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Logo of nihpaAbout Author manuscriptsSubmit a manuscriptHHS Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
Biochim Biophys Acta. Author manuscript; available in PMC 2010 November 1.
Published in final edited form as:
PMCID: PMC2738971

SR/ER-mitochondrial local communication: Calcium and ROS


Mitochondria form junctions with the sarco/endoplasmic reticulum (SR/ER), which support signal transduction and biosynthetic pathways and affect organellar distribution. Recently, these junctions have received attention because of their pivotal role in mediating calcium signal propagation to the mitochondria, which is important for both ATP production and mitochondrial cell death. Many of the SR/ER-mitochondrial calcium transporters and signaling proteins are sensitive to redox regulation and are directly exposed to the reactive oxygen species (ROS) produced in the mitochondria and SR/ER. Although ROS has been emerging as a novel signaling entity, the redox signaling of the SR/ER-mitochondrial interface is yet to be elucidated. We describe here possible mechanisms of the mutual interaction between local Ca2+ and ROS signaling in the control of SR/ER-mitochondrial function.

Keywords: superoxide anion, H2O2, IP3 receptor, ryanodine receptor, SERCA, bioenergetics, apoptosis

1. SR/ER-mitochondrial structural relationship: the platform for local signaling

The cytoplasm is a highly restrictive medium for the diffusion of charged compounds and ions. Thus, intracellular signaling systems utilizing ionic messengers frequently operate via local communication between the sources and targets of the messenger molecules [1, 2].

The SR/ER and mitochondria are probably the two most extensive intracellular membrane systems. The SR/ER takes shape as a luminally continuous reticular network [3], while the mitochondria exist as an array of individual organelles that dynamically change their shape and size via fusion and fission capable of forming luminally continuous regional networks [4]. Due to their abundant presence throughout the cytoplasm and dynamic nature, close encounters between SR/ER and mitochondria are predictable. Indeed, close SR/ER-mitochondrial contacts, where the interorganellar gap is ~10–50 nm have been documented in different tissues [5, 6]. However, it has turned out that the contacts are not just stochastic ‘collisions’ but are sturdy junctions secured by protein tethers [68]. SR/ER fragments remain associated with the mitochondria after fractionation [6, 9], allowing the isolated SR/ER-mitochondrial complexes to sustain local interorganellar Ca2+ communication [6, 10] and collaboration in phospholipid biosynthesis [11].

The hunt for the specific proteins that contribute to the SR/ER mitochondrial linkage has recently intensified. The heterogeneous length of the ER-mitochondrial tethers in non-muscle cells, suggests that several proteins might participate in linking the organelles together [6]. Biochemical and functional measurements have implicated a handful of proteins/protein complexes as ER-mitochondrial coupling elements [DLP-1/DRP1-1[12, 13], autocrine motility factor receptor (AMFR) and BAP-31 [14], PACS2 [15], grp75, a chaperone forming a complex with type 1 voltage-dependent anion channel (VDAC1) and type 1 IP3 receptor (IP3R1) [16], type 3 IP3R (IP3R3) [17] and mitofusin-2 (Mfn2) [18]]. Furthermore, the main Ca2+ conducting channels of both SR/ER (ryanodine receptors (RyRs) and IP3Rs) and outer mitochondrial membrane (VDACs) form multimolecular complexes (a few protein partners are listed in Figure 1) that can also interact with adjacent membranes. This mechanism might be particularly relevant for the IP3Rs that seem to be concentrated at the ER-mitochondrial interface [19] but dispensable for RyR2s that concentrate on the sarcolemmal side the SR in the heart [5]. Additional physical support of the SR/ER mitochondrial associations is provided by cytoskeletal proteins, since both SR/ER and mitochondria bind to microtubules/actin filaments (for a recent review see [20]). Thus, the overlapping spatial arrangements of SR/ER and mitochondria and the interorganellar physical linkage provide a robust platform for local communications between the organelles. However, it is important to note that both SR/ER and mitochondria are continually reorganized, indicating that the SR/ER junctions likely form dynamic structures.

Figure 1
Local calcium signaling between the SR/ER and mitochondria

2. Local calcium signaling: sources and targets and their modulators

Both SR/ER and mitochondria can accumulate Ca2+. The Ca2+ storage capacity of the SR/ER is limited, but its luminal [Ca2+] ([Ca2+]ER) is close to the millimolar range, promoting the protein folding/processing functions [3, 21]. Due to the 3–4 orders of magnitude [Ca2+] gradient across its membrane, to the presence of messenger-operated Ca2+ release channels, to the robust Ca2+ pumping mechanism and to its cell-wide distribution, the SR/ER can serve the generation of rapid cytoplasmic [Ca2+] ([Ca2+]c) transients. On the other hand, the low-affinity, high-capacity and Ca2+-gated Ca2+ uptake mechanism enables mitochondria to respond effectively to local calcium signals.

2.1. SR/ER Ca2+ accumulation

is established via the SERCAs. Out of the seven isoforms usually found in mammalian cells, the predominant subtype in non-muscle cells is SERCA2b, in skeletal muscle SERCA1a and in cardiac muscle SERCA2a. The activity of these high affinity (2b>1a~2a) and velocity (1a>2a>2b) Ca2+ pumps is regulated by substrate/product levels ([ATP]/[ADP], [Pi]), pH, [Ca2+]c and [Ca2+]ER, by regulatory proteins (eg. phospholamban, sarcolipin) and by the redox state of critical reactive thiols [22, 23]. The luminal Ca2+ regulation of SERCA2b involves the ER chaperone ERp57 that is also a target for ROS/redox regulation (see under ROS targets: SERCA).

2.2. SR/ER Ca2+ release

during [Ca2+]c spikes occurs through the RyR and IP3R, which are non-specific cation channels. Both RyRs and IP3Rs exist as tetramers of subunits that contain large N-terminal cytoplasmic domains and luminal loops engaging in numerous regulatory interactions with other proteins [2426]. RyRs mediate Ca2+-induced Ca2+ release from the SR of skeletal (RyR1) and cardiac (RyR2) muscle in the process of excitation contraction coupling or from the SR/ER of smooth muscle, diaphragm and non-muscle cells (predominantly RyR3) [27]. For IP3Rs three different isoforms have been identified, and most cells appear to express multiple isoforms.

RyRs are subject to both cytosolic and luminal Ca2+ regulation. In principle, Ca2+ conductance is directly related with both the local [Ca2+]ER and [Ca2+]c and the Ca2+ regulation is established via proprietary Ca2+ binding domains and via interactions with Ca2+ binding proteins eg. calsequestrin/triadin/junctin complex [25, 28, 29]. Activation of the IP3Rs is also controlled by Ca2+ but it requires the binding of IP3, which is directly modulated by the competitive ‘pseudoligand’ IRBIT [30]. In addition to channel opening, IP3 binding leads to time-dependent inactivation [31]. The [Ca2+]c regulation of IP3Rs is biphasic at physiologically occurring [Ca2+]c ranges with isotype and species-dependent differences in the stimulatory and inhibitory range [24, 30]. [Ca2+]ER regulation of the IP3R involves Ca2+ binding and redox-dependent chaperones including calnexin and ERp44 (see later in more details) [32]. Besides Ca2+ regulation, the cytosolic domain of both RyR and IP3R provides a scaffold to a host of regulatory protein interactions giving rise to macromolecular complexes able to receive inputs from most of the major signaling pathways and to sense the metabolic status of the cell [2426, 29, 30, 33, 34].

Due to the Ca2+-induced activation of both RyR-and IP3R, these channels operate more effectively in clusters than being stochastically dispersed along the SR/ER membranes. Indeed, cardiac and skeletal RyRs are concentrated in dense arrays mostly to the SR interfaces with the T-tubules (junctional SR) [5]. This orderly distribution is ensured by multiple accessory proteins, including FKBP12.6 and is critical for the synchronized activation of the sarcomeres over the excitation-contraction coupling [25]. IP3Rs are also distributed in groups that function as distinct release units [35] and display enhanced clustering when activated [36, 37]. The exact mechanisms of IP3R positioning and group formation are still yet to be clarified ([30]). Recent data suggest higher prevalence of Ca2+ release units [38] and chaperone-mediated (Sigma-1 receptor) protection of activated IP3R3s against proteasomal degradation [17] at the ER-mitochondrial close contacts.

2.3. Mitochondrial Ca2+ uptake

Cytoplasmic Ca2+ has to cross both the outer and inner mitochondrial membranes (OMM, IMM) to enter the matrix. The predominant pathways for Ca2+ diffusion across OMM are the VDACs. Because of their high density and high conductance, VDACs allow rapid Ca2+ diffusion [39]. Still, VDACs may become a Ca2+ transport limiting factor during delivery of microdomain-derived Ca2+ signals to the mitochondria [40, 41]. Evidence has been presented that the VDAC-mediated Ca2+ and solute flux is stimulated by Ca2+ [42, 43]. Additional regulatory mechanisms of the Ca2+ flux are likely to result from the VDAC’s phosphoregulation and direct interaction with several proteins. The primary driving force for Ca2+ entry into the mitochondrial matrix is the inside-negative membrane potential across the IMM maintained by the proton-extruding activity of the electron transport chain (ETC). In most cellular and subcellular paradigms, mitochondrial Ca2+ uptake becomes apparent when the [Ca2+]c is increased to ≥1µM, though [Ca2+]m elevations have also been observed during submicromolar [Ca2+]c increases in some cases [20]. This may be supported by Ca2+-induced allosteric activation of the mitochondrial Ca2+ uptake [44, 45]. The molecular identity of none of the IMM Ca2+ transport mechanisms is known. The most recognized mechanism via which Ca2+ enters the mitochondrial matrix is the Ca2+ uniporter [46]. The uniporter has been functionally characterized by patch-clamping mitoplasts as a ruthenium red-sensitive and highly selective inward-rectifying low-affinity Ca2+-gated Ca2+ channel [47, 48]. Recently, uncoupling proteins 2 and 3 have been implicated in the Ca2+ uniport mostly based on RNA silencing/rescue experiments but this finding remains a subject of discussion [49, 50]. Besides the uniporter, a rapid uptake mode and a mitochondria-located RyR (mtRyR1) have been proposed to participate in the Ca2+ flux across the IMM in cardiomyocytes and neurons but their physiological relevance and regulation is yet to be confirmed [5153]. An important feature of the mitochondrial Ca2+ uptake is the time-dependent sensitization and desensitization observed in different cellular systems (sensitization: [45, 54]; desensitization: [5557]) that might have profound role in establishing the relationship between the periodicity and efficacy of mitochondrial delivery of [Ca2+]c oscillations. The mechanism underlying sensitization or desensitization of mitochondrial Ca2+ uptake is yet to be clarified; currenly proposed paradigms involve Ca2+/CaM for the sensitization [45], PKC-ε [58], p38 MAPK [59] and the H+-flux through the F1F0-ATPase as a positive regulator [60] in the process of desensitization.

2.4. Ca2+ targets and Ca2+ buffering in the mitochondrial matrix

The [Ca2+]m increase enhances oxidative metabolism via activating Ca2+ sensitive matrix dehydrogenases (CSMDH) [6163] that feed reducing equivalents to the ETC and the F1F0-ATPase itself [64]. Therefore, the RyR/IP3R-mediated [Ca2+]m signal is rapidly followed by an increase in mitochondrial [ATP] [65, 66]. With the increased ETC activity mitochondrial superoxide (·O2) generation also increases and as a compensatory mechanism, Ca2+ also activates ·O2- scavenging enzymes (SOD2). However, after a point, the [Ca2+]m increase activates the opening of the permeability transition pore (PTP), allowing indiscriminate exchange of ions and small solutes between the mitochondrial matrix and the cytosol. The PTP opening dissipates mitochondrial membrane potential, might cause swelling and release of apoptotic factors from the intermembrane space (IMS) and initiation of the mitochondria-dependent apoptotic cascade [67]. Importantly, mitochondria can hold elevated [Ca2+]m steadily below the toxic level even during extended mitochondrial Ca2+ accumulation by means of a Pi-dependent dynamic Ca2+ buffering/precipitating system [68, 69]. PTP opening seems to occur when Ca2+ uptake is very fast or when massive amounts of Ca2+ are accumulated.

2.5. Ca2+ extrusion from the mitochondria

is carried out mostly via cation exchangers [3Na+/Ca2+-exchanger (NCX), 3H+/Ca2+-exchanger] and perhaps transient openings of the PTP. The Na+-independent pathways display increased activity under oxidizing conditions in the mitochondrial matrix perhaps as a defense mechanism against the [Ca2+]m rise-associated enhancement of ROS production [70].

2.6. Local SR/ER-mitochondrial Ca2+ dynamics (Figure 1)

Depending on the location of the RyR and IP3R, the released Ca2+ can enter directly the SR/ER-OMM cleft or can spread in by diffusion when released via nearby channels. The former scenario is likely if the IP3R and VDAC are physically coupled [16]. However, only the latter mechanism is plausible in skeletal and cardiac muscle, where RyRs are clustered at the triadic/dyadic clefts instead of the SR/OMM interface. Both mechanisms are competent to expose the mitochondrial Ca2+ uptake sites to high local [Ca2+]c since RyR/IP3R-dependent calcium signal propagation to the mitochondria could persist under [Ca2+]c clamping using a slow Ca2+ chelator (EGTA) in permeabilized cardiac and non-muscle cells [7173]. Also, single mitochondrial [Ca2+]m elevations (“Ca2+ marks”) have been detected in cardiac cells during “elementary” Ca2+ release events (sparks) that fail to significantly elevate the global [Ca2+]c [74]). Nevertheless, synchronous activation of a large number of IP3Rs delivers Ca2+ to the mitochondria more effectively than random openings [19]. During [Ca2+]c oscillations that peak at ≤1 µM in the bulk cytosol, the mitochondrial side of the interface can be exposed to >100µM [Ca2+] if the RyRs/IP3Rs are also in the interface area, and to ~10µM [Ca2+] if the RyRs/IP3Rs are located within 100nm from the contact region.

Disruption of the ER-mitochondrial physical linkage by limited proteolysis uncoupled mitochondria from IP3-induced Ca2+ release [6] and genetic interference with the tethering proteins grp75 and Mfn2 also reduced the efficacy of IP3R-dependent [Ca2+]m signal generation [16, 18]. Conversely, strengthening of the ER-mitochondrial linkage by synthetic linkers and Ca2+-mediated inhibition of mitochondrial motility close to the sites of Ca2+ mobilization led to more effective IP3R-mitochondrial Ca2+ transfer [6, 75]. Notably, Ca2+ has been shown to affect several aspects of ER and mitochondrial dynamics, including mitochondrial motility and fission and ER motility and restructuring, which may also mediate Ca2+-dependent changes in the ER-mitochondrial coupling [7582]. Once Ca2+ entered the mitochondrial matrix, [Ca2+]m increases and Ca2+ removal mechanisms are activated to restore the ‘resting state’. Mitochondrial Ca2+ efflux might locally support ER Ca2+ reloading [83]. Hyperactivity of Ca2+ extrusion might attenuate the beneficial stimulatory effect of the [Ca2+]m rise on oxidative ATP production as it has been observed in association with increased [Na+]c in failing cardiomyocytes [84, 85].

3. Local ROS signaling: sources, targets and mediators

Enhanced production or external exposure to ROS and reactive nitrogen species (RNS) has been proven hazardous to cells causing various types of damage and accelerating cell death or senescence via multiple mechanisms collectively known as oxidative/nitrosative stress. However, a multitude of molecules participating in different cellular signaling or regulatory networks have redox active moieties (e.g. cysteine, methionine, tyrosine) at functional regions endowing them a ROS/RNS-sensor feature. Consequently, endogenously generated ROS and RNS have been emerging not just as harmful products but diffusible modulators of a range of different signaling cascades and even more, as messengers themselves [23, 86]. Many ROS/RNS sources and sensors are localized in the mitochondria or SR/ER and are relevant for calcium signaling. In this review, we focus on ROS as local messengers between the SR/ER and mitochondria (for a review on RNS see [87]). The main ROS sources and targets and pathways of ROS dynamics are depicted in Figure 2.

Figure 2
ROS dynamics and potential ROS signaling targets in the SR/ER and mitochondria

3.1. ROS sources

In principle, ROS are products of consecutive stepwise reduction of molecular oxygen:


where enzyme E1-3 represent the catalyzing enzymes. When E3 are saturated and Fe2+ is available, H2O2 is reduced to H2O via non-enzymatic Fenton reaction yielding a hydroxyl radical OH., the most aggressive ROS:_


E1 represents mitochondrial and non-mitochondrial oxidases, out of which the most significant ·O2 sources are complex I and III of the electron transport chain (ETC) and NADPH oxidases associated with the plasma membrane or the SR/ER ([88, 89]). In the highly reducing environment of mitochondria, a number of potential e donors are capable of reducing O2 to ·O2. However, it is mostly the e leak via complexes I and III that cannot be completely overcome by the powerful mitochondrial ROS scavenging mechanisms ([90]). The relative contribution of respiratory complex I and III to the mitochondrial ·O2 generation appears to be cell/tissue type-and respiratory status-dependent. In cardiac muscle, with fully respiring (state 3) mitochondria and in the lung the ubiquinone (Q) cycle of complex III is the primary source of ·O2 generation. Since autooxidation of Q can happen at either side of IMM, it can increase ·O2 in both the matrix and the IMS. In the brain and in state 4 respiration (absence of ADP) the NADH oxidase complex I is the dominant ·O2 source at the inner side of IMM ([90]). Importantly, complex I has been reported to work as a redox sensor, having highly reactive thiol groups that become S-gluthationylated when GSH/GSSG ratio decreases (e.g. under oxidative stress) causing decreased e flow to complex III but increased ·O2 production [91, 92]. The rate of mitochondrial ·O2 production is determined in principle by simple mass action with an inverse relation to the electron flow over the ETC and with a direct proportionality with the available O2 and electron donor (R·).


In vitro, about 1–4% of the O2 entering the ETC is reduced to ·O2 [88, 90], though some groups have published even smaller fraction (~0.1% [94, 95]). The activity of the tricarboxylic acid cycle (TCA), which provides a vast source of e to the ETC in the form of NADH is a crucial determinant of the rate of mitochondrial ·O2 production.

The highly regulated TCA is an ideal platform to connect mitochondrial ·O2 production with signaling systems; in particular with Ca2+ signaling via the CSMDH. In addition, the [4Fe-4S] iron-sulfur cluster of aconitase is reversibly inactivated by ·O2 (to [3Fe-4S]) serving as a ‘rheostatic’ limit switch on oxidative metabolism and fuel (ATP) production that keeps radical ‘byproducts’ in a non-toxic range [96, 97]. Frataxin, an iron chaperone protein mutated in Friedrich’s ataxia appears to play an important role in maintaining the reversibility of the oxidation of the iron-sulfur cluster of aconitase [98]. Importantly, if sufficient glutamate is present, the activity of aconitase will not limit the NAD(P)H output of the TCA as α- ketoglutarate dehydrogenase (α-KGDH), one of the Ca2+-sensitive rate-limiting enzymes of the TCA cycle can bypass the segment between oxaloacetate and α-KG [99]. Nevertheless, α-KGDH is also inhibited by ROS but with less sensitivity than aconitase. Moreover, the α-KGDH complex can generate significant amount of ·O2 upon excess in [NADH] [100102] making α-KGDH a main regulatory node for mitochondrial energy metabolism and ROS production. ROS regulation of and ROS production by α-KGDH appears to be particularly significant in neuronal cells and is suspected to contribute to ROS-related neuronal degeneration [102, 103].

NADPH oxidases (Nox) have been recognized as the ‘superoxide guns’ of phagocytic immune cells to kill bacteria but later other isoforms of the enzyme have been discovered in the plasma membrane and SR/ER of non-phagocytic cells [104, 105]. It has been recently suggested that the ·O2 generation by an SR-associated Ca2+-sensitive Nox is regulated by cADP-ribose activation of RyR in coronary artery smooth muscle cells and functions as a local calcium signal amplifier [106, 107]. Along this line, an SR-associated NADH oxidase has been found in skeletal muscle that produced ·O2 upon substrate addition and was proposed to be responsible for the sensitization of RyR1 by NADH [108]. However, thus far there is no experimental evidence that ROS derived from SR/ER-resident Nox would affect mitochondria.

Nitric oxide synthase (NOS) is present in various organelles, including the plasma membrane, SR/ER and mitochondria. NOS is an NADPH/FAD oxidoreductase that produces ·NO from the substrate L-arginine. This process requires the co-factor (6R)-5,6,7,8,-tetrahydrobiopterin (BH4). In the absence of the substrate or co-factor, the oxidoreductase function becomes uncoupled from ·NO generation and produces ·O2 [87, 109]. A recent paper indicates that a mitochondrial NOS can be activated by Ca2+ and when uncoupled, it may promote opening of the PTP and apoptosis in cardiomyocytes [110].

Direct H2O2 generation: Besides ·O2 dismutation, H2O2 can also be directly generated in the mitochondria. The 66kDa isoform of the growth factor adapter Shc (p66Shc) has been recently recognized as a key promoting factor in mammalian aging [111]. Upon oxidative stress, p66Shc accumulates to the IMS via a PKCβ-and PIN1 (a peptidyl prolyl isomerase) dependent manner where it displays cytochrome c oxidase activity generating H2O2 and so promotes IMM permeabilization and mitochondria-dependent apoptosis [112] [113]. The OMM-bound monoamino oxidase (MAO) is another local source of H2O2 as a byproduct of the oxidation of biogenic amines and is believed to play a significant role in the mitochondrial damage and PTP activation in several neurological disorders [114].

3.2. ROS scavengers

[ROS] in the intracellular compartments is tightly controlled via scavenging mechanisms to maintain subtoxic levels (Fig 2). ·O2 scavenging is established via dismutation to H2O2 by superoxide dismutases with Mn2+ (SOD2 in the mitochondrial matrix) or Zn2+/Cu2+ (SOD1 in the IMS and cytosol) centers. H2O2 scavenging is established via reduction to H2O using glutathione dependent and independent pathways; the former one being dominant. Both pathways utilize NADPH as e donor provided by the pentose phosphate cycle (P5P in Fig2) in the cytosol or by transhydrogenation from NADH in the mitochondria [115]. Thus, maintaining the level of reducing equivalents in the mitochondrial matrix is critical for both, the oxidative ATP production as well as the balance between oxidized and reduced thiols in the mitochondria. In the glutathione dependent pathway, gluthatione reductase (GR) transfers the electron from NADPH to oxidized glutathione (GSSG) keeping most of it in the reduced state (GSH) under physiological conditions. GSH can then transfer its reducing equivalent to glutathione peroxidase (GPx) that catalyses the reduction of H2O2 to water [115]. Notably, glutathione is synthesized in the cytosol and it is transported into organelles via glutathione transferases. Mitochondrial glutathione is 10–15% of the total cellular glutathione, but its concentration is similar to the cytosolic one (5–10 mM in rat liver) [116].

The glutathione-independent H2O2 scavenging system transfers the electron from NADPH to H2O2 an electron transport chain comprised of thioredoxin reductase (TrxR1 cytosol, TrxR2 mitochondria), thioredoxin (Trx1 and Trx2) and peroxiredoxin (Prx1 and Prx2) in the order TrxR→Trx→Prx→ H2O2. This system has less scavenging power than the glutathione-dependent system; however, the redox state of Trx limits its capability of recovering S-glutathionylated proteins (see below) [23, 90, 115]. Another powerful H2O2 scavenging enzyme is catalase that is predominantly confined to peroxisomes [117] and as such, is probably less relevant in the ER-mitochondrial local redox signaling.

Importantly, ·O2 also readily reacts with nitric oxide (NO) when present and NO· has been even suggested as a ·O2 scavenger in the late ‘80s [118]. However, the reaction product peroxinitrite (ONOO) anion has been proven highly reactive, bearing more oxidative power than its parent molecules; and even more, its solute interactions may further produce aggressive radicals (e.g. hydroxyl and carboxyl radicals) [87]. Moreover, ONOO is membrane permeant thus easily crosses compartmental barriers. ONOO is the center figure in nitrosative stress and has been progressively ‘claiming’ responsibility for effects attributed to ROS. Thus, a part of oxidative effects of ·O2 is actually mediated by ONOO.

3.3. ROS dynamics

Because of its negative charge and high reactivity (redox potential against H2O2 Eh~ +0.94V), ·O2 cannot permeate through lipid membranes and the bulk of it is rapidly dismutated to H2O2 by SODs. The steady state [·O2] in the mitochondrial matrix has been estimated being in the range of 100–200pM based on calculations considering ·O2 production by the ETC and consumption by SOD2 in isolated liver and heart mitochondria [119]. ·O2 can leave mitochondria via anion channels (the VDAC in the OMM [120] and, perhaps through the inner membrane anion channel (IMAC) and VDAC as part of the peripheral benzodiazepine receptor complex at the contact points of IMM and OMM [121, 122]) or via the PTP. Because of its electroneutrality and relatively moderate reactivity (Eh~ +0.38V) H2O2, is considered freely permeable to biological membranes (although see [123, 124] for potential limitations) and thus widely recognized as a diffusible messenger in ROS-associated redox signaling. Thus, the combination of the two ROS intermediates enables both tightly confined short-lasting (microdomain) operations as well as longer lasting more distant signaling actions [125].

In summary, mitochondrial ·O2 production is tightly controlled at membrane-bound main sources utilizing ‘rheostatic’ feedback loops on the reducing equivalent supply (aconitase, αKGDH and powerful clearance/buffer systems (SOD2, GPx) to secure a subtoxic basal setpoint. Both, ·O2 production and clearance are regulated in part, by Ca2+ and thus can be targeted by calcium signaling [126]. The recent visualization of ·O2 ‘flashes’ in single mitochondria indicates that ·O2 in individual mitochondria can be rapidly and independently regulated [127]. However, visualization of cell-wide synchronized oscillations and waves of mitochondrial ROS has also provided evidence for regenerative mechanisms of propagation [122, 128, 129]. Expansion of the local ROS events is likely to utilize rapid “metabolism” of ·O2 to H2O2 that spreads effectively among the individual mitochondria. In contrast with Ca2+, the ROS ‘messengers’ are consumed by their targets.

3.4. ROS targets

The most common molecular targets of ROS are the Cys thiol groups. Protein Cys thiols can be divided into four groups based on their prevalent interactions, which are determined by their immediate intra-and perimolecular environment: those that form permanent structural disulfide bonds, those that coordinate metals, those that are permanently in the reduced state, and those that are reversibly oxidized [130]. The latter group is often referred to as (hyper)reactive thiols and plays the lead role in the oxidative/redox regulation of proteins. Upon exposure to ROS, these reactive thiols may undergo multiple reversible and irreversible oxidative alterations forming sulfenic (R-SOH), sulfinic (R-SO2H) and sulfonic (R-SO3H) acid residues. The formation of the sulfenic goup is reversible but this residue is an instable intermediate that quicly becomes further oxidized to the stable sulfinic or sulfonic groups or forms intra/intermolecular disulphide bonds (R-SS-R) or mixed disulfide bond with GSH (S-gluthationylation→ R-SSG) [115]. The latter pathway dominates under physiological coditions in the cytoplasm because of the abundance of cytoplasmic reduced glutathione (GSH:GSSG~100:1 at millimolar concentrations). Formation of the more oxidized sulfinic and sulfonic acid residues is generally irreversible with the only known exception being the H2O2 scavenger peroxiredoxin that after being oxidized and inactivated by H2O2 (via sulfinylation) is slowly recovered by a novel ATP-dependent enzyme sulfiredoxin [131]. On the other hand, depending on the overall redox tone, the reactive Cys residues of the S-gluthationylated proteins can be reduced back enzymatically by glutaredoxin or thioredoxin. Thus, proteine S-glutathionylation is a reversible oxidative alteration of the reactive Cys thiols triggered by ROS production and controlled by redox enzymes and small molecular weight thiols. As such, it can function as the premier effector pathway in ROS signaling [23, 115, 132]. Indeed, most of the reversible functional effects of ROS on enzymes, ion transporting proteins and other factors involved in intracellular signaling is associated with thiol oxidation [23, 132, 133].

ROS targets can be arbitrarily divided into two groups: ROS scavengers comprising of enzymes that catalyze ROS reduction to water, and ROS effectors that alter their function owing to interaction with ROS. Some ROS scavengers (e.g. GPx4) also interact with ROS effectors and regulate their thiol oxidation. GPx4 (an isoform present both in the cytosol and mitochondrial matrix) has been proven a vital molecular decoder of glutathione-dependent redox signaling as its deletion is embryonic lethal (killing mouse embryos at the same developmental stage as GSH deficiency) [134, 135]. Via metabolizing lipid hydroperoxides (e.g. cardiolipin hydroperoxide in the mitochondria) and counteracting with lypoxygenases as well as cyclooxygenase, GPx4 exerts anti-inflammatory and anti-apoptotic actions [116, 136]. Trx and Grx are also important modulators in the glutathione-dependent redox signaling as they regulate the duration and degree of reversible S-glutathionylation of target signaling molecules [115, 116].

Cell-wide molecular targets of ROS are reviewed elsewhere [23], here we focus on ROS effectors that participate in the local Ca2+ homeostasis between the ER/SR and mitochondria:

SERCA pumps contain numerous free Cys residues (24 in SERCA1, out of which 6 displayed sensitivity to oxidation by peroxynitrite [137]). The reactive thiol groups and disulfide bonds of SERCA affect the pump activity differently, according to their position, accessibility and reactivity [23]. Cys674, a low-pKa reactive thiol at the hinge region of the protein [138] appears to play a key and dual role in the oxidative regulation of SERCA. Mild oxidative conditions lead to S-glutathionylation of Cys674, and increase pump activity, which might be a significant contributor to NO-dependent vasodilatation [139]. Extensive or extended exposure to oxidants that lead to oxidation of other Cys residues and sulphonylation of Cys674 irreversibly inhibit the pump [139141]. The distinct functional changes of the SERCA evoked by mild and harsh oxidative conditions illustrate that graded ROS elevations can serve both as a signaling event and a harmful stress condition acting through a single protein. In addition, the luminal regulation of SERCA2b also appears to involve a redox component: its C-terminal tail and 4th luminal loop forms a complex with the Ca2+-binding chaperone calreticulin and the ERp57 oxidoreductase, respectively, in a [Ca2+]ER and redox dependent manner. At high [Ca2+]ER and oxidizing conditions ERp57 promotes disulfide bridge formation between the L4 thiols that reduces pump activity, while when [Ca2+]ER decreases to <50 µM, ERp57 dissociates from L4 leaving the –SH groups reduced and increasing pump activity [142]. This mechanism increases the energy-efficiency of SERCA2b and protects from Ca2+ depletion-induced ER stress. Thus, ROS production by adjacent mitochondria is likely to facilitate ER store Ca2+ refilling by SERCA pumps but massive ROS generation would engage Ca2+ depletion-induced ER stress that may synergize with other effectors of ROS to initiate apoptosis.

RyR/IP3Rs contain multiple reactive Cys thiols that influence channel gating or assembly. Moreover, many of the regulatory proteins that form complex with the RyR/IP3R also have reactive thiols that affect their interaction with RyR/IP3R [24, 33, 143]. RyR1: out of the three RyR isoforms, RyR1 have the most reactive thiols: 29 free –SH groups out of the 100 Cys moieties in one subunit [144]. Thiol oxidation in general increases channel activity via enhancing intersubunit binding and preventing the binding of the negative regulator calmodulin (both apoCaM and CaCaM) to the receptor [145]. The reactive Cys thiols of the RyR1 act as both cytosolic and luminal (regulating the interaction with triadin) redox sensors [146, 147]. Apparently, the different reactive groups have different sensitivity to oxidation, S-nitrosylation and S-glutathionylation, enabling the RyR1 to distinguish among different redox inputs [23]. RyR2 carries 21 reactive thiols out of 89 Cys moieties per subunit [148]. Similarly to RyR1, the net effect of thiol oxidation is increased/sensitized channel activity [29, 149]. Sensitization of RyR3 by ROS is relevant for the glutamate-induced mitochondrial ROS-dependent long-term potentiation in hippocampal neurons [150].

IP3R On IP3R1 (monomer), ~70% of the 60 Cys residues are kept in reduced state with variable accessibility and variable regulatory significance [151]. Oxidation of certain –SH groups via exposure to thimerosal or GSSG sensitizes IP3R activation by IP3; however, oxidation of other thiol groups, which can be accessed by mersalyl but not the other thiol reagents, sensitize only the agonist binding but inhibit channel function [151, 152]. The sensitizing effect to the agonist has also been reproduced in bilayer systems using purified reconstituted receptors [153155].

IP3-induced ER Ca2+ mobilization can be sensitized with the use of thiol-oxidizing agents (e.g. thimerosal, GSSG) and this sensitization alone is sufficient to generate spontaneous Ca2+ release events and [Ca2+]c oscillations [156, 157]. In addition to the pharmacological thiol reagents, endogenous ·O2 derived from NADPH oxidase activity has also been shown to amplify IP3R-mediated calcium signaling via sensitization of the IP3- induced Ca2+ release in aortic endothelial cells [158]. Also, pharmacological evidence has suggested that mitochondrial ·O2 production is necessary to maintain IP3R-linked [Ca2+]c oscillations in pancreatic acinar cells [159]. Exogenous ROS derived from macrophages augmented IP3R-mediated Ca2+ signaling and Ca2+-dependent apoptosis in pulmonary microvascular endothelial cells, which mechanism might be of relevance in development of atherosclerotic lesions [160]. Similarly, externally added H2O2 enhanced IP3R-dependent GABA-ergic synaptic activity in spinal cord nociceptive interneurons [161]. It has been also suggested that ROS might interfere with IP3 degradation since in vascular SMC the enhancement of IP3-induced Ca2+ release by ·O2 donors could not be observed when nonhydrolysable IP3-analogues were used [162].

RyR/IP3R-associated proteins: FKBP12.6 has been reported to stabilize the RyRs and synchronize their function [25]. FKBP12 association with both RyR2 and RyR1 is sensitive to H2O2, involving a specific thiol group on the RyR-FKBP interaction site [163]. ERp44, a thioredoxin family protein previously known to be involved in oxidative protein folding via reinforcing the Ero1α/oxidoreductase system has been shown to directly interact with the reduced Cys residues on the 3rd luminal loop (L3V) of IP3R1 in a Ca2+ and redox state dependent manner and inhibit the channel [32]. This inhibitory effect has been proposed to support ER Ca2+ refilling and in turn, to protect protein folding when the redox balance in the ER shifts toward reducing conditions. Since silencing experiments revealed a basal inhibition of IP3R1 by ERp44 [32], one could expect that exposure to ROS and oxidizing the free –SH groups in the L3V luminal loop would be a way to enhance IP3R1 activity.

The ROS sensitivity of both RyR and IP3R indicates that feedback amplification of the Ca2+ release at the SR/ER mitochondrial interface might be induced by mitochondria-derived ROS that is induced by the calcium signal propagation to the mitochondria. This mechanism can be particularly important during pathological conditions when the PTP opens and the discharge of the ER Ca2+ pool further supports rapid execution of the cell [3].

Other targets at the SR/ER-mitochondrial interface: Cytoplasmic kinases and phosphatases are among the most established ROS-dependent regulators of the SR/ER and mitochondrial Ca2+ transporting proteins. Practically all the major cytosolic kinase signaling pathways are modulated by ROS [164] but here we discuss only the cAMP-dependent kinases (PKA) and the Ca2+/calmodulin-dependent kinases (CaMK) that can associate with the Ca2+ transporting proteins via anchoring proteins (AKAPs [165, 166]) or CaM. Moreover, SR/ER Ca2+ release channels can form macromolecular complexes with phosphatases PP1, PP2A [165, 167, 168]. Kinases and their counteracting phosphatases have different sensitivities to ROS modulation (mostly inhibition) and often the phosphatases are the more sensitive targets. Therefore, moderately oxidizing conditions inhibit only phosphatases to increase net phosphorylation, while massive exposure to ROS also inhibits kinases to suspend phosphorylation [169, 170]. For example, calcineurin (CaN), a Ca2+-activated protein phosphatase associates with RyR/IP3R via the FKBP12 proteins [171]. Ca2+-activated CaN undergoes time-dependent inactivation that can be suspended by SOD1, suggesting a ·O2 -dependent mechanism [172]. This mechanism has been implicated in the pathogenesis of amyotrophic lateral sclerosis [173]. Thus, CaN can integrate Ca2+ and ROS signals and mediate both Ca2+-activated stabilization of RyR/IP3R through FKBP dephosphorylation and ROS-induced increase in channel activation through phosphorylation [23, 174]. Furthermore, H2O2 also promotes dissociation CaN from RyR/IP3R suspending the local interaction with the channels under oxidative stress [163, 171].

Mitochondrial targets: ROS (feedback) regulation of proteins/protein complexes that contribute to mitochondrial ROS production (αKGDH, aconitase, respiratory complex I) have been discussed earlier under ‘ROS sources’. It is important to note however, that while the direct ·O2 producer complex I increases its ROS generating potency upon ROS exposure (diverting the electrons to this direction from the ETC), the production of reducing equivalents by TCA decreases. Thus, exposure to excessive ROS will lead to progressive decay of ΔΨm and matrix acidification promoting PTP activation (see below).

While considerable data argues for the physiological redox regulation of the SR/ER Ca2+ transporting proteins, much less is known about ROS-regulation of mitochondrial Ca2+ transporters. VDAC has been shown to contribute to the ·O2 -induced increase in OMM permeability, which leads to the loss of cytochrome c from the IMS [175], however no evidence has been presented so far for the VDACs physiological redox regulation as a Ca2+ pore. A recent report showing that silencing of the cytosolic glutaredoxin-1 caused oxidative modification of –SH groups of VDAC gives a hint that probably VDAC is actively protected from oxidation and could be a subject of a local redox regulation [176]. The Ca2+ uniporter and the Ca2+ exchangers have no known redox regulation.

The most extensively studied mitochondrial ROS target that conducts Ca2+ is the PTP. In highlights, the PTP opening is promoted by high [Ca2+]m, ROS, adenine nucleotide depletion, decreased ΔΨm, and elevated phosphate levels in the mitochondrial matrix [67, 177, 178]. Increased electron flow through complex I of the ETC associated with enhanced ·O2 generation promotes PTP via a quinone-mediated regulation [179, 180]. The molecular composition of the PTP is still uncertain (see [67, 177, 178]). Cyclophyilin D (CypD), a matrix peptidyl-prolyl cis-trans isomerase modulates the Ca2+ sensitivity of the PTP [181183] and it is the target of the so far most specific pharmacological PTP inhibitor, cyclosporine A (CSA). However, PT could be evoked by quinones or ROS even after genetic ablation of CypD [182]. The actual pore forming component(s) of the PTP are also elusive. The long-proposed adenine nucleotide translocator (ANT) that modulates the sensitivity of the PTP to Ca2+ and adenine nucleotides have been proven dispensible for CSA sensitive Ca2+-or ROS-dependent PTP activation based on genetic knockout studies [184]. Most recently, the mitochondrial phosphate carrier (mPiC) has been emerging as a pore candidate (perhaps in conjunction with ANT), since it also binds CypD in a CSA-sensitive manner and displays sensitivity to quinones; however this role of mPiC requires further establishment [178, 185, 186].

The PTP has at least two different redox sensing sites; one that reacts to the glutathione oxidation level (decreased matrix GSH/GSSG ratio) and another that senses the oxidation level of matrix pyridine NAD(P) [187190].

Importantly, opening of the PTP is reversible [191]; however, depending on the nature/strength of the activating stimuli it may become fixed in the open conformation ultimately disabling, osmotically destroying the mitochondrion and deliberating mitochondrial pro-apoptotic factors to the cytosol [177, 178]. Reversible (transient) openings of PTP have been suggested to be central players in the ROS-induced ROS release mechanisms underlying the regenerative ROS waves observed in cardiac muscle [129]. Transient opening of the PTP does not allow escape of significant amount of pyridine nucleotides but causes a temporary uncoupling, giving a boost to ROS production by the ETC and giving way out to the superoxide from the matrix. In a similar but rather progressive manner, enhanced NOX-derived ROS-and Ca2+-induced mitochondrial ROS production has been shown to be a central player in the neuronal degeneration associated with the deficiency of PINK1, a protein regulating mitochondrial Ca2+ extrusion and mutated in familiar Parkinson’s disease [192]. A striking ROS effect on mitochondrial fragmentation indicates that mitochondria shaping proteins or their regulators also represent a ROS target [193, 194].

4. Local Ca2+ and ROS cross-signaling between SR/ER and mitochondria

Sources as well as targets of Ca2+ and ROS are brought to close distances at focal contacts of the organelles, making Ca2+ and ROS effective local messengers. However, testing the local crosstalk between these signaling systems is technically challenging, hence to date only a few studies have provided direct evidence in this direction. In astrocytes, ROS-induced transient focal mitochondrial depolarizations (flickers) reflecting transient PTP openings have been shown to depend on Ca2+ stored in the ER. The persistence of these depolarizations at increased [Ca2+]c buffering suggested that local Ca2+ delivery from the ER to the mitochondria might have caused them and that this local Ca2+ transfer became sensitized by the ROS derived from the mitochondria. Although the observed phenomenon was based on an artificial mitochondrial ROS source (the membrane potential probe TMRE) it depicted a scenario where mitochondria-derived ROS could induce local ER Ca2+ release events that in turn, ‘hit back’ on the mitochondria to promote PTP activation [195]. The same group has also reported increased spark frequency in cardiomyocytes at regions of mitochondrial ROS generation (TMRE illumination), which was consistent with local redox sensitization of RyR2 [125]. A very recent study has given detailed account on the effect of different ROS on the spatiotemporal parameters of sparks. Using pharmacological inhibitors (respectively TMPyP and myxothiazol), it also provided evidence that basal cytosolic and mitochondrial ROS production are determinants of basal spark parameters in cardiomyocytes [196].

5. Future perspectives

Compelling evidence supports a mutual relationship between calcium and ROS signaling, which affects Ca2+ and ROS metabolism and the activation of their respective effectors. SR/ER is the main intracellular source of Ca2+ and mitochondria are important modulators of the Ca2+ dynamics. On the other hand, mitochondria and in a lesser extent, SR/ER represents a source and breakdown site of ROS. The present description of the structural and functional coupling between mitochondria and SR/ER illustrates that the SR/ER mitochondrial junction might serve as the center stage for Ca2+-ROS interactions. However, the testing of this idea will require new approaches to target and visualize the components of the local interorganellar coupling. This task is further complicated by the permanent reorganization of SR/ER and mitochondria, which likely involves several Ca2+ and ROS sensitive factors. Further consideration and studies of the local involvement of RNS will be also required. Currently available data have provided evidence that the Ca2+-ROS interplay is relevant for both physiological homeostasis and for a range of pathological conditions when either the Ca2+ or ROS regulations are disturbed. In several grave human disorders from acute ischemia-reperfusion conditions of the heart and brain to the slowly progressing neurodegenerative diseases, the local interaction of Ca2+ and ROS at the mitochondria is gaining notice. These observations give reasons for studying the derangement of the Ca2+ and ROS signaling and structure of the SR/ER-mitochondrial junction as a contributor in the development of a range of diseases.


We would like to thank Drs. Suresh K. Joseph and Pál Pacher for helpful discussions.


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1. Clapham DE. Calcium signaling. Cell. 2007;131:1047–1058. [PubMed]
2. Rizzuto R, Pozzan T. Microdomains of intracellular Ca2+: molecular determinants and functional consequences. Physiol Rev. 2006;86:369–408. [PubMed]
3. Berridge MJ. The endoplasmic reticulum: a multifunctional signaling organelle. Cell Calcium. 2002;32:235–249. [PubMed]
4. Yaffe MP. Dynamic mitochondria. Nat Cell Biol. 1999;1:E149–E150. [PubMed]
5. Franzini-Armstrong C. ER-mitochondria communication. How privileged? Physiology (Bethesda) 2007;22:261–268. [PubMed]
6. Csordas G, Renken C, Varnai P, Walter L, Weaver D, Buttle KF, Balla T, Mannella CA, Hajnoczky G. Structural and functional features and significance of the physical linkage between ER and mitochondria. J Cell Biol. 2006;174:915–921. [PMC free article] [PubMed]
7. Hayashi T, Martone ME, Yu Z, Thor A, Doi M, Holst MJ, Ellisman MH, Hoshijima M. Three-dimensional electron microscopy reveals new details of membrane systems for Ca2+ signaling in the heart. J Cell Sci. 2009;122:1005–1013. [PMC free article] [PubMed]
8. Boncompagni S, Rossi AE, Micaroni M, Beznoussenko GV, Polishchuk RS, Dirksen RT, Protasi F. Mitochondria are linked to calcium stores in striated muscle by developmentally regulated tethering structures. Mol Biol Cell. 2009;20:1058–1067. [PMC free article] [PubMed]
9. Shore GC, Tata JR. Two fractions of rough endoplasmic reticulum from rat liver. I. Recovery of rapidly sedimenting endoplasmic reticulum in association with mitochondria. J Cell Biol. 1977;72:714–725. [PMC free article] [PubMed]
10. Garcia-Perez C, Hajnoczky G, Csordas G. Physical coupling supports the local Ca2+ transfer between sarcoplasmic reticulum subdomains and the mitochondria in heart muscle. J Biol Chem. 2008;283:32771–32780. [PubMed]
11. Rusinol AE, Cui Z, Chen MH, Vance JE. A unique mitochondria-associated membrane fraction from rat liver has a high capacity for lipid synthesis and contains pre-Golgi secretory proteins including nascent lipoproteins. J Biol Chem. 1994;269:27494–27502. [PubMed]
12. Pitts KR, Yoon Y, Krueger EW, McNiven MA. The dynamin-like protein DLP1 is essential for normal distribution and morphology of the endoplasmic reticulum and mitochondria in mammalian cells. Mol Biol Cell. 1999;10:4403–4417. [PMC free article] [PubMed]
13. Varadi A, Cirulli V, Rutter GA. Mitochondrial localization as a determinant of capacitative Ca2+ entry in HeLa cells. Cell Calcium. 2004;36:499–508. [PubMed]
14. Wang HJ, Guay G, Pogan L, Sauve R, Nabi IR. Calcium regulates the association between mitochondria and a smooth subdomain of the endoplasmic reticulum. J Cell Biol. 2000;150:1489–1498. [PMC free article] [PubMed]
15. Simmen T, Aslan JE, Blagoveshchenskaya AD, Thomas L, Wan L, Xiang Y, Feliciangeli SF, Hung CH, Crump CM, Thomas G. PACS-2 controls endoplasmic reticulum-mitochondria communication and Bid-mediated apoptosis. Embo J. 2005;24:717–729. [PubMed]
16. Szabadkai G, Bianchi K, Varnai P, De Stefani D, Wieckowski MR, Cavagna D, Nagy AI, Balla T, Rizzuto R. Chaperone-mediated coupling of endoplasmic reticulum and mitochondrial Ca2+ channels. J Cell Biol. 2006;175:901–911. [PMC free article] [PubMed]
17. Hayashi T, Su TP. Sigma-1 receptor chaperones at the ER-mitochondrion interface regulate Ca(2+) signaling and cell survival. Cell. 2007;131:596–610. [PubMed]
18. de Brito OM, Scorrano L. Mitofusin 2 tethers endoplasmic reticulum to mitochondria. Nature. 2008;456:605–610. [PubMed]
19. Csordas G, Thomas AP, Hajnoczky G. Quasi-synaptic calcium signal transmission between endoplasmic reticulum and mitochondria. Embo J. 1999;18:96–108. [PubMed]
20. Spat A, Szanda G, Csordas G, Hajnoczky G. High- and low-calcium-dependent mechanisms of mitochondrial calcium signalling. Cell Calcium. 2008;44:51–63. [PMC free article] [PubMed]
21. Gorlach A, Klappa P, Kietzmann T. The endoplasmic reticulum: folding, calcium homeostasis, signaling, and redox control. Antioxid Redox Signal. 2006;8:1391–1418. [PubMed]
22. Periasamy M, Kalyanasundaram A. SERCA pump isoforms: their role in calcium transport and disease. Muscle Nerve. 2007;35:430–442. [PubMed]
23. Hidalgo C, Donoso P. Crosstalk between calcium and redox signaling: from molecular mechanisms to health implications. Antioxid Redox Signal. 2008;10:1275–1312. [PubMed]
24. Foskett JK, White C, Cheung KH, Mak DO. Inositol trisphosphate receptor Ca2+ release channels. Physiol Rev. 2007;87:593–658. [PMC free article] [PubMed]
25. Zalk R, Lehnart SE, Marks AR. Modulation of the ryanodine receptor and intracellular calcium. Annu Rev Biochem. 2007;76:367–385. [PubMed]
26. Joseph SK, Hajnoczky G. IP(3) receptors in cell survival and apoptosis: Ca(2+) release and beyond. Apoptosis. 2007;12:951–968. [PubMed]
27. Fleischer S. Personal recollections on the discovery of the ryanodine receptors of muscle. Biochem Biophys Res Commun. 2008;369:195–207. [PubMed]
28. Gyorke S, Terentyev D. Modulation of ryanodine receptor by luminal calcium and accessory proteins in health and cardiac disease. Cardiovasc Res. 2008;77:245–255. [PubMed]
29. Meissner G. Molecular regulation of cardiac ryanodine receptor ion channel. Cell Calcium. 2004;35:621–628. [PubMed]
30. Mikoshiba K. IP3 receptor/Ca2+ channel: from discovery to new signaling concepts. J Neurochem. 2007;102:1426–1446. [PubMed]
31. Hajnoczky G, Thomas AP. The inositol trisphosphate calcium channel is inactivated by inositol trisphosphate. Nature. 1994;370:474–477. [PubMed]
32. Higo T, Hattori M, Nakamura T, Natsume T, Michikawa T, Mikoshiba K. Subtype-specific and ER lumenal environment-dependent regulation of inositol 1,4,5-trisphosphate receptor type 1 by ERp44. Cell. 2005;120:85–98. [PubMed]
33. Meissner G. Regulation of mammalian ryanodine receptors. Front Biosci. 2002;7:d2072–d2080. [PubMed]
34. Choe CU, Ehrlich BE. The inositol 1,4,5-trisphosphate receptor (IP3R) and its regulators: sometimes good and sometimes bad teamwork. Sci STKE. 2006;2006:re15. [PubMed]
35. Bootman MD, Lipp P, Berridge MJ. The organisation and functions of local Ca(2+) signals. J Cell Sci. 2001;114:2213–2222. [PubMed]
36. Wilson BS, Pfeiffer JR, Smith AJ, Oliver JM, Oberdorf JA, Wojcikiewicz RJ. Calcium-dependent clustering of inositol 1,4,5-trisphosphate receptors. Mol Biol Cell. 1998;9:1465–1478. [PMC free article] [PubMed]
37. Tateishi Y, Hattori M, Nakayama T, Iwai M, Bannai H, Nakamura T, Michikawa T, Inoue T, Mikoshiba K. Cluster formation of inositol 1,4,5-trisphosphate receptor requires its transition to open state. J Biol Chem. 2005;280:6816–6822. [PubMed]
38. Marchant JS, Ramos V, Parker I. Structural and functional relationships between Ca2+ puffs and mitochondria in Xenopus oocytes. Am J Physiol Cell Physiol. 2002;282:C1374–C1386. [PubMed]
39. Tan W, Colombini M. VDAC closure increases calcium ion flux. Biochim Biophys Acta. 2007 [PMC free article] [PubMed]
40. Rapizzi E, Pinton P, Szabadkai G, Wieckowski MR, Vandecasteele G, Baird G, Tuft RA, Fogarty KE, Rizzuto R. Recombinant expression of the voltage-dependent anion channel enhances the transfer of Ca2+ microdomains to mitochondria. J Cell Biol. 2002;159:613–624. [PMC free article] [PubMed]
41. Csordas G, Madesh M, Antonsson B, Hajnoczky G. tcBid promotes Ca(2+) signal propagation to the mitochondria: control of Ca(2+) permeation through the outer mitochondrial membrane. Embo J. 2002;21:2198–2206. [PubMed]
42. Israelson A, Abu-Hamad S, Zaid H, Nahon E, Shoshan-Barmatz V. Localization of the voltage-dependent anion channel-1 Ca2+-binding sites. Cell Calcium. 2007;41:235–244. [PubMed]
43. Bathori G, Csordas G, Garcia-Perez C, Davies E, Hajnoczky G. Ca2+-dependent control of the permeability properties of the mitochondrial outer membrane and voltage-dependent anion-selective channel (VDAC) J Biol Chem. 2006;281:17347–17358. [PubMed]
44. Kroner H. Ca2+ ions, an allosteric activator of calcium uptake in rat liver mitochondria. Arch Biochem Biophys. 1986;251:525–535. [PubMed]
45. Csordas G, Hajnoczky G. Plasticity of mitochondrial calcium signaling. J Biol Chem. 2003;278:42273–42282. [PubMed]
46. Gunter TE, Pfeiffer DR. Mechanisms by which mitochondria transport calcium. Am J Physiol. 1990;258:C755–C786. [PubMed]
47. Kirichok Y, Krapivinsky G, Clapham DE. The mitochondrial calcium uniporter is a highly selective ion channel. Nature. 2004;427:360–364. [PubMed]
48. Michels G, Khan IF, Endres-Becker J, Rottlaender D, Herzig S, Ruhparwar A, Wahlers T, Hoppe UC. Regulation of the Human Cardiac Mitochondrial Ca2+ Uptake by 2 Different Voltage-Gated Ca2+ Channels. Circulation. 2009 [PubMed]
49. Brookes PS, Parker N, Buckingham JA, Vidal-Puig A, Halestrap AP, Gunter TE, Nicholls DG, Bernardi P, Lemasters JJ, Brand MD. UCPs--unlikely calcium porters. Nat Cell Biol. 2008;10:1235–1237. author reply 1237-40. [PMC free article] [PubMed]
50. Trenker M, Malli R, Fertschai I, Levak-Frank S, Graier WF. Uncoupling proteins 2 and 3 are fundamental for mitochondrial Ca2+ uniport. Nat Cell Biol. 2007;9:445–452. [PMC free article] [PubMed]
51. Sparagna GC, Gunter KK, Sheu SS, Gunter TE. Mitochondrial calcium uptake from physiological-type pulses of calcium. A description of the rapid uptake mode. J Biol Chem. 1995;270:27510–27515. [PubMed]
52. Beutner G, Sharma VK, Giovannucci DR, Yule DI, Sheu SS. Identification of a ryanodine receptor in rat heart mitochondria. J Biol Chem. 2001;276:21482–21488. [PubMed]
53. Beutner G, Sharma VK, Lin L, Ryu SY, Dirksen RT, Sheu SS. Type 1 ryanodine receptor in cardiac mitochondria: transducer of excitation-metabolism coupling. Biochim Biophys Acta. 2005;1717:1–10. [PubMed]
54. Rohacs T, Tory K, Dobos A, Spat A. Intracellular calcium release is more efficient than calcium influx in stimulating mitochondrial NAD(P)H formation in adrenal glomerulosa cells. Biochem J. 1997;328(Pt 2):525–528. [PubMed]
55. Maechler P, Kennedy ED, Wang H, Wollheim CB. Desensitization of mitochondrial Ca2+ and insulin secretion responses in the beta cell. J Biol Chem. 1998;273:20770–20778. [PubMed]
56. Moreau B, Nelson C, Parekh AB. Biphasic regulation of mitochondrial Ca2+ uptake by cytosolic Ca2+ concentration. Curr Biol. 2006;16:1672–1677. [PubMed]
57. Collins TJ, Lipp P, Berridge MJ, Bootman MD. Mitochondrial Ca(2+) uptake depends on the spatial and temporal profile of cytosolic Ca(2+) signals. J Biol Chem. 2001;276:26411–26420. [PubMed]
58. Pinton P, Leo S, Wieckowski MR, Di Benedetto G, Rizzuto R. Long-term modulation of mitochondrial Ca2+ signals by protein kinase C isozymes. J Cell Biol. 2004;165:223–232. [PMC free article] [PubMed]
59. Szanda G, Koncz P, Rajki A, Spat A. Participation of p38 MAPK and a novel-type protein kinase C in the control of mitochondrial Ca2+ uptake. Cell Calcium. 2008;43:250–259. [PubMed]
60. Moreau B, Parekh AB. Ca2+ -dependent inactivation of the mitochondrial Ca2+ uniporter involves proton flux through the ATP synthase. Curr Biol. 2008;18:855–859. [PubMed]
61. Denton RM, McCormack JG. On the role of the calcium transport cycle in heart and other mammalian mitochondria. FEBS Lett. 1980;119:1–8. [PubMed]
62. Hajnoczky G, Robb-Gaspers LD, Seitz MB, Thomas AP. Decoding of cytosolic calcium oscillations in the mitochondria. Cell. 1995;82:415–424. [PubMed]
63. Robb-Gaspers LD, Burnett P, Rutter GA, Denton RM, Rizzuto R, Thomas AP. Integrating cytosolic calcium signals into mitochondrial metabolic responses. Embo J. 1998;17:4987–5000. [PubMed]
64. Territo PR, Mootha VK, French SA, Balaban RS. Ca(2+) activation of heart mitochondrial oxidative phosphorylation: role of the F(0)/F(1)-ATPase. Am J Physiol Cell Physiol. 2000;278:C423–C435. [PubMed]
65. Jouaville LS, Pinton P, Bastianutto C, Rutter GA, Rizzuto R. Regulation of mitochondrial ATP synthesis by calcium: evidence for a long-term metabolic priming. Proc Natl Acad Sci U S A. 1999;96:13807–13812. [PubMed]
66. Bell CJ, Bright NA, Rutter GA, Griffiths EJ. ATP regulation in adult rat cardiomyocytes: time-resolved decoding of rapid mitochondrial calcium spiking imaged with targeted photoproteins. J Biol Chem. 2006;281:28058–28067. [PubMed]
67. Bernardi P, Forte M. The mitochondrial permeability transition pore. Novartis Found Symp. 2007;287:157–164. discussion 164-9. [PubMed]
68. Nicholls DG. Mitochondria and calcium signaling. Cell Calcium. 2005;38:311–317. [PubMed]
69. Chalmers S, Nicholls DG. The relationship between free and total calcium concentrations in the matrix of liver and brain mitochondria. J Biol Chem. 2003;278:19062–19070. [PubMed]
70. Bernardi P. Mitochondrial transport of cations: channels, exchangers, and permeability transition. Physiol Rev. 1999;79:1127–1155. [PubMed]
71. Szalai G, Csordas G, Hantash BM, Thomas AP, Hajnoczky G. Calcium signal transmission between ryanodine receptors and mitochondria. J Biol Chem. 2000;275:15305–15313. [PubMed]
72. Rizzuto R, Brini M, Murgia M, Pozzan T. Microdomains with high Ca2+ close to IP3-sensitive channels that are sensed by neighboring mitochondria. Science. 1993;262:744–747. [PubMed]
73. Sharma VK, Ramesh V, Franzini-Armstrong C, Sheu SS. Transport of Ca2+ from sarcoplasmic reticulum to mitochondria in rat ventricular myocytes. J Bioenerg Biomembr. 2000;32:97–104. [PubMed]
74. Pacher P, Thomas AP, Hajnoczky G. Ca2+ marks: miniature calcium signals in single mitochondria driven by ryanodine receptors. Proc Natl Acad Sci U S A. 2002;99:2380–2385. [PubMed]
75. Yi M, Weaver D, Hajnoczky G. Control of mitochondrial motility and distribution by the calcium signal: a homeostatic circuit. J Cell Biol. 2004;167:661–672. [PMC free article] [PubMed]
76. Brough D, Schell MJ, Irvine RF. Agonist-induced regulation of mitochondrial and endoplasmic reticulum motility. Biochem J. 2005;392:291–297. [PubMed]
77. Rintoul GL, Filiano AJ, Brocard JB, Kress GJ, Reynolds IJ. Glutamate decreases mitochondrial size and movement in primary forebrain neurons. J Neurosci. 2003;23:7881–7888. [PubMed]
78. Subramanian K, Meyer T. Calcium-induced restructuring of nuclear envelope and endoplasmic reticulum calcium stores. Cell. 1997;89:963–971. [PubMed]
79. Cereghetti GM, Stangherlin A, Martins de Brito O, Chang CR, Blackstone C, Bernardi P, Scorrano L. Dephosphorylation by calcineurin regulates translocation of Drp1 to mitochondria. Proc Natl Acad Sci U S A. 2008;105:15803–15808. [PubMed]
80. Han XJ, Lu YF, Li SA, Kaitsuka T, Sato Y, Tomizawa K, Nairn AC, Takei K, Matsui H, Matsushita M. CaM kinase I alpha-induced phosphorylation of Drp1 regulates mitochondrial morphology. J Cell Biol. 2008;182:573–585. [PMC free article] [PubMed]
81. Cribbs JT, Strack S. Reversible phosphorylation of Drp1 by cyclic AMP-dependent protein kinase and calcineurin regulates mitochondrial fission and cell death. EMBO Rep. 2007;8:939–944. [PubMed]
82. Hom JR, Gewandter JS, Michael L, Sheu SS, Yoon Y. Thapsigargin induces biphasic fragmentation of mitochondria through calcium-mediated mitochondrial fission and apoptosis. J Cell Physiol. 2007;212:498–508. [PubMed]
83. Malli R, Frieden M, Osibow K, Zoratti C, Mayer M, Demaurex N, Graier WF. Sustained Ca2+ transfer across mitochondria is Essential for mitochondrial Ca2+ buffering, sore-operated Ca2+ entry, and Ca2+ store refilling. J Biol Chem. 2003;278:44769–44779. [PubMed]
84. Liu T, O'Rourke B. Enhancing mitochondrial Ca2+ uptake in myocytes from failing hearts restores energy supply and demand matching. Circ Res. 2008;103:279–288. [PMC free article] [PubMed]
85. Maack C, Cortassa S, Aon MA, Ganesan AN, Liu T, O'Rourke B. Elevated cytosolic Na+ decreases mitochondrial Ca2+ uptake during excitation-contraction coupling and impairs energetic adaptation in cardiac myocytes. Circ Res. 2006;99:172–182. [PMC free article] [PubMed]
86. Droge W. Free radicals in the physiological control of cell function. Physiol Rev. 2002;82:47–95. [PubMed]
87. Pacher P, Beckman JS, Liaudet L. Nitric oxide and peroxynitrite in health and disease. Physiol Rev. 2007;87:315–424. [PMC free article] [PubMed]
88. Chance B, Sies H, Boveris A. Hydroperoxide metabolism in mammalian organs. Physiol Rev. 1979;59:527–605. [PubMed]
89. Giorgio M, Trinei M, Migliaccio E, Pelicci PG. Hydrogen peroxide: a metabolic by-product or a common mediator of ageing signals? Nat Rev Mol Cell Biol. 2007;8:722–728. [PubMed]
90. Turrens JF. Mitochondrial formation of reactive oxygen species. J Physiol. 2003;552:335–344. [PubMed]
91. Taylor ER, Hurrell F, Shannon RJ, Lin TK, Hirst J, Murphy MP. Reversible glutathionylation of complex I increases mitochondrial superoxide formation. J Biol Chem. 2003;278:19603–19610. [PubMed]
92. Beer SM, Taylor ER, Brown SE, Dahm CC, Costa NJ, Runswick MJ, Murphy MP. Glutaredoxin 2 catalyzes the reversible oxidation and glutathionylation of mitochondrial membrane thiol proteins: implications for mitochondrial redox regulation and antioxidant DEFENSE. J Biol Chem. 2004;279:47939–47951. [PubMed]
93. Turrens JF, Freeman BA, Levitt JG, Crapo JD. The effect of hyperoxia on superoxide production by lung submitochondrial particles. Arch Biochem Biophys. 1982;217:401–410. [PubMed]
94. Hansford RG, Hogue BA, Mildaziene V. Dependence of H2O2 formation by rat heart mitochondria on substrate availability and donor age. J Bioenerg Biomembr. 1997;29:89–95. [PubMed]
95. St-Pierre J, Buckingham JA, Roebuck SJ, Brand MD. Topology of superoxide production from different sites in the mitochondrial electron transport chain. J Biol Chem. 2002;277:44784–44790. [PubMed]
96. Armstrong JS, Whiteman M, Yang H, Jones DP. The redox regulation of intermediary metabolism by a superoxide-aconitase rheostat. Bioessays. 2004;26:894–900. [PubMed]
97. Bulteau AL, Ikeda-Saito M, Szweda LI. Redox-dependent modulation of aconitase activity in intact mitochondria. Biochemistry. 2003;42:14846–14855. [PubMed]
98. Bulteau AL, O'Neill HA, Kennedy MC, Ikeda-Saito M, Isaya G, Szweda LI. Frataxin acts as an iron chaperone protein to modulate mitochondrial aconitase activity. Science. 2004;305:242–245. [PubMed]
99. Tretter L, Adam-Vizi V. Inhibition of Krebs cycle enzymes by hydrogen peroxide: A key role of [alpha]-ketoglutarate dehydrogenase in limiting NADH production under oxidative stress. J Neurosci. 2000;20:8972–8979. [PubMed]
100. Starkov AA, Fiskum G, Chinopoulos C, Lorenzo BJ, Browne SE, Patel MS, Beal MF. Mitochondrial alpha-ketoglutarate dehydrogenase complex generates reactive oxygen species. J Neurosci. 2004;24:7779–7788. [PubMed]
101. Tretter L, Adam-Vizi V. Generation of reactive oxygen species in the reaction catalyzed by alpha-ketoglutarate dehydrogenase. J Neurosci. 2004;24:7771–7778. [PubMed]
102. Tretter L, Adam-Vizi V. Alpha-ketoglutarate dehydrogenase: a target and generator of oxidative stress. Philos Trans R Soc Lond B Biol Sci. 2005;360:2335–2345. [PMC free article] [PubMed]
103. Zundorf G, Kahlert S, Bunik VI, Reiser G. alpha-Ketoglutarate dehydrogenase contributes to production of reactive oxygen species in glutamate-stimulated hippocampal neurons in situ. Neuroscience. 2009;158:610–616. [PubMed]
104. Bedard K, Krause KH. The NOX family of ROS-generating NADPH oxidases: physiology and pathophysiology. Physiol Rev. 2007;87:245–313. [PubMed]
105. Geiszt M, Leto TL. The Nox family of NAD(P)H oxidases: host defense and beyond. J Biol Chem. 2004;279:51715–51718. [PubMed]
106. Zhang F, Jin S, Yi F, Xia M, Dewey WL, Li PL. Local production of O2- by NAD(P)H oxidase in the sarcoplasmic reticulum of coronary arterial myocytes: cADPR-mediated Ca2+ regulation. Cell Signal. 2008;20:637–644. [PMC free article] [PubMed]
107. Yi XY, Li VX, Zhang F, Yi F, Matson DR, Jiang MT, Li PL. Characteristics and actions of NAD(P)H oxidase on the sarcoplasmic reticulum of coronary artery smooth muscle. Am J Physiol Heart Circ Physiol. 2006;290:H1136–H1144. [PubMed]
108. Xia R, Webb JA, Gnall LL, Cutler K, Abramson JJ. Skeletal muscle sarcoplasmic reticulum contains a NADH-dependent oxidase that generates superoxide. Am J Physiol Cell Physiol. 2003;285:C215–C221. [PubMed]
109. Forstermann U. Janus-faced role of endothelial NO synthase in vascular disease: uncoupling of oxygen reduction from NO synthesis and its pharmacological reversal. Biol Chem. 2006;387:1521–1533. [PubMed]
110. Dedkova EN, Blatter LA. Characteristics and function of cardiac mitochondrial nitric oxide synthase. J Physiol. 2008 [PubMed]
111. Migliaccio E, Giorgio M, Mele S, Pelicci G, Reboldi P, Pandolfi PP, Lanfrancone L, Pelicci PG. The p66shc adaptor protein controls oxidative stress response and life span in mammals. Nature. 1999;402:309–313. [PubMed]
112. Giorgio M, Migliaccio E, Orsini F, Paolucci D, Moroni M, Contursi C, Pelliccia G, Luzi L, Minucci S, Marcaccio M, Pinton P, Rizzuto R, Bernardi P, Paolucci F, Pelicci PG. Electron transfer between cytochrome c and p66Shc generates reactive oxygen species that trigger mitochondrial apoptosis. Cell. 2005;122:221–233. [PubMed]
113. Pinton P, Rimessi A, Marchi S, Orsini F, Migliaccio E, Giorgio M, Contursi C, Minucci S, Mantovani F, Wieckowski MR, Del Sal G, Pelicci PG, Rizzuto R. Protein kinase C beta and prolyl isomerase 1 regulate mitochondrial effects of the life-span determinant p66Shc. Science. 2007;315:659–663. [PubMed]
114. Toninello A, Salvi M, Pietrangeli P, Mondovi B. Biogenic amines and apoptosis: minireview article. Amino Acids. 2004;26:339–343. [PubMed]
115. Bindoli A, Fukuto JM, Forman HJ. Thiol chemistry in peroxidase catalysis and redox signaling. Antioxid Redox Signal. 2008;10:1549–1564. [PMC free article] [PubMed]
116. Kulinsky VI, Kolesnichenko LS. Mitochondrial glutathione. Biochemistry (Mosc) 2007;72:698–701. [PubMed]
117. Schrader M, Fahimi HD. Peroxisomes and oxidative stress. Biochim Biophys Acta. 2006;1763:1755–1766. [PubMed]
118. McCall TB, Boughton-Smith NK, Palmer RM, Whittle BJ, Moncada S. Synthesis of nitric oxide from L-arginine by neutrophils. Release and interaction with superoxide anion. Biochem J. 1989;261:293–296. [PubMed]
119. Cadenas E, Davies KJ. Mitochondrial free radical generation, oxidative stress, and aging. Free Radic Biol Med. 2000;29:222–230. [PubMed]
120. Han D, Antunes F, Canali R, Rettori D, Cadenas E. Voltage-dependent anion channels control the release of the superoxide anion from mitochondria to cytosol. J Biol Chem. 2003;278:5557–5563. [PubMed]
121. Papadopoulos V, Baraldi M, Guilarte TR, Knudsen TB, Lacapere JJ, Lindemann P, Norenberg MD, Nutt D, Weizman A, Zhang MR, Gavish M. Translocator protein (18kDa): new nomenclature for the peripheral-type benzodiazepine receptor based on its structure and molecular function. Trends Pharmacol Sci. 2006;27:402–409. [PubMed]
122. Aon MA, Cortassa S, Marban E, O'Rourke B. Synchronized whole cell oscillations in mitochondrial metabolism triggered by a local release of reactive oxygen species in cardiac myocytes. J Biol Chem. 2003;278:44735–44744. [PubMed]
123. Bienert GP, Schjoerring JK, Jahn TP. Membrane transport of hydrogen peroxide. Biochim Biophys Acta. 2006;1758:994–1003. [PubMed]
124. Bienert GP, Moller AL, Kristiansen KA, Schulz A, Moller IM, Schjoerring JK, Jahn TP. Specific aquaporins facilitate the diffusion of hydrogen peroxide across membranes. J Biol Chem. 2007;282:1183–1192. [PubMed]
125. Davidson SM, Duchen MR. Calcium microdomains and oxidative stress. Cell Calcium. 2006;40:561–574. [PubMed]
126. Hopper RK, Carroll S, Aponte AM, Johnson DT, French S, Shen RF, Witzmann FA, Harris RA, Balaban RS. Mitochondrial matrix phosphoproteome: effect of extra mitochondrial calcium. Biochemistry. 2006;45:2524–2536. [PMC free article] [PubMed]
127. Wang W, Fang H, Groom L, Cheng A, Zhang W, Liu J, Wang X, Li K, Han P, Zheng M, Yin J, Mattson MP, Kao JP, Lakatta EG, Sheu SS, Ouyang K, Chen J, Dirksen RT, Cheng H. Superoxide flashes in single mitochondria. Cell. 2008;134:279–290. [PMC free article] [PubMed]
128. Romashko DN, Marban E, O'Rourke B. Subcellular metabolic transients and mitochondrial redox waves in heart cells. Proc Natl Acad Sci U S A. 1998;95:1618–1623. [PubMed]
129. Zorov DB, Filburn CR, Klotz LO, Zweier JL, Sollott SJ. Reactive oxygen species (ROS)-induced ROS release: a new phenomenon accompanying induction of the mitochondrial permeability transition in cardiac myocytes. J Exp Med. 2000;192:1001–1014. [PMC free article] [PubMed]
130. Sanchez R, Riddle M, Woo J, Momand J. Prediction of reversibly oxidized protein cysteine thiols using protein structure properties. Protein Sci. 2008;17:473–481. [PubMed]
131. Woo HA, Chae HZ, Hwang SC, Yang KS, Kang SW, Kim K, Rhee SG. Reversing the inactivation of peroxiredoxins caused by cysteine sulfinic acid formation. Science. 2003;300:653–656. [PubMed]
132. Mieyal JJ, Gallogly MM, Qanungo S, Sabens EA, Shelton MD. Molecular mechanisms and clinical implications of reversible protein S-glutathionylation. Antioxid Redox Signal. 2008;10:1941–1988. [PMC free article] [PubMed]
133. Hool LC, Corry B. Redox control of calcium channels: from mechanisms to therapeutic opportunities. Antioxid Redox Signal. 2007;9:409–435. [PubMed]
134. Shi ZZ, Osei-Frimpong J, Kala G, Kala SV, Barrios RJ, Habib GM, Lukin DJ, Danney CM, Matzuk MM, Lieberman MW. Glutathione synthesis is essential for mouse development but not for cell growth in culture. Proc Natl Acad Sci U S A. 2000;97:5101–5106. [PubMed]
135. Yant LJ, Ran Q, Rao L, Van Remmen H, Shibatani T, Belter JG, Motta L, Richardson A, Prolla TA. The selenoprotein GPX4 is essential for mouse development and protects from radiation and oxidative damage insults. Free Radic Biol Med. 2003;34:496–502. [PubMed]
136. Seiler A, Schneider M, Forster H, Roth S, Wirth EK, Culmsee C, Plesnila N, Kremmer E, Radmark O, Wurst W, Bornkamm GW, Schweizer U, Conrad M. Glutathione peroxidase 4 senses and translates oxidative stress into 12/15-lipoxygenase dependent- and AIF-mediated cell death. Cell Metab. 2008;8:237–248. [PubMed]
137. Sharov VS, Dremina ES, Galeva NA, Williams TD, Schoneich C. Quantitative mapping of oxidation-sensitive cysteine residues in SERCA in vivo and in vitro by HPLC-electrospray-tandem MS: selective protein oxidation during biological aging. Biochem J. 2006;394:605–615. [PubMed]
138. Bishop JE, Squier TC, Bigelow DJ, Inesi G. (Iodoacetamido)fluorescein labels a pair of proximal cysteines on the Ca2+-ATPase of sarcoplasmic reticulum. Biochemistry. 1988;27:5233–5240. [PubMed]
139. Adachi T, Weisbrod RM, Pimentel DR, Ying J, Sharov VS, Schoneich C, Cohen RA. S-Glutathiolation by peroxynitrite activates SERCA during arterial relaxation by nitric oxide. Nat Med. 2004;10:1200–1207. [PubMed]
140. Grover AK, Samson SE. Effect of superoxide radical on Ca2+ pumps of coronary artery. Am J Physiol. 1988;255:C297–C303. [PubMed]
141. Suzuki YJ, Edmondson JD, Ford GD. Inactivation of rabbit muscle creatine kinase by hydrogen peroxide. Free Radic Res Commun. 1992;16:131–136. [PubMed]
142. Li Y, Camacho P. Ca2+-dependent redox modulation of SERCA 2b by ERp57. J Cell Biol. 2004;164:35–46. [PMC free article] [PubMed]
143. Zissimopoulos S, Lai FA. Redox regulation of the ryanodine receptor/calcium release channel. Biochem Soc Trans. 2006;34:919–921. [PubMed]
144. Eu JP, Sun J, Xu L, Stamler JS, Meissner G. The skeletal muscle calcium release channel: coupled O2 sensor and NO signaling functions. Cell. 2000;102:499–509. [PubMed]
145. Hamilton SL, Reid MB. RyR1 modulation by oxidation and calmodulin. Antioxid Redox Signal. 2000;2:41–45. [PubMed]
146. Pessah IN, Feng W. Functional role of hyperreactive sulfhydryl moieties within the ryanodine receptor complex. Antioxid Redox Signal. 2000;2:17–25. [PubMed]
147. Xia R, Stangler T, Abramson JJ. Skeletal muscle ryanodine receptor is a redox sensor with a well defined redox potential that is sensitive to channel modulators. J Biol Chem. 2000;275:36556–36561. [PubMed]
148. Xu L, Eu JP, Meissner G, Stamler JS. Activation of the cardiac calcium release channel (ryanodine receptor) by poly-S-nitrosylation. Science. 1998;279:234–237. [PubMed]
149. Zima AV, Blatter LA. Redox regulation of cardiac calcium channels and transporters. Cardiovasc Res. 2006;71:310–321. [PubMed]
150. Huddleston AT, Tang W, Takeshima H, Hamilton SL, Klann E. Superoxide-induced potentiation in the hippocampus requires activation of ryanodine receptor type 3 and ERK. J Neurophysiol. 2008;99:1565–1571. [PubMed]
151. Joseph SK, Nakao SK, Sukumvanich S. Reactivity of free thiol groups in type-I inositol trisphosphate receptors. Biochem J. 2006;393:575–582. [PubMed]
152. Joseph SK, Ryan SV, Pierson S, Renard-Rooney D, Thomas AP. The effect of mersalyl on inositol trisphosphate receptor binding and ion channel function. J Biol Chem. 1995;270:3588–3593. [PubMed]
153. Thrower EC, Duclohier H, Lea EJ, Molle G, Dawson AP. The inositol 1,4,5-trisphosphate-gated Ca2+ channel: effect of the protein thiol reagent thimerosal on channel activity. Biochem J. 1996;318(Pt 1):61–66. [PubMed]
154. Kaplin AI, Ferris CD, Voglmaier SM, Snyder SH. Purified reconstituted inositol 1,4,5-trisphosphate receptors. Thiol reagents act directly on receptor protein. J Biol Chem. 1994;269:28972–28978. [PubMed]
155. Bultynck G, Szlufcik K, Kasri NN, Assefa Z, Callewaert G, Missiaen L, Parys JB, De Smedt H. Thimerosal stimulates Ca2+ flux through inositol 1,4,5-trisphosphate receptor type 1, but not type 3, via modulation of an isoform-specific Ca2+-dependent intramolecular interaction. Biochem J. 2004;381:87–96. [PubMed]
156. Bootman MD, Taylor CW, Berridge MJ. The thiol reagent, thimerosal, evokes Ca2+ spikes in HeLa cells by sensitizing the inositol 1,4,5-trisphosphate receptor. J Biol Chem. 1992;267:25113–25119. [PubMed]
157. Missiaen L, Taylor CW, Berridge MJ. Spontaneous calcium release from inositol trisphosphate-sensitive calcium stores. Nature. 1991;352:241–244. [PubMed]
158. Hu Q, Zheng G, Zweier JL, Deshpande S, Irani K, Ziegelstein RC. NADPH oxidase activation increases the sensitivity of intracellular Ca2+ stores to inositol 1,4,5-trisphosphate in human endothelial cells. J Biol Chem. 2000;275:15749–15757. [PubMed]
159. Camello-Almaraz MC, Pozo MJ, Murphy MP, Camello PJ. Mitochondrial production of oxidants is necessary for physiological calcium oscillations. J Cell Physiol. 2006;206:487–494. [PubMed]
160. Madesh M, Hawkins BJ, Milovanova T, Bhanumathy CD, Joseph SK, Ramachandrarao SP, Sharma K, Kurosaki T, Fisher AB. Selective role for superoxide in InsP3 receptor-mediated mitochondrial dysfunction and endothelial apoptosis. J Cell Biol. 2005;170:1079–1090. [PMC free article] [PubMed]
161. Takahashi A, Mikami M, Yang J. Hydrogen peroxide increases GABAergic mIPSC through presynaptic release of calcium from IP3 receptor-sensitive stores in spinal cord substantia gelatinosa neurons. Eur J Neurosci. 2007;25:705–716. [PMC free article] [PubMed]
162. Suzuki YJ, Ford GD. Superoxide stimulates IP3-induced Ca2+ release from vascular smooth muscle sarcoplasmic reticulum. Am J Physiol. 1992;262:H114–H116. [PubMed]
163. Zissimopoulos S, Docrat N, Lai FA. Redox sensitivity of the ryanodine receptor interaction with FK506-binding protein. J Biol Chem. 2007;282:6976–6983. [PubMed]
164. McCubrey JA, Franklin RA. Reactive oxygen intermediates and signaling through kinase pathways. Antioxid Redox Signal. 2006;8:1745–1748. [PubMed]
165. Wong W, Scott JD. AKAP signalling complexes: focal points in space and time. Nat Rev Mol Cell Biol. 2004;5:959–970. [PubMed]
166. Lygren B, Carlson CR, Santamaria K, Lissandron V, McSorley T, Litzenberg J, Lorenz D, Wiesner B, Rosenthal W, Zaccolo M, Tasken K, Klussmann E. AKAP complex regulates Ca2+ re-uptake into heart sarcoplasmic reticulum. EMBO Rep. 2007;8:1061–1067. [PubMed]
167. Tu H, Tang TS, Wang Z, Bezprozvanny I. Association of type 1 inositol 1,4,5-trisphosphate receptor with AKAP9 (Yotiao) and protein kinase A. J Biol Chem. 2004;279:19375–19382. [PubMed]
168. Marx SO, Reiken S, Hisamatsu Y, Jayaraman T, Burkhoff D, Rosemblit N, Marks AR. PKA phosphorylation dissociates FKBP12.6 from the calcium release channel (ryanodine receptor): defective regulation in failing hearts. Cell. 2000;101:365–376. [PubMed]
169. Humphries KM, Pennypacker JK, Taylor SS. Redox regulation of cAMP-dependent protein kinase signaling: kinase versus phosphatase inactivation. J Biol Chem. 2007;282:22072–22079. [PubMed]
170. Franklin RA, Rodriguez-Mora OG, Lahair MM, McCubrey JA. Activation of the calcium/calmodulin-dependent protein kinases as a consequence of oxidative stress. Antioxid Redox Signal. 2006;8:1807–1817. [PubMed]
171. Cameron AM, Steiner JP, Roskams AJ, Ali SM, Ronnett GV, Snyder SH. Calcineurin associated with the inositol 1,4,5-trisphosphate receptor-FKBP12 complex modulates Ca2+ flux. Cell. 1995;83:463–472. [PubMed]
172. Wang X, Culotta VC, Klee CB. Superoxide dismutase protects calcineurin from inactivation. Nature. 1996;383:434–437. [PubMed]
173. Ferri A, Gabbianelli R, Casciati A, Celsi F, Rotilio G, Carri MT. Oxidative inactivation of calcineurin by Cu,Zn superoxide dismutase G93A, a mutant typical of familial amyotrophic lateral sclerosis. J Neurochem. 2001;79:531–538. [PubMed]
174. Bultynck G, Vermassen E, Szlufcik K, De Smet P, Fissore RA, Callewaert G, Missiaen L, De Smedt H, Parys JB. Calcineurin and intracellular Ca2+-release channels: regulation or association? Biochem Biophys Res Commun. 2003;311:1181–1193. [PubMed]
175. Madesh M, Hajnoczky G. VDAC-dependent permeabilization of the outer mitochondrial membrane by superoxide induces rapid and massive cytochrome c release. J Cell Biol. 2001;155:1003–1015. [PMC free article] [PubMed]
176. Saeed U, Durgadoss L, Valli RK, Joshi DC, Joshi PG, Ravindranath V. Knockdown of cytosolic glutaredoxin 1 leads to loss of mitochondrial membrane potential: implication in neurodegenerative diseases. PLoS ONE. 2008;3:e2459. [PMC free article] [PubMed]
177. Bernardi P, Krauskopf A, Basso E, Petronilli V, Blachly-Dyson E, Di Lisa F, Forte MA. The mitochondrial permeability transition from in vitro artifact to disease target. Febs J. 2006;273:2077–2099. [PubMed]
178. Halestrap AP. What is the mitochondrial permeability transition pore? J Mol Cell Cardiol. 2009 [PubMed]
179. Walter L, Miyoshi H, Leverve X, Bernard P, Fontaine E. Regulation of the mitochondrial permeability transition pore by ubiquinone analogs. A progress report. Free Radic Res. 2002;36:405–412. [PubMed]
180. Fontaine E, Eriksson O, Ichas F, Bernardi P. Regulation of the permeability transition pore in skeletal muscle mitochondria. Modulation By electron flow through the respiratory chain complex i. J Biol Chem. 1998;273:12662–12668. [PubMed]
181. Baines CP, Kaiser RA, Purcell NH, Blair NS, Osinska H, Hambleton MA, Brunskill EW, Sayen MR, Gottlieb RA, Dorn GW, Robbins J, Molkentin JD. Loss of cyclophilin D reveals a critical role for mitochondrial permeability transition in cell death. Nature. 2005;434:658–662. [PubMed]
182. Basso E, Fante L, Fowlkes J, Petronilli V, Forte MA, Bernardi P. Properties of the permeability transition pore in mitochondria devoid of Cyclophilin D. J Biol Chem. 2005;280:18558–18561. [PubMed]
183. Nakagawa T, Shimizu S, Watanabe T, Yamaguchi O, Otsu K, Yamagata H, Inohara H, Kubo T, Tsujimoto Y. Cyclophilin D-dependent mitochondrial permeability transition regulates some necrotic but not apoptotic cell death. Nature. 2005;434:652–658. [PubMed]
184. Kokoszka JE, Waymire KG, Levy SE, Sligh JE, Cai J, Jones DP, MacGregor GR, Wallace DC. The ADP/ATP translocator is not essential for the mitochondrial permeability transition pore. Nature. 2004;427:461–465. [PMC free article] [PubMed]
185. Di Lisa F, Bernardi P. A CaPful of mechanisms regulating the mitochondrial permeability transition. J Mol Cell Cardiol. 2009 [PubMed]
186. Leung AW, Varanyuwatana P, Halestrap AP. The mitochondrial phosphate carrier interacts with cyclophilin D and may play a key role in the permeability transition. J Biol Chem. 2008;283:26312–26323. [PMC free article] [PubMed]
187. Costantini P, Chernyak BV, Petronilli V, Bernardi P. Modulation of the mitochondrial permeability transition pore by pyridine nucleotides and dithiol oxidation at two separate sites. J Biol Chem. 1996;271:6746–6751. [PubMed]
188. Costantini P, Colonna R, Bernardi P. Induction of the mitochondrial permeability transition by N-ethylmaleimide depends on secondary oxidation of critical thiol groups. Potentiation by copper-ortho-phenanthroline without dimerization of the adenine nucleotide translocase. Biochim Biophys Acta. 1998;1365:385–392. [PubMed]
189. Halestrap AP. Calcium, mitochondria and reperfusion injury: a pore way to die. Biochem Soc Trans. 2006;34:232–237. [PubMed]
190. Leung AW, Halestrap AP. Recent progress in elucidating the molecular mechanism of the mitochondrial permeability transition pore. Biochim Biophys Acta. 2008;1777:946–952. [PubMed]
191. Crompton M, Costi A, Hayat L. Evidence for the presence of a reversible Ca2+- dependent pore activated by oxidative stress in heart mitochondria. Biochem J. 1987;245:915–918. [PubMed]
192. Gandhi S, Wood-Kaczmar A, Yao Z, Plun-Favreau H, Deas E, Klupsch K, Downward J, Latchman DS, Tabrizi SJ, Wood NW, Duchen MR, Abramov AY. PINK1-associated Parkinson's disease is caused by neuronal vulnerability to calcium-induced cell death. Mol Cell. 2009;33:627–638. [PMC free article] [PubMed]
193. Pletjushkina OY, Lyamzaev KG, Popova EN, Nepryakhina OK, Ivanova OY, Domnina LV, Chernyak BV, Skulachev VP. Effect of oxidative stress on dynamics of mitochondrial reticulum. Biochim Biophys Acta. 2006;1757:518–524. [PubMed]
194. Jendrach M, Mai S, Pohl S, Voth M, Bereiter-Hahn J. Short- and long-term alterations of mitochondrial morphology, dynamics and mtDNA after transient oxidative stress. Mitochondrion. 2008;8:293–304. [PubMed]
195. Jacobson J, Duchen MR. Mitochondrial oxidative stress and cell death in astrocytes--requirement for stored Ca2+ and sustained opening of the permeability transition pore. J Cell Sci. 2002;115:1175–1188. [PubMed]
196. Yan Y, Liu J, Wei C, Li K, Xie W, Wang Y, Cheng H. Bidirectional regulation of Ca2+ sparks by mitochondria-derived reactive oxygen species in cardiac myocytes. Cardiovasc Res. 2008;77:432–441. [PubMed]