al., 2004; Han et al., 2005; Petrosillo et al., 2005; Sparagna et al., 2007; He and Han, 2014; Mulligan et al., 2014). Barth syndrome (BTHS) is an inherited cardiomyopathy, associated with skeletal myopathy, growth retardation and neutropenia and occurs at an estimated frequency of about 1 case per 300,000–400,000 births (Barth et al., 1996; Cantlay et al., 1999; Steward et al., 2010). BTHS is caused by a mutation in the TAZ gene, encoding for Tafazzin, a mitochondrial acyltransferase, involved in the biogenesis of CL (Bione et al., 1996; Dudek and Maack, 2017).

Signaling pathways monitor the physiological state of mitochondria, and trigger a cellular response to various stress conditions. In this review, we describe how alterations in the CL pool cause mitochondrial dysfunction and trigger retrograde signaling pathways. We will discuss the requirement of CL for cellular signaling pathways, such as protein kinase C (PKC) signaling. Upon mitochondrial stress, CL is externalized on the outer mitochondrial membrane forming a binding platform for the specific recruitment of signaling molecules. CL microdomains play a role in autophagy, apoptosis and inflammasome signaling.

### CL IS ESSENTIAL FOR MITOCHONDRIAL FUNCTIONS

### CL Species Composition and Biosynthesis

Mitochondrial membranes have a characteristic lipid composition, with a high CL content in the IM, where it contributes up to 20% of total lipids (Gebert et al., 2009; Schlame and Greenberg, 2016; Tatsuta and Langer, 2016). The outer mitochondrial membrane also contains CL, where it constitutes up to 3% of the total lipid content (de Kroon et al., 1997). Peroxisomes are the only other organelles, in which a significant amount of CL (2–4% CL of the total phospholipids) was identified (Wriessnegger et al., 2009). CL is composed of a glycerol head group and two phosphatidylglyceride backbones. In total, four fatty acids chains are bound to CL, which differ in length and saturation. In most mammalian tissues, a multitude of different fatty acids bound to CL form a highly diversified CL pool (Maguire et al., 2017). In some tissues, like the mammalian heart, the CL pool consists of a defined species composition with linoleic acid (18:2) being the predominant form for all four acyl chains (Hoch, 1992). It seems that tissue specific metabolism and energy requirements, determine specific fatty acid composition of CL (Paradies et al., 2014). A significant adaption of the CL pool to external stress signaling has been reported in several studies. In yeast, increased total CL levels were found under temperature stress (Luévano-Martínez et al., 2015). A recent study observed increased total levels of CL and increased linoleic acid (18:2) content in CL, in skeletal muscle in response to overload stimuli (Fajardo et al., 2017).

The IM is the location of the CL biosynthesis and remodeling pathway (**Figure 1A**). Phosphatidic acid (PA) is imported from the ER and transported across the IMS with the help of the protein complex PRELID-TRIAP1 (Connerth et al., 2012; Potting et al., 2013; Tatsuta and Langer, 2016). After activation of PA by the CDP-DAG synthase TAMM41 (Kutik et al., 2008), the phosphatidylglycerol phosphate synthase (PGS1) catalyzes the committed step by converting CDP-DAG to phosphatidylglycerol phosphate (PGP) (Shen and Dowhan, 1998; Zhong and Greenberg, 2003; He and Greenberg, 2004). Phosphatidylglycerol (PG) is formed by the phosphatase PTPMT1 (Protein-tyrosine phosphatase mitochondrial 1) (Xiao et al., 2011; Zhang et al., 2011). A second molecule of CDP-DAG is used by the Cardiolipin Synthase (CLS1) to form premature cardiolipin (pCL) (Chang et al., 1998; Chen et al., 2006; Lu et al., 2006). After its initial synthesis CL is remodeled by the exchange of fatty acid moieties (**Figure 1B**). Several phospholipases (iPLA2, iPLA2γ, iPLA2-VIA) have been suggested to deacytylate pCL to form monolysocardiolipin (MLCL) (Mancuso et al., 2007; Malhotra et al., 2009; Yoda et al., 2010; Hsu et al., 2013). The coenzyme A independent acyltransferase Tafazzin then mediates the reacylation to form mature CL (Bissler et al., 2002; Houtkooper et al., 2009a). The gene encoding for Tafazzin is mutated in Barth Syndrome, and the resulting inactivation of this enzyme causes a block in the generation of mature forms of CL (Lu et al., 2016). Despite the remarkable tissue specific composition of CL, isolated Tafazzin was found to have little or no specificity toward fatty acids (Houtkooper et al., 2009b). The specific remodeling of CL arises from the physical properties of the lipid environment and is ensured by the tissue specific availability of substrate fatty acids and the packing conditions of lipids within the membrane (Schlame et al., 2017). Interestingly, Tafazzin is not the only enzyme, capable of CL remodeling. Under pathological conditions, ROS induced upregulation of Acyl-CoA:lysocardiolipin acyltransferase 1 (ALCAT1) causes an aberrant remodeling of CL. ALCAT1 generates CL species with long and polyunsaturated acyl chains such as docosahexaenoic acid (DHA), which are prone to ROS (reactive oxygen species) induced oxidation (Li et al., 2010). ALCAT1 upregulation is associated with hyperthyroid cardiomyopathy and diabetes. The four fatty acid moieties in close proximity make CL susceptible to peroxidation by ROS. As damaged CL is harmful to mitochondria it has been suggested that oxidized CL is rapidly degraded. The CL phospholipase HSD10 has been identified recently to mediate the degradation of oxidized CL (Boynton and Shimkets, 2015).

### The Role of CL in Mitochondrial Protein Transport

Ninety-nine percent of mitochondrial proteins are encoded in the nucleus and transported into mitochondria after their translation in the cytosol. These proteins are transported by specialized

translocases across mitochondrial membranes (**Figure 2A**). The Translocase of Outer Membrane (TOM) serves as a central entry gate for almost all mitochondrial proteins (Neupert and Herrmann, 2007; Dudek et al., 2013b). Protein import is highly regulated by posttranslational modification. Upon a metabolic switch to glycolysis in yeast, Protein Kinase A phosphorylates the TOM receptor subunit Tom70, exerting an inhibitory effect on protein import (Rao et al., 2011; Schmidt et al., 2011). PKA also reduces TOM assembly, whereas Casein kinase and Cyclin dependent kinase (Cdk1) promotes assembly of the TOM complex (Rao et al., 2012; Harbauer et al., 2014). After their import, β-barrel proteins are integrated into the outer membrane by the Sorting and Assembly Machinery (SAM) (Gebert et al., 2009). Inner membrane proteins are transported by the Translocase of inner membrane TIM23 and the associated Presequence Translocase-associated protein import motor (PAM) drives the import of proteins into the matrix (Dudek et al., 2013b). The TIM22 complex in the IM integrates metabolite carriers into the inner membrane. TOM, SAM, TIM22, TIM23 and the associated motor (PAM) require CL for their structural integrity indicating that CL is essential for mitochondrial protein import (Gallas et al., 2006; Tamura et al., 2006; Kutik et al., 2008). AGK has been recently identified a novel structural constituent of the TIM22 complex (Kang et al., 2017; Vukotic et al., 2017). Interestingly, AGK has a catalytic domain, which catalyzes the phosphorylation of DAG to PA. As PA is a precursor for CL, it has been suggested that AGK contributes to CL biosynthesis (Waggoner et al., 2004; Tatsuta and Langer, 2016). Defective mitochondrial import causes protein accumulation in the cytosol, and triggers the unfolded protein response activated by protein mistargeting (UPRam) in yeast (Wrobel et al., 2015). This signaling pathway reduces cytosolic translation and activates the proteasomal degradation pathway. An increase in proteasomal activity in response to a defect in the intermembrane space was also found in mammals suggesting that a similar pathway might also exist in mammals (Papa and Germain, 2011). After import and maturation, cofactors are integrated into mitochondrial proteins, which then acquire their native conformation. CL also plays a role in cofactor integration, as reduced enzymatic activity of Fe-S containing proteins has been reported in CL-deficient yeast (Patil et al., 2013).

## The Role of CL in Mitochondrial Morphology

With four acyl chains bound to one head group, CL adopts a cone-shaped structure and segregates into regions of locally high membrane curvature or even induces membrane bends (Huang et al., 2006). CL is anticipated to localize in the highly bended regions within the cristae structures and preferentially resides in the monolayer facing the matrix side. In addition, the distribution of CL is determined by the lipid scaffolding proteins prohibitin-1 and prohibitin-2 (Merkwirth et al., 2008). Prohibitins belong to the stomatin/prohibitin/flotillin/HflK/C (SPFH) family, which also includes lipid raft-associated proteins. Prohibitin-1 and prohibitin-2 share more than 50% identical amino acids and form large hetero-oligomeric ring-like structures. The somatin like protein 2 (SLP-2) binds to prohibitins and was found to directly interact with cardiolipin (Da Cruz et al., 2008; Osman et al., 2009). Prohibitin complexes are suggested to segregate CL into specialized membrane domains (Merkwirth et al., 2012). A protein with similarities to the family of Hsp40 co-chaperones, DNAJC19, was identified as a prohibitin interaction partner. As DNAJC19 plays a detrimental role for the CL acetylation pattern, it was suggested that the prohibitin/DNAJC19 complex may segregate a specific membrane domain which facilitates Tafazzin mediated CL remodeling (Richter-Dennerlein et al., 2014).

MitoPLD converts CL into phosphatidic acid (PA), which facilitates MFN-mediated fusion. Fission is mediated by recruitment of DRP1 oligomers to the OM, where it interacts with CL. DRP1 oligomerization is intensively regulated by posttranslational modification. OM, outer membrane; IM, inner membrane; IMS, intermembrane space.

The role of CL in the mitochondrial morphology explains the morphological changes found in cardiolipin-deficient cells. CL deficiency results in an increase in mitochondrial size and absent or disorganized cristae structures (Xu et al., 2006; Acehan et al., 2007, 2009). Mitochondrial morphology is essential for normal mitochondrial function and plays a role in development, aging and apoptosis (Chan, 2006). A large amount of studies implicate changes of mitochondrial morphology in cardiovascular diseases (Brady et al., 2006; Cribbs and Strack, 2007; Williamson et al., 2010).

The complex mitochondrial morphology is determined by several protein structures, in which CL plays a role. The structural organization of the IM and the cristae structures are formed by the mitochondrial contact site and cristae organizing system (MICOS), located at the cristae junctions (Darshi et al., 2011; Gómez and Hagen, 2012). MICOS-deficient mitochondria show a reduction of the number of cristae junctions, resulting in stacks of lamellar cristae. The MICOS complex is formed by nine subunits and interaction partners in the inner membrane, exposing their functional domains into the IMS (Alkhaja et al., 2011; Darshi et al., 2011; Harner et al., 2011; von der Malsburg et al., 2011; Ding et al., 2015; Guarani et al., 2015; Ott et al., 2015; van der Laan et al., 2016). Two proteins MIC26 (APOO) and MIC27 (APOOL) are structurally related to the family of apolipoproteins. MIC27 directly interacts with CL and this interaction was found to be essential for MIC27 assembly into the MICOS complex (Weber et al., 2013; Friedman et al., 2015). The MICOS complex is a central component of a large interaction network which includes several complexes in the IM and OM (Rampelt et al., 2016). MICOS interacts with the protein import and assembly machinery (TOM, SAM in the OM and the IMS resident MIA40), and fusion proteins (OPA1 (see below) and SLC25A46, a mammalian homolog of the yeast fusion adaptor Ugo1) (Abrams et al., 2015; Janer et al., 2016).

Two opposing processes maintain the reticular network of mitochondria: fusion and fission. Fusion (merging of two mitochondria) is mediated by a set of dynamin related GTPases: the mitofusins (MFN1, MFN2) in the OM and OPA1 in the IM (**Figure 2B**) (de Brito and Scorrano, 2008; Westermann, 2010; Shirihai et al., 2015; Schrepfer and Scorrano, 2016). Their activity is counteracted by fission (segregation of two mitochondria), mediated by DRP1, which also belongs to the family of dynamin related GTPases. Fission and fusion maintains the homogeneity of mitochondria within a cell (Sesaki and Jensen, 1999) and has been shown to be functionally dependent on CL. The fusion protein OPA1 is anchored to the IM by an N-terminal transmembrane domain, exposing the GTPase domain into the IMS. Alternative splicing of OPA1 give rise to two long forms L-OPA1, which are processed by proteases to three short forms S-OPA1 (Ishihara et al., 2006; Anand et al., 2014). A balanced formation of both forms is required for the formation of active dimers and maintaining a reticulated morphology of mitochondria. CL was found to be necessary for the dimerization and induction of the GTPase activity (DeVay et al., 2009; Meglei and McQuibban, 2009). A recent study suggests that the IMS domain of OPA1 contains a CL-binding site. It has been proposed, that after fusion of the OM, this binding domain allows OPA1 to interact with CL in the opposing IM. Therefore, CLand OPA1-mediated membrane bridging will induce fusion of the IM (Ban et al., 2017). Mitochondrial fission is mediated by DRP1, which is recruited from the cytosol to oligomerize into helical structures. Activation of its GTPase mediates constriction of the outer membrane and induces mitochondrial fission. DRP1 also shows a strong affinity to CL and recent studies indicate that CL binding enhances oligomerization and GTP hydrolysis (Bustillo-Zabalbeitia et al., 2014; Stepanyants et al., 2015). A balanced equilibrium of fusion and fission is essential for maintaining mitochondrial morphology and necessary for mitochondrial inheritance, as recent data in yeast indicate (Böckler et al., 2017). The correct mitochondrial morphology is also a prerequisite for the many interactions of mitochondria with other organelles. Mitofusins act as tethering molecules, which mediate the contact to the ER (Schrepfer and Scorrano, 2016). In cardiac tissue, mitofusins are important for the SRmitochondrial Ca 2+ transmission and required for regulating mitochondrial metabolism (Chen et al., 2012).

Based on the functional dependence of the fission and fusion machinery on specific lipids, regulated synthesis and degradation of specific lipids may part of a signaling pathway on mitochondrial membranes. MitoPLD is an OM resident phospholipase, converting cardiolipin (CL) to phosphatidic acid (PA). The generation of PA was found to facilitate mitofusindependent fusion of the outer membrane (Choi et al., 2006). PA also recruits the PA phosphatase Lipin 1b, which converts PA to diacylglycerol (DAG). By an unknown mechanism DAG then mediates reduced fusion and increased fission (Huang et al., 2011). Also the phospholipase PA-PLA1 was shown to play a role in hydrolyzing PA and counteracting MitoPLD (Baba et al., 2014). The generation of lipid signals might be important to locally define regions of membrane fusion and fission.

### The Implication of Mitochondrial Morphology for Cellular Signaling

CL plays a fundamental role in forming the shape of mitochondria and mitochondrial morphology is strongly affected in CL-deficient cells. Interestingly, correct mitochondrial morphology is a prerequisite for many cellular signaling pathways, as these pathways are strongly affected in cells with disturbed mitochondrial morphology. Fragmentation of mitochondria in mouse skeletal muscle, induced by the mutation of genes involved in mitochondrial morphology (see above) was reported to cause severe changes insulin signaling, without interference with the contractile functions (Sebastián et al., 2012; Touvier et al., 2015; Wai et al., 2015). Other studies describe skeletal muscle atrophy induced by enforced expression of the fission machinery, which causes AMPK activation and FOXO3 signaling resulting in autophagy activation (Romanello et al., 2010).

Increased fission in apoptotic cells coincides with the release of cytochrome c and the execution of apoptosis. Inactivating DRP1 causes a delay in cytochrome c release, caspase activation and a block in apoptosis (Frank et al., 2001; Breckenridge et al., 2003; Barsoum et al., 2006). Mitochondrial fission is also necessary for mitophagy in many cell types, as mitochondria are too large for autophagic removal (Arnoult et al., 2005; Twig et al., 2008). Dedicated signaling pathways control mitochondrial morphology during mitophagy. Starvation activates PKA and mediates phosphorylation of DRP1. Phospho-DRP1 is retained in the cytosol resulting in mitochondrial elongation which excludes elongated mitochondria from mitophagy (Cribbs and Strack, 2007; Gomes et al., 2011). DRP1 acts as a central hub in the regulation of mitochondrial morphology and is modified by phosphorylation ubiquitination, sumylation, and S-nitrosylation (Karbowski et al., 2007; Zunino et al., 2007; Cereghetti et al., 2008; Han et al., 2008; Cho et al., 2009; Wu et al., 2011). The stressinduced mitochondrial hyperfusion pathway (SIMH) triggers mitochondrial hyperfusion in response to cellular stress. MFN1 and long isoforms of OPA1 have been found to be required for the SIMH response, which establishes a compensating increase in OXPHOS and ATP levels (Tondera et al., 2009). Interestingly, the CL binding protein Stomatin-like protein 2 (SLP-2) was found crucial for regulating OPA1 processing upon SIMH activation (Christie et al., 2011). SLP-2 was also found to be involved in affect supercomplex stabilization and in segregating cardiolipin into lipid domains the inner membrane (Mitsopoulos et al., 2015).

### Role of CL in the Respiratory Chain and Metabolism

The oxidative metabolism in the cell generates NADH and FADH2, which are oxidized by the respiratory chain in the mitochondrial inner membrane. The respiratory chain consists of five complexes (complex I–V) involved in the electron transport to molecular oxygen. The electron transport is coupled with proton export across the IM, generating the membrane potential, which serves as an energy source for the production of ATP by the F1Fo-ATPase. Evidence for CL playing a role in the structural integrity and enzymatic activity exists for all complexes. Structural analysis found specific binding sites for CL in complex I (Fiedorczuk et al., 2016), complex III (Lange et al., 2001; Palsdottir et al., 2003), and complex IV (Itoh et al., 2007). The succinate dehydrogenase (complex II) is strongly reduced in BTHS, indicating a role of CL in the structural integrity of the complex (Dudek et al., 2015). For complex III and complex IV an active role of CL in the proton translocation has been suggested (Eble et al., 1990; Morelli et al., 2013; Duncan et al., 2016). Respiratory chain complexes assemble into large supercomplexes, in which complex I binds a dimer of complex III and several copies of complex IV (**Figure 3**; Letts et al., 2016; Wu et al., 2016). Supercomplex formation increases the efficiency of the electron translocation and minimizes the risk for the generation of ROS (Schagger and Pfeiffer, 2000). Structural analysis found about 200 CL molecules to be associated with the bovine mitochondrial supercomplex (Pfeiffer et al., 2003; Zhang et al., 2005; Mileykovskaya et al., 2012; Bazan et al., 2013). Analysis of CL-deficient mitochondria indicates a remodeling of supercomplexes with a shift from higher- to lower molecular weight complexes. Therefore, it has been concluded, that CL is required for the structural integrity of supercomplexes (Dudek et al., 2013a, 2015; Huang et al., 2015). CL is also essential for the structure of the family of mitochondrial carrier proteins, which includes the phosphate carrier (PiC), the pyruvate carrier, the tricarboxylate carrier and the carnitine/acylcarnitine translocase (Claypool et al., 2006, 2008; Claypool, 2009).

A large number of phosphorylation sites has been reported for respiratory chain subunits indicating that reversible

phosphorylation plays a prominent role in regulation of respiration (Covian and Balaban, 2012). Other covalent modifications of respiratory chain subunits include the modification by lysine acylation and succinylation, which is reversed by the family of mitochondria-localized NAD+ dependent deacetylases SIRT3, SIRT4, and SIRT5.  SIRT5 regulates the respiratory chain by desuccinylation (Zhang et al., 2017). A specific binding of an N-terminal amphipathic helix in SIRT5 to CL was recently found to be essential for its activity (Zhang et al., 2017).

## SIGNALING PATHWAYS AFFECTED BY CL Mitochondria Play a Central Role in ROS Signaling

The structural remodeling of the respiratory chain in BTHS is associated with the aberrant transfer of electrons onto O2, forming superoxide (.O − 2 ) and other forms of ROS (Dudek et al., 2015). When ROS defense mechanisms are overwhelmed, ROS directly reacts with lipids, proteins and DNA (Davies, 1995). ROS also targets proteins of the respiratory chain, affecting its structure and initiates a vicious cycle by further generating more ROS (Nickel et al., 2014, 2015). ROS-induced cellular damage is associated with many forms of cardiac disease in aging (Gómez and Hagen, 2012), ischemia/reperfusion (Ostrander et al., 2001) and heart failure (Saini-Chohan et al., 2009).

A role for ROS as a second messenger in cellular signaling pathways has also been described. ROS activates signaling proteins by changing their redox status. The most susceptible molecular targets of ROS are cysteine and methionine residues. Oxidation of cysteine causes intra- or intermolecular disulfide formation or nitrosylation and glutathiolation. Other posttranslational redox modifications include the hydroxylation of proline and arginine and the nitration of aromatic amino acids. ROS have been integrated as second messengers in many fundamental signaling pathways, modulating the cellular response. Activation of many pathways including the PI3K/Akt pathway and TNFα signaling are associated with an increase in ROS levels (Nogueira et al., 2008). The balance between ROS generating and ROS detoxifying mechanisms are crucial for the activation of downstream pathways. Hypoxia is associated with an increase in ROS levels in many cell types and ROS was identified as an essential contributor to the stabilization of the transcription factor HIF-1α (Diebold and Chandel, 2016). Other transcription factors, which are reported to be activated by ROS include NF-κB, FOXO3A, p53, and PGC1α (Gloire and Piette, 2009; Chae et al., 2013; Marinho et al., 2014). Upstream signaling components are also regulated by ROS, like the Protein kinase C (PKC), which is directly activated by oxidation of cysteine residues (Gopalakrishna et al., 1997). In cardiac hypertrophy AMPK and many MAPKs including p38, JNK, apoptosis-signaling kinase 1 (ASK-1) and ERK1/2 (Giordano, 2005) are activated by ROS. Mitochondria have the ability to significantly amplify a local increase in ROS levels. In a process called ROS-induced ROS release the opening of the mitochondrial permeability transition pore (MPT) and the inner membrane anion channel (IMAC) induces a ROS burst, which not only induces redox-sensitive signaling pathways but also triggers a similar ROS burst in neighboring mitochondria (Zorov et al., 2014).

### Role of CL in MCU and Ca2+ Signaling

Mitochondrial morphology is critical for many signaling pathways. During excitation-contraction coupling, Ca2+ emitted from the sarcoplasmic reticulum is transmitted into the mitochondrion. Ryanodine receptors (RyR) in the sarcoplasmic reticulum are in close proximity to mitochondria and the mitochondrial calcium uniporter (MCU) in the inner membrane. The formation of this functional microdomain has been considered to be essential for the efficient transmission of Ca2+ into mitochondria. Transmission of Ca2+ into mitochondria was absent in fragmented mitochondria (Szabadkai et al., 2004).

Ca2+ plays a prominent role in the regulation of mitochondrial metabolism allowing adaptation to increased energy demands during increased cardiac workload (Dorn and Maack, 2013). Ca2+-mediated activation of the pyruvate dehydrogenase, the isocitrate dehydrogenase and the αketogrutarate dehydrogenase triggers a significant increase in the metabolic flux in the Krebs cycle. At the same time respiration is increased by activation of complex III, IV and V. The Ca2+ conducting pore of the mitochondrial Ca2+ uniporter (MCU) is formed by MCU and EMRE (Baughman et al., 2011; De Stefani et al., 2011; Sancak et al., 2013; Oxenoid et al., 2016). A novel form of regulation has been discovered recently with the finding that oxidative stress causes S-glutathionylation and persistent activity of MCU (Dong et al., 2017). MCUb was described as a dominant negative component of the Ca2+ channel (Raffaello et al., 2013). Regulation of MCU as a calcium induced calcium channel is mediated the Ca2+ sensitive subunits MICU1 and MICU2 (Petrungaro et al., 2015). This regulatory unit is bound to MCU complex via its interaction with EMRE (Tsai et al., 2016). Recently, MICU1 and MICU2 was found to interact with CL, indicating that the binding to CL also contributes to the membrane association (Kamer et al., 2017). Studies in cells, deficient of the regulatory subunits MICU1 and MICU2 revealed a deregulated calcium uptake into mitochondria (Patron et al., 2014). If deregulated Calcium uptake plays a role in BTHS has not been investigated.

### Mitochondrial Protein Kinase C Signaling Depends on CL

Human cells have fifteen isoforms of the Protein kinase C (PKC). Overstimulation of some isoforms are associated with hypertrophy in cardiomyocytes, other forms are associated with a cardioprotective function (Mellor and Parker, 1998). One of the first data of a direct role of CL in PKC signaling was generated in yeast. CL-deficient yeast was found to have a defect in the integrity of the cell wall. Cell wall biogenesis and intracellular turgor pressure are regulated by the PKC-Slt2 and HOG pathway, respectively. CL deficiency caused a defect in the phosphorylation and activation of the MAPK Slt2, the downstream effector of PKC (Zhong et al., 2005, 2007; Zhou et al., 2009). Overexpression of individual genes involved in the PKC-Slt2 pathway rescued the mutant phenotype. As deletion of components of the HOG pathway also rescued the phenotype, it was concluded, that the balanced homeostasis of both pathways is affected in CL-deficient yeast (Patil and Greenberg, 2013). The PKC-Slt2 and the HOG pathway in yeast are also involved in the induction of mitophagy. Recent data indicates that defective activation of both pathways contribute to the described mitophagy defect in CL-deficient yeast (Shen et al., 2017). It is presently unclear, if a direct binding of CL may regulate PKC and Slt2 in yeast or loss of CL has an indirect effect on the components of the Slt2 pathway.

In the heart, several targets for PKC have been identified including the myofilament proteins troponin, cMyBP-C, titin, and the cytoskeletal protein desmin (Hidalgo et al., 2009; Kooij et al., 2010). Although most members of the PKC family are cytosolic proteins, the two diacyglycerol (DAG) sensitive but Ca2+ insensitive PKCε and PKCδ localize to the inner mitochondrial membrane (Budas et al., 2010; Yang et al., 2012). These proteins were found to interact with the cytosolic chaperone HSP90 and the mitochondrial import receptors TOM20 and TOM70. This led to the model of a regulated import into mitochondria upon mitochondrial stress, such as ischemia and reperfusion. Activation of mammalian PKCε was found to be dependent on CL (Konno et al., 1989; Shen et al., 2017). In mitochondria PKC was suggested to regulate mitochondrial proteins, involved in glycolysis, TCA cycle, β-oxidation, and ion transport and it physically interacts with regulatory subunits of the respiratory chain (Ping et al., 2001; Baines et al., 2003). Interestingly, PKCδ was shown to phosphorylate and activate the phospholipid scramblase 3 (PLS3), which is an important mediator of CL externalization (see below), revealing an interesting interdependence of CL and PKC (He et al., 2007). By supporting complex IV activity and preventing pathological ROS release, PKCε was suggested to have a cardioprotective effect (Guo et al., 2007; Yu et al., 2008).

### CL is Precursor for Lipid Mediators Released under Stress Conditions

Lipid mediators such as prostaglandins, thromboxanes, leukotrienes play a role in many pathological processes such as inflammation, fewer, allergy and other immune responses. These signaling molecules are derived from phosphatidylserine (PS) by PLA2-mediated (Ca2+ dependent phospholipases A2) release of polyunsaturated fatty acids, which are subsequently oxidized by cyclooxygenases (COX) and lipoxygenases (LOX). Mitochondrial CL was recently found to be the origin of a new class of mitochondrial mediators. In a very similar process, cytochrome c-mediated oxidation of CL has been proposed to play a role in the production of specific lipid mediators under stress conditions. Oxidized CL was suggested to be a precursor for iPLA2γ-mediated release of oxidized polyunsaturated fatty acids and the accumulation was inhibited by specific inhibitors of iPLA2γ (Tyurina et al., 2014; Liu et al., 2017). Typical mediators, released by this process are 9-Hydroxyoctadecadienoic acid (9-HODE) and 13-HODE. A functional role for lipid mediators in mitochondria is not known, yet. Targets for 9-HODE and 13-HODE have been found predominantly in the cytosol where they were proposed to activate G2A, a G protein-coupled receptor and TRPV1 (Transient receptor potential vanilloid 1), which is expressed in the peripheral and central nervous systems (Patwardhan et al., 2009; Ogawa et al., 2010).

### EXTERNALIZED CL FORMS BINDING SITES FOR SIGNALING MOLECULES

### Mechanisms of CL Translocation into the OM

The specific exposure of phosphatidylserine (PS) on the plasma membrane serves as a well-studied "death signal" in apoptosis. In healthy cells, PS is maintained in an asymmetrical distribution on the cytosolic site of the plasma membrane. The activation of a scramblase upon apoptosis externalizes PS on the outer surface forming a recognition signal for macrophages. The unique biochemical properties of CL have fostered the hypothesis that CL enriched membrane sites might also serve as a binding site for specific protein complexes. Two negative charges allow the specific recruitment of protein interaction partners. CL, which is translocated from the IM and exposed on the OM under stress conditions, will serve as a binding site for signaling molecules and establishes a signaling hub for mitophagy and apoptosis.

In the inner mitochondrial membrane CL shows an asymmetrical distribution with a majority of CL facing the matrix. Phospholipid scramblases (PLS) are a small family of phospholipid translocators, which mediate the translocation of phospholipids between the two monolayers of a lipid bilayer (**Figure 4A**). Mitochondrial phospholipid scramblase-3 (PLS-3) is involved in the redistribution of CL within the membrane, which is considered a necessary step for translocation of CL to the outer membrane. Remarkably, inactivation of PLS-3 causes increased resistance to apoptosis, highlighting the importance of CL translocation for cellular signaling (Liu et al., 2003; Van et al., 2007).

Two IMS proteins, with an established role in energy metabolism, were recently found to have a novel function in the transport of CL into the outer membrane. The Mitochondrial creatine kinase (MtCK) provides the cellular energy buffer phosphocreatine by ATP dependent phosphorylation of creatine. The nucleoside diphosphate kinase (NDPK-D/NM23- H4) converts nucleoside diphosphates to triphosphates by transferring the terminal phosphate group. Both enzymes were found to form large oligomeric complexes in the IMS, exposing specific binding sites for CL (**Figure 4B**). By binding to CL, these proteins were shown to mediate CL transfer between the inner and outer membrane (Lacombe et al., 2009; Schlattner et al., 2013).

### CL Externalization in Mitophagy

Mitophagy is the selective degradation of dysfunctional mitochondria by autophagy. Mitophagy plays an essential role during cardiac development and maintains cardiac function in the adult heart (Billia et al., 2011; Hoshino et al., 2013; Kubli et al., 2013; Gong et al., 2015). During mitophagy, damaged mitochondria are enclosed by a double membrane structure, the phagophore, which forms the autophagosome and fuses with the lysosome to target its content for degradation (Youle and van der Bliek, 2012). Due to their size, mitochondria have to be separated from their reticular network in order to be incorporated into the autophagosomes. Mitochondrial fission has been shown to be a prerequisite for mitophagy in many cell types. Inhibition of mitochondrial fission decreases the level of mitophagy (Thomas and Jacobson, 2012). The two CL dependent proteins DRP1 and OPA1 activity mediate the balanced fusion and fission of mitochondria (see above). Interestingly, another CL binding protein is involved in mitochondrial fission during mitophagy. The mitochondrial human immunity-related GTPase, IRGM, was shown to play an essential role in mitophagy induced fission and translocates from the cytosol to mitochondria, upon mitophagy induction, where it binds to CL (Singh et al., 2010).

The concept of CL externalization forming an essential platform for mitophagy was emerging from the observation that CL translocates to the OM upon CCCP treatment, a well-studied inducer of mitophagy in mammalian cells. Both the content and species composition of CL in the OM were increased upon mitophagy induction (Chu et al., 2013). Externalized CL plays a central role in mitophagy. Phagophore formation is promoted by activation the class III phosphatidylinositol 3-kinase Vps34, which generates phosphatidylinositol-3-phosphate PtdIns(3)P, required for the recruitment of further effectors (Kaur and Debnath, 2015). Vps34 forms a complex with Beclin 1, Vps15, and Ambra1. Beclin 1 is a central regulator of this complex and multiple proteins interact with Beclin 1 to induce or inhibit autophagy (Kroemer et al., 2010). Beclin 1 has been shown to directly interact with CL on the OM, indicating a direct involvement of CL in mitophagy (Huang et al., 2012).

Elongation of the isolation membrane is mediated by two ubiquitin-like protein-conjugation systems. Conjugation of Atg12 to Atg5 allows the formation of the Atg12-Atg5-Atg16 complex, which itself functions as an E3 ligase for LC3. LC3 is the mammalian ortholog of Atg8 and is conjugated to phosphatidylethanolamine (PE) in a reaction involving Atg7 (E1-like) and Atg3 (E2-like) to form membrane-bound LC3- II. Lipidated LC3-II is recruited to both the outer and inner surfaces of the autophagosomal membrane and remains on the autophagosomes until fusion with lysosomes. LC3-II contains basic surface patches which bind to CL (**Figure 5A**; Chu et al., 2013). Consequently, reducing CL levels by knockdown of genes involved in CL biosynthesis causes reduced mitophagy in primary neurons, SH-SY5Y, and HeLa cells (Chu et al., 2013). Recent data suggest that also LC3B, a second member of the Atg8 family, is also specifically recruited to the OM by its interaction with externalized CL (Antón et al., 2016).

## CL Peroxidation Triggers Cytochrome c Release during Apoptosis

Cytochrome c, mediates electron transfer from complex III to complex IV in the respiratory chain. A second function has been discovered during apoptosis, where cytochrome c acts as a catalyst for peroxidation of cardiolipin (CL). Externalization of CL on the IMS side of the IM increases availability of CL for cytochrome c. It has been proposed that conformational changes alter the interaction of CL with cytochrome c causing a change in the coordination of heme (Rajagopal et al., 2012; Vincelli et al., 2013). The resulting increase in peroxidase activity targets the closely bound unsaturated fatty acids in CL and results in a substantial increase in oxidized CL (Abe et al., 2011). Cytochrome c loses its interaction with oxidized CL, allowing cytochrome c release during apoptosis (**Figure 5B**; He, 2010). CL peroxidation precedes the internal/mitochondrial apoptotic pathway (see below) and peroxidized CL was also found to be a major contributor to the opening of the Mitochondrial Permeability Transition Pore (MPTP). The MPTP consists of the OM protein VDAC, the IM protein adenine nucleotide transporter (ANT) and the peripherally associated IMS protein peptidyl-prolyl cis-trans isomerase cyclophilin D. Direct interactions of CL with ANT explain the role of peroxidized CL in triggering the opening of the MPTP (Schlame et al., 2012; Powers et al., 2013).

A major contributor of cytochrome c peroxidase activity has been identified in the protein p66Shc. p66Shc is a splice variant of the growth factor adapter Shc (Francia et al., 2009). Activation of the p66Shc pathway was found in hypergylcemia, which significantly increases the severity of cardiovascular

diseases. Activated PKCβ induces phosphorylation of p66Shc (Pinton et al., 2007). Phosphorylated p66Shc translocates from the cytosol into the mitochondria where it binds to components of the TOM and TIM protein translocases. Proapoptotic stimuli destabilize this complex and induce the release of p66Shc . During apoptosis, p66Shc plays an active role in supporting the peroxidase activity of cytochrome c and the generation of reactive oxygen species. Therefore, p66Shc might have an essential role in the oxidation of CL.

### Role of CL during Execution of Apoptosis

Apoptosis is a controlled process of cell death and maintains the homeostasis of cell populations in tissues and occurs as a defense mechanism in immune reactions or in damaged cells. A role for apoptosis in the heart is described during cardiac development, in ischemia, infarction and in the end stage of heart failure (Bennett, 2002). Apoptosis is executed on two main apoptotic pathways: the extrinsic and the intrinsic pathway. The extrinsic pathway is triggered by the activation of the death receptor such as CD95/FAS/APO-1 in the plasma membrane, followed by the formation of the death-inducing signaling complex (DISC) and the subsequent activation of caspase-8. Caspase-8-mediated processing of Bid to form tBid is necessary to trigger the intrinsic pathway. The intrinsic apoptotic pathway is triggered, when diverse stimuli converge at the mitochondria and induce Bcl-2 family proteins. Bcl-2 proteins are involved in the permeabilization of the outer membrane and the release of proapoptotic factors, including cytochrome c and SMAC/DIABLO. It has been speculated that the externalization of CL into the outer membrane forms a binding platform for the recruitment of multi-protein complexes, which are required for the execution of apoptosis.

Proteins of the Bcl-2 family play a key role in the regulation of apoptosis and can be divided into the pro-apoptotic proteins (Bax and Bak), the anti-apoptotic (Bcl-2, Bcl-XL, Bcl-W, and Mcl1) and the pro-apoptotic BH3-only proteins (Bid, Bim, Bad, Puma and Noxa). The balanced homeostasis of Bcl-2 family proteins is an important regulator of apoptosis. CL has been suggested to serve as a receptor to recruit tBid to the mitochondrial outer membrane (**Figure 5B**). tBid binding to the mitochondria was significantly decreased in mitochondria from cells deficient in CL (Lutter et al., 2000). Upon membrane binding, tBid promotes Bax membrane insertion. The recruitment and oligomerization of Bax and Bak in the outer mitochondrial membrane was also found to be a CL dependent process (Sorice et al., 2004; Lovell et al., 2008; Lucken-Ardjomande et al., 2008). CL-mediated changes in the structure and curvature of the membrane may be a prerequisite for Bax and Bak oligomerization and pore formation.

Execution of apoptosis depends on caspases, a family of cysteine proteases, which are synthesized in the cell as inactive procaspases. In an amplifying proteolytic cascade, active caspases cleave, and activate, other procaspases, resulting in the targeted degradation of key proteins in the cell. Procaspase-8 predominantly localizes in the cytosol. Apoptotic triggers induce the translocation of active caspase-8 to the mitochondria. Mitochondrial recruitment was shown to be dependent on the presence of CL on the outer mitochondrial membrane (**Figure 5B**). Lymphoblastoid cells from BTHS patients have been shown to be resistant to apoptotic triggers (Gonzalvez et al., 2008).

### Role of CL in inflammasome Activation

Myocardial injury is commonly associated with inflammatory signaling. Upregulation of cytokines (TNF), interleukins (IL-1, IL-6), and chemokines (MCP-1) has been observed in heart failure. Inflammatory signaling was shown to increase fibrosis, is involved in myocardial remodeling, myocyte hypertrophy and decreased contractility and has been considered as a driving force for heart failure progression. The inflammasome is a cytosolic protein complex consisting of the receptor NLRP3, the procaspase-1 and ASC, and is essential for caspase-1 activation (Butts et al., 2015). Upon inflammasome activation, caspase-1 is activated and triggers the processing of pro-IL18 and pro-IL-1β into their active forms (Gurung et al., 2015). In cardiac myocytes, IL-1β induces calcium release from the sarcoplasmic reticulum, causing cardiac deficient contractility. The caspase-1-mediated inflammatory form of a programmed cell death (pyroptosis) is responsible for loss of cardiomyocytes and causes heart failure progression. Knockout of NLRP3 in mice results in smaller infarct sizes in an experimental model of acute myocardial infarction (Mezzaroma et al., 2011).

The inflammasome is activated by external molecular signals summarized as PAMPs (pathogen associated molecular patterns, LPS, proteoglycans, double stranded RNA) or endogenous signals, which are called DAMPS (danger associated molecular patterns, ATP, DNA, Chaperones). The finding of mitochondrial derived DAMPs (mito-DAMPs) including ROS, NAD+ and mitochondrial Ca2+ release lead to the hypothesis that mitochondrial dysfunction is a contributor to inflammasome activation (Guo et al., 2015; Chakraborty et al., 2017). Mitochondria play a prominent role for inflammasome activity as they provide CL, which was found to bind to the leucin rich LRR domain in NLRP3. In an in vitro setting, CL was found sufficient for activation of NLRP3. In CLS1 knockdown cells with reduced CL levels, NLRP3 activity was significantly alleviated (Iyer et al., 2013).

## CONCLUSIONS

The mitochondrial phospholipid CL is essential for a large array of functions. Therefore, CL forms multiple interactions with different proteins. As the interaction of CL with proteins serves multiple different functions, a uniform binding site has not been identified. In many of the described binding sites, CL's phosphate groups forms electrostatic interactions with positively charged residues, particularly with Arginine or Lysine (Rytömaa and Kinnunen, 1995). The acyl chains form hydrophobic interactions with Leucin, Isoleucine, and Valine (Kalanxhi and Wallace, 2007). A recent analysis of 62 CL interacting proteins also revealed a role for Glycine in the binding sites, indicating an increased requirement for structural flexibility in the CL binding site (Planas-Iglesias et al., 2015).