Redox Homeostasis and Mitochondrial Dynamics.

Please cite this article in press as: Willems et al., Redox Homeostasis and Mitochondrial Dynamics, Cell Metabolism (2015), http://dx.doi.org/10.1016/ j.cmet.2015.06.006

Cell Metabolism

Perspective Redox Homeostasis and Mitochondrial Dynamics Peter H.G.M. Willems,1 Rodrigue Rossignol,2 Cindy E.J. Dieteren,3 Michael P. Murphy,4 and Werner J.H. Koopman1,* 1Department of Biochemistry, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, 6500HB Nijmegen, The Netherlands 2University of Bordeaux, Maladies Rares: Ge ´ ne´tique et Me´tabolisme (MRGM), 330000 Bordeaux, France 3Department of Cell Biology, Radboud Institute for Molecular Life Sciences, Radboud University Medical Center, 6500HB Nijmegen, The Netherlands 4MRC Mitochondrial Biology Unit, Wellcome Trust/MRC Building, Cambridge CB2 0XY, UK *Correspondence: [email protected] http://dx.doi.org/10.1016/j.cmet.2015.06.006

Within living cells, mitochondria are considered relevant sources of reactive oxygen species (ROS) and are exposed to reactive nitrogen species (RNS). During the last decade, accumulating evidence suggests that mitochondrial (dys)function, ROS/RNS levels, and aberrations in mitochondrial morphology are interconnected, albeit in a cell- and context-dependent manner. Here it is hypothesized that ROS and RNS are involved in the short-term regulation of mitochondrial morphology and function via non-transcriptional pathways. We review the evidence for such a mechanism and propose that it allows homeostatic control of mitochondrial function and morphology by redox signaling. Introduction Mitochondria are motile organelles that are present in virtually every mammalian cell and display a continuous cycle of fission and fusion (Westermann, 2010; Chan, 2012; Birsa et al., 2013). These mitochondrial dynamics are mediated by an extensive protein machinery and, in combination with chemiosmosisrelated mitochondrial swelling/shrinking and removal of damaged mitochondria by mitophagy, determine net mitochondrial morphology (Westermann, 2010; Twig and Shirihai, 2011; Picard et al., 2013; Archer, 2013). At the structural level, mitochondria consist of an outer membrane (MOM) and a highly folded inner membrane (MIM), which protrudes into the mitochondrial matrix compartment (cristae). The MIM and MOM are separated by the mitochondrial intermembrane space (IMS) and are partially connected via contact sites, involved in cristae organization (Mannella et al., 2013; Pfanner et al., 2014). Mitochondrial fission (Figure 1A) is guided by contact sites between mitochondria and endoplasmic reticulum (ER) tubules (Friedman et al., 2011) and executed by Dynamin-related protein 1 (Drp1), a cytosolic guanosine triphosphatase that is recruited from the cytosol to the MOM (Lackner and Nunnari, 2010; Chan, 2012). Currently, four adaptor proteins have been identified that mediate Drp1 recruitment (Otera et al., 2010; Chan, 2012; Losoń et al., 2013): human fission protein 1 (hFis1), mitochondrial fission factor (Mff), and two mitochondrial elongation factors (MIEF1/MiD51 and MIEF2/MiD49). Mitochondrial fusion (Figure 1A) occurs by sequential merging of the MOM and MIM (Mishra et al., 2014). This process is ATP-dependent, requires a sufficiently negative trans-MIM electrical potential (Dc), and is mediated by two mitofusins (Mfn1/Mfn2; MOM-fusion) and the optic atrophy 1 protein OPA1 (MIM-fusion; Meeusen et al., 2004; Frezza et al., 2006; Chan, 2012; Belenguer and Pellegrini, 2013; Boissan et al., 2014). Mitochondria can display a large variety of shapes (Figure 1B), ranging from ‘‘giant’’ spherical morphologies (Yoon et al., 2006) to ‘‘hyperfused’’ reticular networks (Mitra et al., 2009). Evidence was provided that changes in mitochondrial volume and/or (ultra)

structure affect the kinetics of biochemical reactions and assembly of protein supercomplexes in the mitochondrial matrix (Lizana et al., 2008; Dieteren et al., 2011; Cogliati et al., 2013; Mannella et al., 2013). Although much progress was made during the last decade (e.g., Benard et al., 2007, 2013; Twig and Shirihai, 2011; Chan, 2012; Picard et al., 2013; Archer, 2013; Jose et al., 2013; Varikmaa et al., 2013; Rambold et al., 2015), it is still unclear how the various mitochondrial morphologies are mechanistically linked to mitochondrial functions. This might relate to the fact that various fission and fusion proteins also have roles outside of mitochondrial dynamics (e.g., Zorzano et al., 2010; Schrader et al., 2012). During the last decade, experimental strategies combining live-cell microscopy and fluorescent reporter molecules were developed to allow unbiased 2D and 3D quantification of mitochondrial morphology and statistical analysis (Figures 1C–1E; reviewed in Tronstad et al., 2014). Quantitative analysis is crucial to discriminate between mitochondrial phenotypes that ‘‘look’’ similar. For instance, visual inspection of mitochondrial microscopy images might not always be suited to distinguish between ‘‘true’’ mitochondrial fragmentation (i.e., cleavage of mitochondrial tubules resulting in smaller mitochondria that are more abundant) and ‘‘apparent’’ fragmentation (i.e., mitochondrial tubules become smaller but mitochondrial number does not increase). Assessing the continuity of the mitochondrial matrix compartment is crucial to functionally understand the sharing of mitochondrial content such as mtDNA, tRNA, mRNA, proteins, and small molecules such as ATP, ADP, and antioxidants. Studies using Dc-sensitive reporter molecules revealed the existence of equipotential mitochondrial ‘‘sub-networks,’’ suggesting that matrix connectivity is not complete throughout mitochondrial filaments that appear continuous (Twig et al., 2008). Matrix connectivity can be experimentally demonstrated in living cells using photo-convertible proteins (Figure 1F; Gurskaya et al., 2006), photobleaching strategies like FRAP (fluorescence recovery after photobleaching) and FLIP (fluorescence loss in photobleaching; e.g., Dieteren et al., 2011; Distelmaier et al., Cell Metabolism 22, August 4, 2015 ª2015 Elsevier Inc. 1


Cell Metabolism

Perspective A

Fusion

B

Physiology

Stress, apoptosis?

Stress, starvation?

Mfn1/2 OPA1

Drp1 Fis1 MIEF1/2 Mff

Irreversible Excessive ROS stress?

Fission C

"Giant"

RAW

COR

LCS

THF

Exogenous H2O2 BSO, FCCP Damaged? Apoptosis

Cell division Resting state? Mitophagy permissive

Spherical

Fragmented

MED

BIN

MSK

Antioxidant sharing Metabolically active?

Mitophagy preventing Antioxidant sharing Stress adapted

Filamentous

Tubular

D

“Hyperfused”

E 200

Intensity (grayvalue)

100

6 4

t=2.095 s

t=2.541 s

t=15.084 s

160 140 120 100 80 60 40

0

20 1

10

100 1000 10000

Area (pixels)

In

Ar

G

F

180

2

ea AR

15 µm

8

te F ns ity

Descriptor value

200

t= 94 s

t=118 s

15 µm

15 µm

t=122 s

Low

High

Figure 1. Mitochondrial Dynamics and Its Analysis in Single Living Mammalian Cells (A) Net mitochondrial morphology primarily results from the balance between fusion and fission. These processes are carried out by a set of eight core proteins involved in fusion (Mfn1, Mfn2, OPA1) and fission (Drp1, Fis1, MIEF1/MiD51, MIEF2/MiD49, Mff). (B) Mitochondria display various net morphologies depending on the cell type and/or metabolic state. In physiological (healthy) conditions (green), mitochondria are motile and continuously fuse and divide. This state is associated with morphologies ranging between fragmented and filamentous. Stress conditions are associated with giant/spherical (red) or hyperfused morphologies (blue) that appear to be linked to apoptosis induction and stress adaptation, respectively. (C) Image processing scheme for quantification of mitochondrial morphology and function in healthy primary human skin fibroblasts (CT5120; Koopman et al., 2008). Mitochondria are stained with TMRM (tetramethylrhodamine methyl ester) and imaged using epifluorescence microscopy. The resulting image (RAW) is background corrected (COR), contrast optimized (LCS), spatially filtered using a top-hat filter (THF), median filtered (MED), and thresholded to yield a binary image (BIN). Combining the BIN and COR image yields a masked image (MSK) suited for image quantification. The color coding reflects TMRM fluorescence intensity ranging from low (blue) to high (red). (D) Quantification of average mitochondrial morphology parameters for the 50 mitochondrial objects in (C) (MSK image). Depicted are mitochondrial area (in pixels), aspect ratio (AR, a measure of mitochondrial length), formfactor (F, a combined measure of mitochondrial length and degree of branching), and intensity (a measure of Dc). The error bars reflect the standard error of the mean (SEM). (E) Relationship between mitochondrial area and TMRM fluorescence intensity. Each point represents an individual mitochondrial object. (F) CT5120 cell expressing a mitochondria-targeted variant of the photo-convertible protein Dendra2. Upon pulse illumination with 405 nm light, the greenemitting form of Dendra2 is converted into red-emitting Dendra2 (white box). The red-emitting Dendra2 will time-dependently redistribute throughout the mitochondrial filaments if their matrix is continuous. Cells were transfected using a baculoviral expression system and imaged by confocal microscopy. (G) CT5120 cell stained with TMRM. Extended illumination of the TMRM induced a synchronous Dc depolarization in a part of the mitochondrial network (subnetwork), as reflected by the reversible loss of mitochondrial TMRM fluorescence. The latter is caused by photo-induced opening of the mitochondrial permeability transition pore (mPTP) and suggests that the subnetwork is electrically coupled (i.e., possesses a continuous mitochondrial inner membrane). Cells were imaged using epifluorescence microscopy.

2015), or analysis of photo-induced reversible mitochondrial permeability transition pore opening (Figure 1G; Ricchelli et al., 2011; Blanchet et al., 2014). Mitochondria generate ATP by oxidative phosphorylation (OXPHOS; Mitchell, 1961) and participate in numerous other metabolic pathways as well as energy sensing, making these organelles hubs of metabolic signal integration (Frohman, 2010; Jose et al., 2013; Ho¨nscher et al., 2014). The OXPHOS 2 Cell Metabolism 22, August 4, 2015 ª2015 Elsevier Inc.

system is MIM embedded and abstracts electrons from NADH (at complex I [CI]) and FADH2 (at complex II [CII]), which are transported to complex III (CIII; by coenzyme Q10) and complex IV (CIV; by cytochrome c). At the latter complex, electrons react with molecular oxygen (O2), yielding H2O. Electron transport is linked to proton (H+) efflux from the mitochondrial matrix (at CI, CIII, and CIV), and the resulting proton-motive force (PMF) is used to drive ATP generation by complex V (CV). The PMF


Cell Metabolism

Perspective consists of an electrical (Dc) and chemical (DpH) component, which are not only essential for ATP production but also sustain many other mitochondrial functions including metabolite/ion exchange and import of mitochondrial precursor proteins from the cytosol (Szabo and Zoratti, 2014). As a consequence of normal OXPHOS, mitochondria generate reactive oxygen species (ROS) in a physiological range (Murphy, 2009). In human disease models, increased mitochondrial ROS production has been linked to pathology induction (Yu et al., 2006; Blanchet et al., 2011; Nakamura and Lipton, 2011; Nakamura et al., 2013; Qi et al., 2011; Halliwell and Gutteridge, 2012; Jose et al., 2013; Koopman et al., 2012, 2013; Niemann et al., 2014). CI and CIII are most likely the major sources of ROS in intact cells (Murphy, 2009). Under certain conditions CII and other mitochondrial and cellular ROS sources can also be relevant (Koopman et al., 2010; Brown and Borutaite, 2012; Mailloux et al., 2013; Quinlan et al., 2013; Woolley et al., 2013). The superoxide anion (O2d ) is the primary ROS generated and is converted into hydrogen peroxide (H2O2) by superoxide dismutases (SODs; Auche`re and Rusnak, 2002). The highly reactive hydroxyl radical (dOH), generated via the Fenton reaction from O2d and H2O2, can initiate formation of lipid (Ld) and lipid peroxyl (LOOd) radicals. H2O2 is converted into H2O by enzymes including peroxiredoxins, catalase (CAT), and glutathione peroxidases (GPX). Proper functioning of these systems further requires the action of various other enzymatic and non-enzymatic systems (Koopman et al., 2010; Brown and Borutaite, 2012; Mailloux et al., 2013; Quinlan et al., 2013; Woolley et al., 2013). In the context of this review it is important to mention that mitochondria are exposed to the reactive nitrogen species (RNS) nitric oxide (NOd) and that NOd can react with O2d to form peroxynitrite (ONOO ). This molecule can be converted into various other reactive species including dOH (Valerio and Nisoli, 2015). Coincident alterations in ROS/RNS levels and mitochondrial morphofunction have been described in various experimental studies (see next section), suggesting that there is a mechanistic link between redox homeostasis and regulation of mitochondrial morphology. Below, we first review the evidence for such a connection and then focus on redox-induced posttranslational modifications of mitochondrial fission/fusion proteins and their binding partners. We conclude by proposing a working model integrating mitochondrial redox homeostasis with mitochondrial morphofunction. Links between Redox Homeostasis and Mitochondrial Morphology Parallel changes in ROS/RNS levels and mitochondrial morphology have been reported in many experimental studies. For example, patient primary fibroblasts with a greatly reduced CI activity display a fragmented mitochondrial phenotype and highly increased ROS levels, whereas cells with a moderately reduced CI activity display normal mitochondrial morphology and moderately increased ROS levels (Koopman et al., 2007; Blanchet et al., 2011). Also in fibroblasts, exogenous H2O2 addition stimulated ubiquitination of Mfn1/Mfn2 (but not of OPA1 or hFis1) paralleled by mitochondrial fragmentation (Rakovic et al., 2011). Conversely, lowering fibroblast ROS levels by the antioxidant Trolox stimulated Mfn2-dependent mitochondrial filamentation and increased OXPHOS protein expression

and enzymatic activity (Distelmaier et al., 2012; Blanchet et al., 2015). Exposing skeletal muscle myoblasts to exogenous H2O2 triggered Dc depolarization and stimulated mitochondrial fragmentation, suggesting that ROS-induced mitochondrial depolarization might be responsible for mitochondrial fragmentation (Fan et al., 2010). In a Drosophila wound-healing model, high ROS levels induced Drp1-mediated mitochondrial fragmentation involving the Rho effector ROCK, whereas low ROS levels promoted mitochondrial filamentation (Muliyil and Narasimha, 2014). Cells devoid of Mfn1 or Mfn2 displayed a fragmented mitochondrial morphology and increased ROS levels (Munõz et al., 2013). Conversely, genetic pro-fusion interventions (inducing mitochondrial elongation) were associated with reduced mitochondrial ROS production, whereas pro-fission interventions (leading to mitochondrial fragmentation) stimulated mitochondrial ROS production (Picard et al., 2013). During hyperglycemic conditions, mitochondrial fragmentation was required to allow increased ROS production (Yu et al., 2006). However, although Drp1 overexpression induced mitochondrial fragmentation in HeLa cells, it was not associated with increased ROS levels (Distelmaier et al., 2012). OPA1 mutations in Drosophila induced aberrations in mitochondrial morphology and increased ROS production and susceptibility to oxidative stressors (Tang et al., 2009), suggesting that mitochondrial filamentation protects against oxidative stress. Glutathione (GSH) plays a central role in ROS-induced redox signaling and stress induction (Mailloux et al., 2013). In the central nervous system (CNS), loss of ganglioside-induced differentiation-associated protein 1 (GDAP1) is compensated by the mitochondrial translocation of GDAP1L1 (a GDAP1 paralog). This recruitment is stimulated by elevated levels of oxidized GSH (GSSG), suggesting that GDAP1-family members can act as redox sensors during cell protection (Niemann et al., 2014). In primary human fibroblasts, inhibition of GSH synthesis by l-buthionine-(S,R)-sulphoximine (BSO, 12.5 mM, 72 hr) shifted the cytosolic and mitochondrial thiol redox environment toward a fully oxidized state (Verkaart et al., 2007). The latter was associated with a reduction in mitochondrial size (‘‘shrinkage’’; Distelmaier et al., 2012), compatible with the proposed link in vivo between oxidation of the mitochondrial matrix environment and mitochondrial tubule shortening (Breckwoldt et al., 2014). Overexpression of BOLA1, a mitochondrial protein that interacts with glutaredoxin 5 (GLRX5), attenuated the BSO-induced effects on thiol redox state and prevented mitochondrial shrinkage (Willems et al., 2013). In contrast, BSO treatment of HeLa cells (100 mM, 24 hr) induced mitochondrial hyperfusion, which might act as a transient protection against apoptosis and mitophagy (Shutt et al., 2012). Mechanistically, the latter study provided evidence that GSSG stimulates generation of disulfide-mediated (Cys684) Mfn oligomers. The different effects of BSO in fibroblasts and HeLa cells suggest that the effects of GSH depletion on mitochondrial morphology depend on BSO concentration, incubation time, and/or cell type (Koopman et al., 2013). Increasing NOd concentrations triggered a dose- and Drp1dependent increase in mitochondrial fragmentation (Cho et al., 2009). Conversely, NOd scavenging stimulated mitochondrial filamentation in human pulmonary arterial endothelial cells (Lee et al., 2013). The above results suggest that increased ROS/RNS levels are linked to a fragmented and (more) Cell Metabolism 22, August 4, 2015 ª2015 Elsevier Inc. 3


Cell Metabolism

Perspective dysfunctional mitochondrial phenotype, albeit in a cell- and context-dependent manner. Most of the experimental evidence points to a mechanism in which low ROS/RNS levels do not affect mitochondrial morphology and/or induce filamentation, whereas too high ROS/RNS levels that cannot be appropriately counterbalanced by the cell’s antioxidant defense system induce mitochondrial fragmentation, swelling, and/or shortening. In this sense, evidence was provided that mild oxidative stress triggers Drp1-dependent mitophagy (Frank et al., 2012). It was proposed that the various mitochondrial structural phenotypes reflect different metabolic and/or (patho)physiological states (Westermann, 2010; Picard et al., 2013). However, it is still not firmly established whether elongated or fused mitochondria perform bioenergetically ‘‘better’’ (or worse) than fragmented ones. In the next section, we discuss the theoretical and empirical evidence linking redox homeostasis to mitochondrial dynamics. Coupling Mechanisms between Redox Homeostasis and Mitochondrial Morphology In principle, mitochondrial dynamics can be controlled by the amount, activity, and localization of the fission/fusion proteins, but also by the lipid composition of mitochondrial membranes (see below). The role of mitochondrial motility and control exerted at the level of protein processing/degradation, mRNA, and transcription/translation are discussed elsewhere (e.g., Benard et al., 2007, 2013; Birsa et al., 2013; Jose et al., 2013; Mishra and Chan, 2014). Here we primarily focus on nontranscriptional regulation by redox-sensitive posttranslational modifications (PTMs), which can occur at a relatively short timescale. To this end, we first generated an inventory of (predicted) PTMs for Drp1, Mfn1, Mfn2, OPA1, hFis1, Mff, MIEF1/MiD51, and MIEF2/MiD49 by integrating information from various online databases and empirical studies (Figure 2). This highlighted various PTMs (i.e., phosphorylation, S-nitrosylation, sumoylation, ubiquitination, tyrosine sulfation, and acetylation). Next, we extended this inventory by also including known and predicted proteinaceous binding partners for Drp1, hFis1, Mfn1/2, and OPA1 (Supplemental Information). Of the core fission/fusion proteins only Drp1 (Cho et al., 2009) and possibly OPA1 (Bossy et al., 2010) are posttranslationally modified in a redox-sensitive manner by S-nitrosylation. In addition, several proteins that bind to the core fission/fusion proteins contain redox-sensitive motifs (Table 1): amyloid-b (S-nitrosylation), CDK5 (S-nitrosylation), Parkin (S-nitrosylation), PKA/ AKAP1 (disulfide bond, nitration), Timm8a (disulfide bond), caspase-3 (S-nitrosylation), paraplegin (nitration), ROMO1 (disulfide bond), and JNK (S-nitrosylation). All of these modifications can be induced by H2O2, NOd, or ONOO , thereby functionally affecting Drp1, OPA1, Mfn2, and mitochondrial structure (Figure 3). The chemistry and biology regarding protein S-nitrosylation (SNO), disulfide bond formation, and nitration is discussed in detail elsewhere (Sacksteder et al., 2006; Nakamura and Lipton, 2011; Halliwell and Gutteridge, 2012; Nakamura et al., 2013; Bak and Weerapana, 2015; Valerio and Nisoli, 2015). Redox Regulation of Drp1 NOd and amyloid-b peptide were demonstrated to induce mitochondrial fragmentation in neurons (Barsoum et al., 2006; Nakamura and Lipton, 2011). Evidence was provided that 4 Cell Metabolism 22, August 4, 2015 ª2015 Elsevier Inc.

NOd elevation increases the GTPase activity of Drp1 by S-nitrosylation of Cys644 in response to amyloid-b peptide (Cho et al., 2009). It was proposed that formation of S-nitrosylated Drp1 (Drp1-SNO) induces Drp1 dimerization and increases GTPase activity (Cho et al., 2009). S-nitrosylation-induced activation of dynamins, including Drp1, was also observed in other studies (see Nakamura and Lipton, 2011 and the references therein). In contrast, another study suggested that Drp1-SNO does not dimerize and displays no increased GTPase activity (Bossy et al., 2010). Although these authors detected OPA1-SNO, they did not observe significant differences in Drp1-SNO levels between brains of age-matched normal and patients with Alzheimer’s (AD) or Parkinson’s disease (PD). Interestingly, experimental evidence suggests that NOd production stimulates the phosphorylation of Ser616, thereby increasing Drp1 activity and MOM recruitment (Bossy et al., 2010). It was proposed that NOd-induced Ser616 phosphorylation is mediated by NOd-stimulated kinases. However, under physiological conditions, Drp1 needs to exist in a non-oxidized form for NOdinduced S-nitrosylation to occur. In this sense, Nakamura and Lipton argued that artifactual Drp1 oxidation might have occurred in the study of Bossy and coworkers (Nakamura and Lipton, 2011). Amyloid-b production was triggered by mitochondria-derived ROS using in vitro and in vivo models (Leuner et al., 2012), leading to activation of Drp1 via stimulating S-nitrosylation (Cho et al., 2009). It was proposed that the ensuing mitochondrial fragmentation (co)contributes to mitochondrial dysfunction induced by amyloid-b (Wang et al., 2008). Ser616 phosphorylation of Drp1 generally promotes mitochondrial fission (e.g., Taguchi et al., 2007; Qi et al., 2011; Gan et al., 2014). Remarkably, CDK5-induced phosphorylation of this serine residue inhibits Drp1 and induces mitochondrial elongation during neuronal maturation under physiological conditions (Cho et al., 2014). It was proposed that this effect was mediated by CDK5-induced reduction of Drp1 oligomerization. Parkin is a cytosolic E3 ubiquitin ligase involved in Drp1 ubiquitination and its subsequent proteasome-dependent degradation (Wang et al., 2011). S-nitrosylation inactivates Parkin, leading to impaired Drp1 degradation and stimulation of mitochondrial fragmentation (Chung et al., 2004; Lutz et al., 2009). PKA-mediated Drp1 phosphorylation inhibits its fission promoting activity, stimulating mitochondrial filamentation (Kim et al., 2011; Merrill et al., 2011). The effect of PKA nitration is currently unknown. Formation of a disulfide bond between the two regulatory subunits of PKA induces activation (Brennan et al., 2006). This suggests that ROS-induced PKA-activation stimulates mitochondrial filamentation by inhibiting Drp1. Timm8a is released from the IMS during apoptotic conditions and binds to cytosolic Drp1 to promote its translocation to the MOM and thereby mitochondrial fragmentation (Arnoult et al., 2005). Timm8a is a protein involved in mitochondrial protein import and contains a disulfide bond that, in yeast, appears to be involved in forming stabilizing intramolecular disulfide bonds important in the formation of the Tim8p-Tim13p complex (Curran et al., 2002). How this is linked to Drp1 translocation is unclear. Although not predicted to contain a redox-sensitive motif, PKCd stimulates Ser616 phosphorylation of Drp1 and stimulates mitochondrial fragmentation under conditions of oxidative stress, providing a mechanism for ROS-induced mitochondrial fragmentation


Cell Metabolism

Perspective A

E

F

B G

C H

D

Figure 2. Domain Structure and Posttranslational Modifications of Core Mitochondrial Fission and Fusion Proteins in Humans (A–H) Domain structure and observed/predicted posttranslational modifications (PTMs) of: (A) Drp1, (B) Mfn1, (C) Mfn2, (D) OPA1, (E) hFis1, (F) Mff, (G) MIEF1/ MiD51, and (H) MIEF2/MiD49. (+), phosphorylation; ( ), dephosphorylation; ADP, ADP binding site; CaN, calcineurin; CC, coiled-coil; DIM, dimerization domain; Drp1, Drp1 interaction; GED, GTPase effector domain; GSK3b, glycogen synthase kinase 3 b; HR, heptad-repeat; LZ, leucine zipper; MTS, mitochondrial targeting sequence; NAL, N6-acetyllysine; NAM, N-acetylmethionine; NO, nitric oxide; NT, nucleotidyltransferase domain; OL, O-linked glycosylation (GlcNac); Olig., oligomerization domain; STyr, sulfotyrosine; T, transmembrane helix; TPR, tetratricopeptide repeat. Data were compiled from UnitprotKB (Homo sapiens; http://www.uniprot.org) and dbPTM (http://dbptm.mbc.nctu.edu.tw): Drp1 (736 aa; O00429), Fis1 (152 aa; Q9Y3D6), Mff (342 aa; Q9GZY8), MIEF1/MiD51 (463 aa; Q9NQG6), MIEF2/MiD49 (454 aa; Q96C03), Mfn1 (741 aa; Q8IWA4), Mfn2 (757 aa; O95140), OPA1 (960 aa; O60313). Abbreviations and literature references are provided in Supplemental Information.

(Qi et al., 2011). Similarly, degradation of the Drp1 Ser637 phosphatase PGAM5 is mediated by Kelch-like ECH-associated protein 1 (KEAP1), which is an important redox stress-sensing protein (Kanamaru et al., 2012). This suggests that the KEAP1PGAM5 complex regulates mitochondrial fission in a redoxdependent manner. Redox Regulation of OPA1 Proteolytic processing of OPA1 is an important means to control its function (e.g., Song et al., 2007; Merkwirth et al., 2008; Ehses et al., 2009; Chan, 2012; Quiro´s et al., 2012; Anand et al., 2014; Mishra et al., 2014). Evidence was provided that OPA1 can be S-nitrosylated (Bossy et al., 2010), although the functional effect of this modification is currently unknown. N-terminal cleavage of OPA1 by caspase-3 induces mitochondrial fragmentation (Loucks et al., 2009). The activity of caspase-3 is inhibited

by S-nitrosylation and increased by denitrosylation during apoptosis activation. This might allow OPA1-mediated regulation of mitochondrial fragmentation by RNS species. Paraplegin is a mitochondrial metalloprotease of the AAA family involved in OPA1 processing (Ehses et al., 2009). Nitration of paraplegin potentially affects its catalytic rate, protein-protein interactions, and phosphotyrosine signaling pathways (Sacksteder et al., 2006; Ehses et al., 2009). This might provide a link between OPA1 processing/function and redox state. The ROMO1 (reactive oxygen species modulator 1) protein was recently discovered as a regulator of cellular ROS generation and redox-dependent modulator of mitochondrial fusion and cristae morphology (Norton et al., 2014). This protein possesses two conserved cysteines (Cys15 and Cys79) that form a redoxsensing module. In its monomeric state, ROMO1 is required to Cell Metabolism 22, August 4, 2015 ª2015 Elsevier Inc. 5


Cell Metabolism

Perspective Table 1. Mechanistic Non-transcriptional Links between Redox Homeostasis and Mitochondrial Dynamics Proteins Protein

Interactor

Function

Redox Modification

Reference

Drp1 and Interactors Drp1

n.a.

Core MOM fission protein.

S-nitrosylation, which stimulates mitochondrial fission. Possibly by enhancing Drp1 MOM recruitment, increasing Drp1 GTPase activity, stimulating Drp1 oligomer formation, or increasing Drp1 phosphorylation at Ser616.

Cho et al., 2009; Bossy et al., 2010

Amyloid-b

Drp1

S-Nitrosylates Drp1 at Cys644 (activating) and possibly promotes Drp1 phosphorylation at Ser616 (activating).

Disulfide bond. Extracellular binding and reduction of copper (Cu) results in a corresponding oxidation at Cys144 and at Cys158 and the formation of a disulfide bond. In vitro, the APP-Cu complex in the presence of hydrogen peroxide results in an increased production of amyloid-b-containing peptides.

Cho et al., 2009

CDK5

Drp1

Phosphorylates Drp1 at Ser616 and inhibits its activity, probably by reducing Drp1 oligomerization.

S-nitrosylation activates CDK5.

Qu et al., 2011; Cho et al., 2014

Parkin

Drp1

E3-ubiquitin ligase for Drp1.

S-nitrosylation inhibits the ubiquitin E3 ligase activity of Parkin. Loss of Parkin impairs Drp1 ubiquitinylation and increases Drp1-dependent mitochondrial fragmentation.

Chung et al., 2004; Lutz et al., 2009

PKA/AKAP1

Drp1

Mediates cAMP-dependent phosphorylation of Drp1 (inactivation) at Ser637 (Ser656 in rat).

Disulfide bond (regulatory a and b subunit) and nitration (regulatory b subunit). Formation of a disulfide bond between the two regulatory subunits of PKA induces activation.

Brennan et al., 2006; Kim et al., 2011; Merrill et al., 2011

PKCd

Drp1

PKCd binds to Drp1 and phosphorylates it (activation) at Ser579 (Ser616 in human).

No redox motif. Drp1 phosphorylation is associated with its translocation to MOM under oxidative stress conditions.

Qi et al., 2011

Timm8a

Drp1

Binds to cytosolic Drp1 when released from the IMS during apoptosis and promotes Drp1 transition to the MOM and mitochondrial fragmentation.

Disulfide bond. Effect unknown.

Arnoult et al., 2005

OPA1 and Interactors OPA1

n.a.

Core MIM fusion protein.

S-nitrosylation (possibly), the effect of which is unknown.

Bossy et al., 2010

Caspase-3

OPA1

N-terminal cleavage of OPA1 by caspase-3 leads to mitochondrial fragmentation.

S-nitrosylation. Caspase-3 is S-nitrosylated on its catalytic site Cys in unstimulated human cell lines and denitrosylated upon activation of the Fas apoptotic pathway, associated with an increase in intracellular caspase activity.

Loucks et al., 2009

Paraplegin

OPA1

Involved in OPA1 processing. Interacts with AFG3L2; the interaction is required for the efficient assembly of mitochondrial CI.

Nitration. Effect unknown. Potentially affecting catalytic rate, protein-protein interactions and phosphotyrosine signaling pathways.

Ehses et al., 2009; Sacksteder et al., 2006

ROMO1

OPA1

Monomeric ROMO1 is required to maintain mitochondrial fusion.

Disulfide bond (intermolecular). ROMO1 is inactivated by GSH-dependent disulfide bond formation in oxidizing environments leading to OPA1 cleavage and cristae shape alteration.

Norton et al., 2014

Phosphorylates Mfn2 at Ser27, leading to recruitment of ubiquitin ligase (E3) Huwe1/Mule/ARF-BP1/ HectH9/E3Histone/Lasu1 to Mfn2 and ubiquitin-mediated proteasomal Mfn2 degradation.

S-nitrosylation (JNK1, JNK3). S-nitrosylation of JNK1 and JNK3 inhibits their activity.

Park et al., 2000; Pei et al., 2008; Leboucher et al., 2012

Mitofusin Interactors JNK

Mfn2

(Continued on next page)

6 Cell Metabolism 22, August 4, 2015 ª2015 Elsevier Inc.


Cell Metabolism

Perspective Table 1.

Continued

Protein

Interactor

Function

Redox Modification

Reference

Parkin

Mfns

E3 ubiquitin ligase for Mfns. PINK1 phosphorylates Mfn2 and promotes its Parkin-mediated ubiquitination and breakdown.

S-nitrosylation inhibits Parkin’s ubiquitin E3 ligase activity. This could inhibit Parkin-mediated Mfn breakdown and stimulate mitochondrial filamentation.

Chung et al., 2004; Glauser et al., 2011; Chen and Dorn, 2013

PKA

Mfn2

Mfn2 is an important suppressor of vascular smooth muscle cell proliferation. PKA phosphorylation at Ser442 suppresses growth independent of Mfn2 morphology effects.

Disulfide bond (regulatory a and b subunit) and nitration (regulatory b subunit). Formation of a disulfide bond between the two regulatory subunits of PKA induces activation.

Brennan et al., 2006; Zhou et al., 2010

Databases used: HGNC (HUGO Gene Nomenclature Committee) database of human gene names (http://www.genenames.org), HPRD (Human Protein Reference Database, http://www.hprd.org), SNObase (S-Nitrosylation database, http://www.nitrosation.org), UnitprotKB (only reviewed protein entries were included, http://www.uniprot.org), STRING (Search Tool for the Retrieval of Interacting Genes/Proteins, http://string-db.org), MINT (the Molecular INTeraction database, http://mint.bio.uniroma2.it/mint/Welcome.do). For abbreviations, see Supplemental Information.

maintain mitochondrial fusion and normal cristae morphology. ROMO1 can be inactivated in an oxidizing environment by formation of an intermolecular disulfide bond causing oligomerization. This inactivation is GSH dependent and paralleled by reduced mitochondrial fusion, OPA1 cleavage, impairment of OPA1 oligomerization, and cristae shape alterations. It was proposed that ROMO1 might mediate the coupling between mitochondrial ROS levels, OPA1 oligomerization, and cristae junction integrity (Norton et al., 2014). Redox Regulation of Mfns Mfn2 is phosphorylated at Ser27 by c-Jun N-terminal kinase (JNK), which leads to recruitment of the ubiquitin ligase (E3) Huwe1/Mule/ARF-BP1/HectH9/E3Histone/Lasu1 to Mfn2 (Leboucher et al., 2012). As a consequence Mfn2 is ubiquitinated and subsequently degraded, leading to mitochondrial fragmentation and apoptotic cell death. Activity of JNK1 and JNK3 is inhibited by S-nitrosylation (Park et al., 2000), providing a mechanism to prevent Mfn2 ubiquitination and mitochondrial fragmentation. Enigmatically, exogenously added NOd appears to inhibit the S-nitrosylation of JNK3 induced by endogenous NOd during cerebral ischemia and reperfusion in rat hippocampus (Pei et al., 2008). In addition to Drp1 (see above), Parkin also acts as an E3 ubiquitin ligase for Mfn1 (Glauser et al., 2011). PINK1 phosphorylates Mfn2 and promotes its Parkinmediated ubiquitination and breakdown (Chen and Dorn,

2013). S-nitrosylation inhibits Parkin’s ubiquitin E3 ligase activity (Chung et al., 2004), which could inhibit Parkin-mediated Mfn1 breakdown and stimulate mitochondrial filamentation. This might provide a mechanism for RNS-induced stimulation of mitochondrial filamentation. PKA can be activated by disulfide bond formation (see above) and phosphorylates Mfn2 at Ser422. However, this phosphorylation appears not to affect mitochondrial morphology (Zhou et al., 2010). Other Links between Redox State and Mitochondrial Morphology Mitochondrial lipids like diacylglycerol (DAG), phosphatidylethanolamine (PE), phosphatidic acid (PA), and cardiolipin (CL) play an important role in mitochondrial morphology regulation (reviewed in Ha and Frohman, 2014). Peroxidation of CL is catalyzed by a CL-specific peroxidase of CL-bound cytochrome c (cyt-c; which shuttles protons from CIII to CIV; Kagan et al., 2005). CL peroxidation is essential for the release of proapoptotic factors from mitochondria into the cytosol (see Ren et al., 2014 and the references therein). With respect to mitochondrial morphology, evidence in yeast demonstrated that CL plays a role in the recruitment to mitochondria of fusionand fission-promoting proteins (see Ha and Frohman, 2014 and the references therein). dOH-induced peroxidation and carbonylation of MIM lipids and proteins, respectively, was linked to Dc depolarization (Iqbal and Hood, 2014). The latter Figure 3. Non-transcriptional Pathways Linking Redox Homeostasis to Mitochondrial Fission and Fusion Mitochondria are exposed to superoxide (O2d ) and nitric oxide (NOd) that are converted into hydrogen peroxide (H2O2) and peroxinitrite (ONOO ). Current literature and database analysis highlighted that these reactive species can induce nitration (green), S-nitrosylation (SNO; red) or affect disulfide bonds (S-S; blue). Drp1 and OPA1 (as well as proteinaceous binding partners of Drp1, OPA1, and Mfn2) are direct targets of these modifications, inducing functional alterations. Depolarization of Dc induced by the hydroxyl radical (dOH) was demonstrated to functionally affect OPA1 cleavage and thereby its function. Abbreviations and literature references are provided in the Supplemental Information.

Cell Metabolism 22, August 4, 2015 ª2015 Elsevier Inc. 7


Cell Metabolism

Perspective Figure 4. Interaction between Mitochondrial Dynamics, Redox Homeostasis, and Function Working model linking mitochondrial morphology, ROS/RNS production, and functional state (arrows a–e) in an interacting mechanism (Homeostatic circuit I). This circuit allows ROS/RNS signaling and is augmented by two additional mechanisms that suppress ROS/RNS levels (Homeostatic circuit II) and oxidative stress (Homeostatic circuit III). ROS/RNS from the cytosol, endogenous antioxidants, and various other parameters (lower box) exert a modulating effect on this system (see Regulation of Mitochondrial Morphology by Cellular Redox State for further details).

affects Dc-dependent OPA1 processing/function and mitochondrial morphology (Guillery et al., 2008). Regulation of Mitochondrial Morphology by Cellular Redox State Although the field is still at an early stage, we hypothesize here that mitochondrial morphology, redox homeostasis, and morphology are bi-directionally linked (Figure 4; arrows a–f). The main ROS/RNS involved in this system are H2O2, NOd, and ONOO . With respect to reactivity, dOH reacts very fast with virtually all biomolecules, whereas other ROS such as O2d , H2O2, and NOd are more selective, or display intermediate reactivity (ONOO ; Halliwell and Gutteridge, 2012). This chemical reactivity allows mitochondria-generated dOH and O2d to act more locally, whereas H2O2 and NOd can function as both a cytosolic and extracellular messenger, which generally induce reversible and irreversible atomic modifications at low and high levels, respectively (Winterbourn, 2008). Non-transcriptional regulation of mitochondrial fission/fusion proteins by ROS and RNS might provide the cell with a homeostatic mechanism (Figure 4; homeostatic circuit I) that sustains proper mitochondrial morphology, function, and redox homeostasis. It is highly likely that the balance between these entities depends on the cell type and experimental context. As a second line of defense against oxidative stress, it has been proposed that a mild increase in mitochondrial ROS production induces activation of long-lasting cytoprotective pathways by nuclear transcriptional changes. This process of ‘‘mitohormesis’’ is thought to protect cells against subsequent stress-inducing events (Yun and Finkel, 2014) and includes upregulation of endogenous antioxidant enzymes that maintain redox homeostasis (homeostatic circuit II) and avoid oxidative stress by reversing or preventing oxidative modification of DNA, proteins, and lipids (homeostatic circuit III). In this mechanism ROS/RNS generated outside the 8 Cell Metabolism 22, August 4, 2015 ª2015 Elsevier Inc.

mitochondrion can modulate homeostatic circuit I to exert control or trigger oxidative stress. Similarly, the properties of this circuit are influenced by various other parameters including substrate availability, oxygen pressure, environmental toxins, and/or the presence of pathogenic mutations (Figure 4). Mitochondrial fragmentation has been linked to mitochondrial inheritance during cell division and apoptosis and appears to be required for removal of dysfunctional mitochondria by (ROS-induced) mitophagy (Arnoult et al., 2005; Twig and Shirihai, 2011; Frank et al., 2012; Picard et al., 2013; Mishra and Chan, 2014). A fragmented mitochondrial phenotype is often observed during pathophysiological conditions in humans including PD/AD, Huntington’s disease, ischemia/reperfusion injury, drug-induced tissue damage, and inherited mitochondrial metabolic disorders (Archer, 2013). This suggests that reversal of mitochondrial fragmentation, possibly by genetic or pharmacological modulation of endogenous antioxidant levels or application of cell-permeable exogenous antioxidants (Figure 4), could be a valid therapeutic strategy in human disease (e.g., Koopman et al., 2007; Lackner and Nunnari, 2010; Blanchet et al., 2011, 2015; Distelmaier et al., 2012; Archer, 2013; Guo et al., 2013; Reddy, 2014). However, for such strategies to be successful, a detailed understanding of the various homeostatic circuits is required. In this respect, we expect that application of targeted (anti)oxidants, (targeted) ROS-generating molecules, and (targeted) ROS/RNS reporter molecules will be of great value to assess and modulate the levels of (local) ROS/RNS and their downstream effects (Skulachev, 2005; Forkink et al., 2010, 2014; Cocheme´ et al., 2011; Smith et al., 2011; Prime et al., 2012). Conclusion Accumulating evidence suggests that cellular and mitochondrial redox homeostasis is linked to mitochondrial dynamics in a celland tissue-dependent manner. During healthy and pathological conditions, mitochondria generate ROS and are exposed to RNS, which act as signaling and/or damaging molecules. Here we propose that (local) ROS and/or RNS regulate mitochondrial dynamics through acting on mitochondrial fusion and fission proteins. This would permit (auto)regulation of mitochondrial morphology and function by redox-mediated signaling.


Cell Metabolism

Perspective Understanding this mechanism is crucial to unravel the pathological events in the many human disorders where redox homeostasis, mitochondrial function, and structure are disturbed, as well as to develop successful rational intervention strategies. SUPPLEMENTAL INFORMATION Supplemental Information includes one table and can be found with this article online at http://dx.doi.org/10.1016/j.cmet.2015.06.006. ACKNOWLEDGMENTS We apologize to those authors whose articles we were unable to cite because of space limitations. This research was supported by the Centres for Systems Biology Research initiative (CSBR09/013V) of NWO (The Netherlands Organisation for Scientific Research) and an NWO-VENI grant (863.13.019) awarded to C.E.J.D. We thank Dr. J.J. Esseling and Dr. F. Distelmaier (RIMLS, Department of Biochemistry 286) for performing microscopy experiments. REFERENCES Anand, R., Wai, T., Baker, M.J., Kladt, N., Schauss, A.C., Rugarli, E., and Langer, T. (2014). The i-AAA protease YME1L and OMA1 cleave OPA1 to balance mitochondrial fusion and fission. J. Cell Biol. 204, 919–929. Archer, S.L. (2013). Mitochondrial dynamics—mitochondrial fission and fusion in human diseases. N. Engl. J. Med. 369, 2236–2251. Arnoult, D., Rismanchi, N., Grodet, A., Roberts, R.G., Seeburg, D.P., Estaquier, J., Sheng, M., and Blackstone, C. (2005). Bax/Bak-dependent release of DDP/TIMM8a promotes Drp1-mediated mitochondrial fission and mitoptosis during programmed cell death. Curr. Biol. 15, 2112–2118. Auche`re, F., and Rusnak, F. (2002). What is the ultimate fate of superoxide anion in vivo? J. Biol. Inorg. Chem. 7, 664–667. Bak, D.W., and Weerapana, E. (2015). Cysteine-mediated redox signalling in the mitochondria. Mol. Biosyst. 11, 678–697. Barsoum, M.J., Yuan, H., Gerencser, A.A., Liot, G., Kushnareva, Y., Gra¨ber, S., Kovacs, I., Lee, W.D., Waggoner, J., Cui, J., et al. (2006). Nitric oxide-induced mitochondrial fission is regulated by dynamin-related GTPases in neurons. EMBO J. 25, 3900–3911.

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Collapse of redox homeostasis during aging and stress.

Circadian clock proteins regulate neuronal redox homeostasis and neurodegeneration.

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