Calmodulin regulation (calmodulation) of voltage-gated calcium channels

Calmodulin regulation (calmodulation) of the family of voltage-gated Ca_V1-2 channels comprises a prominent prototype for ion channel regulation, remarkable for its powerful Ca²⁺ sensing capabilities, deep in elegant mechanistic lessons, and rich in biological and therapeutic implications. This field thereby resides squarely at the epicenter of Ca²⁺ signaling biology, ion channel biophysics, and therapeutic advance. This review summarizes the historical development of ideas in this field, the scope and richly patterned organization of Ca²⁺ feedback behaviors encompassed by this system, and the long-standing challenges and recent developments in discerning a molecular basis for calmodulation. We conclude by highlighting the considerable synergy between mechanism, biological insight, and promising therapeutics.The ancient Ca²⁺ sensor protein calmodulin (CaM) has emerged as a pervasive modulator of ion channels (Saimi and Kung, 2002), manifesting remarkable Ca²⁺ sensing capabilities in this context (Dunlap, 2007; Tadross et al., 2008) and furnishing essential Ca²⁺ feedback in numerous biological settings (Alseikhan et al., 2002; Xu and Wu, 2005; Mahajan et al., 2008; Adams et al., 2010; Morotti et al., 2012). Such modulation was discovered via mutations in CaM that altered the motile behavior of Paramecium, stemming from blunted activation of Ca²⁺-dependent Na⁺ current or K⁺ current (Kink et al., 1990). Since then, numerous ion channels have been found to be modulated by CaM, as extensively reviewed (Budde et al., 2002; Saimi and Kung, 2002; Wei et al., 2003; Halling et al., 2006; Tan et al., 2011; Nyegaard et al., 2012). This paper focuses on the CaM regulation (calmodulation) of voltage-gated Ca²⁺ channels, which are extensively regulated by intracellular Ca²⁺ binding to CaM (Ca²⁺/CaM; Lee et al., 1999; Peterson et al., 1999; Zühlke et al., 1999; Haeseleer et al., 2000; DeMaria et al., 2001; Budde et al., 2002; Halling et al., 2006). Such feedback regulation by Ca²⁺/CaM demonstrates versatile functional capabilities, represents a prominent prototype for ion-channel modulation, and holds far-reaching biological consequences. These attributes contribute much to the standing of voltage-gated Ca²⁺ channels as the queen of ion channels, molecules that not only sculpt membrane electrical waveforms but also serve as gatekeepers of the ubiquitous second messenger Ca²⁺ (Yue, 2004).

Classic history of discovery

The earliest signs of Ca²⁺-dependent regulation of Ca²⁺ channels came from data showing that increased Ca²⁺ could accelerate the inactivation of Ca²⁺ currents in Paramecium (Brehm and Eckert, 1978), invertebrate neurons (Tillotson, 1979), and insect muscle (Ashcroft and Stanfield, 1981). These results gave birth to the concept of Ca²⁺-dependent inactivation (CDI), the then counterintuitive notion that entities other than transmembrane voltage could modulate the rapid gating of ion channels (Eckert and Tillotson, 1981).Methodological advances permitting routine isolation and recording of Ca²⁺ currents within vertebrate preparations clearly revealed the existence of CDI of cardiac L-type Ca²⁺ currents (carried by Ca_V1.2 channels) in both multicellular preparations (Kass and Sanguinetti, 1984; Mentrard et al., 1984) and isolated myocytes (Lee et al., 1985). Classic data illustrating CDI of cardiac L-type currents is shown in Fig. 1 (A–C). Fig. 1 A (Ca) shows Ca²⁺ current from frog cardiocytes, elicited by a test voltage pulse to +80 mV (Mentrard et al., 1984). CDI becomes apparent upon Ca²⁺ entry during a voltage prepulse to +80 mV (Fig. 1 B), which causes sharply attenuated Ca²⁺ current in a subsequent test pulse (diminished area of red shading). To exclude voltage depolarization itself as the cause of inactivation, a prepulse to +120 mV (Fig. 1 C) can be demonstrated to produce far less inactivation within a subsequent test-pulse current. Because this prepulse imposed strong depolarization, but diminished Ca²⁺ entry via decreased driving force, the reemergence of test Ca²⁺ current suggests that the strong inactivation evident in Fig. 1 B could be attributed to CDI. Data of this type specifies an inactivation mechanism that depends on voltage as a U-shaped function, now considered a hallmark of CDI (Eckert and Tillotson, 1981).Open in a separate windowFigure 1.Classic U-shaped signature of CDI. (A–C) Early voltage-clamp recordings from a multicellular preparation of frog atrial trabeculae cells demonstrates CDI of Ca²⁺ currents. All listed voltages are relative to resting potential E_R (∼−80 mV) Adapted from Mentrard et al. (1984). (A) Ca²⁺ currents (shaded pink) evoked in response to an +80-mV depolarizing test pulse show robust inactivation. Fast capacitive transient was estimated by blocking these Ca²⁺ currents with 3 mM Co²⁺ solution (Co). Vertical bar corresponds to 0.5 µA of current; horizontal bar, 100 ms. (B) The Ca²⁺ current in response to the test pulse is sharply attenuated when preceded by an +80 mV prepulse. This reduction in current amplitude reflects the inactivation of Ca²⁺ channels. Format is as in A. (C) Further increase in the prepulse potential to +120 mV restores Ca²⁺ current amplitude (compare with B). Here, the diminished Ca²⁺ entry during the prepulse was insufficient to trigger CDI. The format is as in A. (D–F) Single channel recordings of L-type Ca²⁺ channels from adult rat ventricular myocytes also exhibit U-shaped dependence of Ca²⁺ channel inactivation. Adapted from Imredy and Yue (1994) with permission from Elsevier. (D) Depolarizing voltage pulse to +20 mV evokes representative elementary Ca²⁺ currents. Ensemble average currents are shown on the bottom. (E) When depolarizing pulse was preceded by +40 mV prepulse, the elementary Ca²⁺ currents are sparser and the first opening is delayed. This reduction in open probability parallels the sharp attenuation of macroscopic Ca²⁺ currents seen in B. The ensemble average is shown on the bottom. (F) Further increase in prepulse voltage to +120 mV led to a reversal of this gating pattern, once again highlighting the U-shaped dependence of CDI. Ensemble average is shown on the bottom.Still, the actual nature of effects of Ca²⁺ upon gating remained unclear. Fig. 1 (D–F) reproduces the profile of CDI at the single-molecule level (Yue et al., 1990; Imredy and Yue, 1992, 1994). Resolving data such as these is technically challenging, owing to the diminutive size of unitary Ca²⁺ currents (∼0.3 pA) and the sub-millisecond gating timescale involved. These data demonstrate the existence of U-shaped inactivation in the gating of a single cardiac L-type Ca²⁺ channel fluxing Ca²⁺, as present in an adult rat ventricular myocyte (Imredy and Yue, 1994). These results established that Ca²⁺ influx of a single channel suffices to trigger CDI, and demonstrated the clear correspondence of single-channel and multicellular behavior (Fig. 1). Thus, CDI emerged as a legitimate molecular-level modulatory process.

Advent of Ca²⁺ channel calmodulation

The Ca²⁺ sensor mediating Ca²⁺ regulation of Ca²⁺ channels has long been a matter of debate, with proposals of direct binding of Ca²⁺ to the channel complex (Standen and Stanfield, 1982; Plant et al., 1983; Eckert and Chad, 1984), and of Ca²⁺-dependent phosphorylation and/or dephosphorylation of channels (Chad and Eckert, 1986; Armstrong, D., C. Erxleben, D. Kalman, Y. Lai, A. Nairn, and P. Greengard. 1988. Abstracts of papers presented at the forty-second annual meeting of the Society of General Physiologists. Abstr. 20). The road toward identification of the actual Ca²⁺ sensor began with the cloning and expression of recombinant Ca²⁺ channels, thereby enabling structure–function studies (Snutch and Reiner, 1992). This phase of discovery was initiated by a bioinformatic observation (Babitch, 1990), pointing out that the carboxy terminus of voltage-gated Ca²⁺ channels contains a region with weak homology to an EF hand Ca²⁺-binding motif (Fig. 2 A, EF1). After the recognition of EF1, the existence of a second EF hand–like motif became apparent (Fig. 2 A, EF2). This led to chimeric channel analysis (de Leon et al., 1995), wherein segments of the carboxy termini of L-type (Ca_V1.2) and R-type (Ca_V2.3) Ca²⁺ channels were swapped. These two types of channels exhibit strong and weak CDI, respectively, under conditions of strong intracellular Ca²⁺ buffering (Liang et al., 2003), making them advantageous for chimeric analysis. Results from these experiments revealed that the proximal third of the carboxy terminus (Fig. 2 A, CI region) was important for CDI, and that EF1 of Ca_V1.2 channels was essential for the strong CDI in these channels (de Leon et al., 1995). One possible explanation was that EF1 might be the Ca²⁺ binding site for CDI postulated previously (Standen and Stanfield, 1982; Plant et al., 1983; Eckert and Chad, 1984). However, mutations within the Ca²⁺ binding loop of EF1 failed to eliminate CDI (Zhou et al., 1997; Peterson et al., 2000), focusing the search for the Ca²⁺ sensor elsewhere.Open in a separate windowFigure 2.Calmodulation: the ingredients and the flavors. (A) Channel diagram illustrates overall arrangement of structural landmarks critical for CDI. The Ca²⁺-inactivation (CI) region, spanning ∼160 residues of the channel carboxy terminus, is highly conserved across Ca_V1/2 channel families and is elemental for CDI. The proximal segment (PCI) of the CI region includes the dual vestigial EF hand (EF) segments (shaded purple and blue-green). The IQ domain, a canonical CaM binding motif critical for CDI, is located just downstream of the PCI segment. The NSCaTE element in the amino terminus of Ca_V1.2/1.3 channels is an N-lobe Ca²⁺/CaM effector site. The CBD element (gray) in the carboxy terminus of Ca_V2 channels is a CaM binding segment thought to be critical for CDI. (B) The table outlines functional bipartition of CaM in Ca_V channels and the corresponding spatial Ca²⁺ selectivities. In Ca_V1.2 and Ca_V1.3 channels, both the C-lobe and N-lobe of CaM enable fast CDI with local Ca²⁺ selectivity. Latent CDI of Ca_V1.4 channels is revealed upon deletion of an autoinhibitory domain in the distal carboxy terminus. Throughout the Ca_V2 channel family, the N-lobe of CaM evokes slow CDI with global Ca²⁺ spatial selectivity. In Ca_V2.1 channels, the C-lobe of CaM supports ultra-fast facilitation (CDF) with local Ca²⁺ selectivity. No C-lobe triggered modulation has been reported for Ca_V2.2 and Ca_V2.3 channels.Targets of CaM binding themselves often resemble CaM, a bilobed molecule with each lobe composed of two EF hands (Jarrett and Madhavan, 1991). The dual vestigial EF hands in the carboxy terminus of channels (Fig. 2 A) may be seen as resembling a lobe of CaM, which suggests that CaM itself might be the Ca²⁺ sensor for CDI. Initial studies exploring this notion, however, showed that pharmacological inhibition of CaM did not eliminate CDI of L-type Ca²⁺ channels (Imredy and Yue, 1994; Zühlke and Reuter, 1998). Nonetheless, deletions within the Ca_V1.2 channel CI region identified an IQ motif (Fig. 2 A) as a critical element in CDI (Zühlke and Reuter, 1998), and mutagenesis of this IQ domain weakens CDI (Qin et al., 1999). As IQ motifs often bind Ca²⁺-free CaM (apoCaM; Jurado et al., 1999), the possibility that CaM acts as a CDI sensor reemerged. Definitive evidence came with studies involving a Ca²⁺-insensitive mutant form of CaM (CaM₁₂₃₄), in which point mutations within all four EF hands eliminate Ca²⁺ binding (Xia et al., 1998). If “preassociation” of apoCaM with target molecules were a prerequisite for subsequent regulation by Ca²⁺ binding to this apoCaM, then CaM₁₂₃₄ could act as a dominant negative to silence regulation, as seen for small-conductance Ca²⁺-activated K channels (Xia et al., 1998). Indeed, when Ca_V1.2 channels were coexpressed with CaM₁₂₃₄ or its analogues, CDI was ablated or strongly suppressed (Peterson et al., 1999; Zühlke et al., 1999). Additionally, apoCaM was found to bind to the Ca_V1.2 CI region, with critical dependence upon the IQ segment (Erickson et al., 2001; Pitt et al., 2001; Erickson et al., 2003). In retrospect, apoCaM preassociation with Ca_V1.2 channels would sterically protect CaM from pharmacological effects (Dasgupta et al., 1989), rationalizing the prior insensitivity of CDI to such small-molecule perturbation. In all, these data firmly substantiated CaM as the sensor for Ca_V1.2 channel Ca²⁺ regulation.Initially, it was believed that only Ca_V1.2 L-type channels (Fig. 2 B) were subject to CaM-mediated Ca²⁺ regulation (Zamponi, 2003). However, this system of calmodulation was gradually recognized to pertain to most of the Ca_V1 and Ca_V2 (but not Ca_V3) branches of the Ca_V channel superfamily (Liang et al., 2003; Fig. 2 B). P-type (Ca_V2.1) channels were found to be Ca²⁺ regulated (Lee et al., 1999), and a Ca²⁺/CaM binding site downstream of the IQ element (CBD; Fig. 2 A) was argued to be important. In this initial study, no role for CaM preassociation was recognized, and no role for the IQ domain was found. Moreover, Ca²⁺ regulation of Ca_V2.1 channels manifests as a facilitation of current (Ca²⁺-dependent facilitation; CDF), followed by a slowly developing CDI. Thus, it appeared possible that the Ca²⁺ regulation of Ca_V2.1 channels might diverge mechanistically from that of Ca_V1.2 channels. However, apoCaM was later found to preassociate with Ca_V2.1 channels (Erickson et al., 2001), Ca²⁺-insensitive mutant CaM molecules were found to eliminate Ca²⁺ regulation in Ca_V2.1 channels (DeMaria et al., 2001; Lee et al., 2003), and the IQ domain was determined to be structurally essential for this regulation (DeMaria et al., 2001; Lee et al., 2003; Kim et al., 2008; Mori et al., 2008). That said, the role of the CBD segment remains contentious, with some studies still arguing for this segment’s importance in CDI (Lee et al., 2003). In contrast, deletion of the entire carboxy terminus after the IQ domain (including the CBD element) completely spared CDF and CDI of Ca_V2.1 channels (DeMaria et al., 2001; Chaudhuri et al., 2005). Overall, Ca_V2.1 and Ca_V1.2 channels appear to be Ca²⁺ regulated by a largely conserved scheme.Soon thereafter, calmodulation was established for the remaining members of the Ca_V2 class of channels (Fig. 2 B, Ca_V2.2 and Ca_V2.3), which indicates that this modulatory system pertains throughout the Ca_V1-2 channel superfamily (Liang et al., 2003). Strong CaM-mediated CDI was found in Ca_V1.3 channels (Xu and Lipscombe, 2001; Shen et al., 2006; Yang et al., 2006); a latent capacity for CaM-mediated CDI was observed in Ca_V1.4 channels (Singh et al., 2006; Wahl-Schott et al., 2006); and potential indications of CaM-mediated regulation of Ca_V1.1 channels have been described (Stroffekova, 2008, 2011).A few more nuanced points nonetheless merit attention. First, some types of calmodulation are insensitive to strong intracellular Ca²⁺ buffering (e.g., Ca_V1.2 CDI), whereas others are not (Ca_V2.3 CDI; Fig. 2 B). This contrasting sensitivity enables the use of chimeric channels under increased Ca²⁺ buffering to identify key structural determinants (de Leon et al., 1995), despite the existence of calmodulation across the channel superfamily. Second, the mechanisms underlying CDF in Ca_V1.2 channels remain mysterious, as the CDF of native channels of the heart is weak to start, and attenuated by blockade of Ca²⁺ release from neighboring ryanodine receptor channels (RYRs; Wu et al., 2001). Additionally, CDF of Ca_V1.2 channels is strongly manifest only in recombinant channels containing point mutations within the IQ domain, and only when they are expressed in frog oocytes (Zühlke et al., 2000). Curiously, CDF is not observed in recombinant Ca_V1.2 channels expressed in mammalian cell lines (Peterson et al., 1999). In contrast, CDI of Ca_V1.2 channels is strong and universally observed across experimental platforms. Third, Ca²⁺/CaM-dependent dephosphorylation by calcineurin was initially suggested as a mechanism for CDI of Ca²⁺ channels (Chad and Eckert, 1986); however, this proposition has remained controversial as previously reviewed (Budde et al., 2002). Recently, calcineurin was found to preassemble with the Ca_V1.2 channel complex through the scaffolding protein AKAP79/150 bound to the distal carboxy terminus of the channel (Oliveria et al., 2007). Furthermore, two additional maneuvers were reported to impede CDI of these channels (Oliveria et al., 2012): the overexpression of a mutant AKAP79/150 incapable of binding calcineurin, and inhibition of calcineurin activity. However, others have found that direct inhibition of calcineurin has no effect on the CDI of L-type channels within multiple neuronal preparations (Branchaw et al., 1997; Victor et al., 1997; Zeilhofer et al., 1999). Moreover, deletion of the entire distal carboxy terminus (including the AKAP79/150 binding segment) of Ca_V1.2 channels preserves strong CDI (de Leon et al., 1995; Erickson et al., 2001; Crump et al., 2013). Similarly, short variants of Ca_V1.3 channels lacking the AKAP harboring distal carboxy termini (Marshall et al., 2011) also fully support CDI (Xu and Lipscombe, 2001; Shen et al., 2006; Yang et al., 2006; Bock et al., 2011). Altogether, it may be that calcineurin and AKAP79/150 are indirect and context-dependent modulators of CDI, rather than direct effector molecules (Budde et al., 2002).

Functional bipartition of CaM and selectivity for local/global Ca²⁺ sources

The calmodulatory mechanism for Ca²⁺ channels supports remarkable forms of Ca²⁺ decoding. This feature echoes earlier discoveries of “functional bipartition” of CaM in Paramecium (Kink et al., 1990; Saimi and Kung, 2002). Here, “under-excitable” behavioral strains had mutations only in the N-lobe of CaM, whereas “over-excitable” strains had mutations only in the C-lobe. Thus, one lobe of CaM can mediate signaling to one set of functions, whereas the other lobe signals to an alternative set of operations. It was unclear, however, whether this bipartition extended to mammals. The requirement that Ca²⁺ regulation of mammalian Ca²⁺ channels requires apoCaM preassociation permitted direct exploration of this question. Coexpressing channels with “half mutant” CaM molecules (CaM₁₂ and CaM₃₄, where only C- or N-terminal lobes, respectively, bind Ca²⁺) revealed that the individual lobes of CaM can evoke distinct components of channel regulation (Fig. 2 B). In most Ca_V1 channels, the C-lobe induces a kinetically rapid phase of CDI, whereas the N-lobe yields a slower component (Peterson et al., 1999; Yang et al., 2006; Dick et al., 2008). For Ca_V1.2 channels, early studies that utilized high intracellular Ca²⁺ buffering found only minimal N-lobe–mediated CDI (Peterson et al., 1999). However, when interrogated under low intracellular Ca²⁺ buffering, slow but recognizable N-lobe–mediated CDI of Ca_V1.2 channels emerges (Dick et al., 2008; Simms et al., 2013). Most strikingly, in Ca_V2.1 channels, the lobes of CaM produce opposing polarities of regulation: the C-lobe of CaM triggers a kinetically rapid CDF, whereas the N-lobe evokes a slower CDI (DeMaria et al., 2001; Lee et al., 2003). Ca_V2.2 and Ca_V2.3 channels manifest CDI triggered mainly by the N-lobe of CaM (Liang et al., 2003). Whether this bipartition orchestrates divergent classes of behavior in higher-order animals remains an intriguing and open question (Wei et al., 2003).Beyond simply splitting the Ca²⁺ signal, however, the calmodulation of mammalian Ca²⁺ channels revealed that the lobes of CaM can selectively decode Ca²⁺ in different ways. Hints as to this capability came from the differential sensitivity of calmodulation in various channels to Ca²⁺ buffering. Processes evoked by the C-lobe of CaM are invariably insensitive to introduction of strong Ca²⁺ buffering, whereas N-lobe–mediated processes in Ca_V2 channels can be eliminated by the same maneuver. Strong buffering would only spare Ca²⁺ signals near the cytoplasmic mouth of channels where strong point-source Ca²⁺ influx would overwhelm buffer capacity (Neher, 1986; Stern, 1992). This argues that local Ca²⁺ influx through individual channels triggers C-lobe signaling; in other words, the C-lobe of CaM exhibits a “local Ca²⁺ selectivity.” In contrast, a spatially global elevation of Ca²⁺ (present in the absence of strong Ca²⁺ buffering) is required for N-lobe signaling of Ca_V2 channels (DeMaria et al., 2001; Lee et al., 2003; Liang et al., 2003); thus, the N-lobe in this context exhibits a “global selectivity.” The existence of global selectivity is notable, given that local Ca²⁺ influx yields far larger Ca²⁺ increases near a channel than does globally sourced Ca²⁺ (Tay et al., 2012). One exception to this pattern is the local Ca²⁺ selectivity of the N-lobe component of CDI in Ca_V1.2/1.3 channels (see two paragraphs below). Corroboration of spatial Ca²⁺ selectivity is provided by the ability of single Ca_V1.2 channels to undergo CDI (Fig. 1, D–F). By definition, only a local Ca²⁺ source is present in the single-channel configuration; thus, the presence of CDI in this context indicates local Ca²⁺ selectivity. Additionally, single-channel records of Ca_V2.1 channels exhibit CDF (driven by C-lobe), but not CDI (triggered by N-lobe; Chaudhuri et al., 2007), which is consistent with the proposed differential selectivities for this channel. Fig. 2 B summarizes the arrangement of spatial Ca²⁺ selectivities according to the lobes of CaM.The mechanisms underlying these contrasting spatial Ca²⁺ selectivities can be interpreted as emergent behaviors of a system in which a lobe of apoCaM must transiently detach from a preassociation site before binding Ca²⁺, and where a Ca²⁺-bound lobe of CaM must associate with a channel effector site to mediate regulation (Tadross et al., 2008). When the slow Ca²⁺-unbinding kinetics of the C-lobe of CaM are imposed on this scenario, local Ca²⁺ sensitivity invariably results. In contrast, when the rapid Ca²⁺-unbinding kinetics of the N-lobe are interfaced with this architecture, global Ca²⁺ selectivity arises if the channel preferentially binds apoCaM versus Ca²⁺/CaM, as in the case of Ca_V2 channels (Fig. 2 B). When the channel preferentially interacts with Ca²⁺/CaM, local Ca²⁺ selectivity emerges, as in the context of Ca_V1 channels (Fig. 2 B). The relative roles of the affinities and kinetics of Ca²⁺ binding to the two lobes of CaM in mediating local and global Ca²⁺ selectivities have been considered at length elsewhere (Tadross et al., 2008).As an explicit example of how spatial Ca²⁺ selectivity of N-lobe regulation is specified, we consider the effects of an N-terminal spatial Ca²⁺-transforming element (NSCaTE) present only on the amino terminus of Ca_V1.2/1.3 channels (Fig. 2 A). NSCaTE is a Ca²⁺/CaM binding site whose presence enhances the aggregate channel affinity for Ca²⁺/CaM over apoCaM (Ivanina et al., 2000; Dick et al., 2008), endowing the N-lobe component of CDI of these channels with a largely local Ca²⁺ selectivity of N-lobe CDI in these channels. Elimination of the NSCaTE site in these channels, which tilts channels toward a preference for apoCaM, then switches their N-lobe CDI to a global profile. Conversely, donation of NSCaTE to Ca_V2 channels, which causes channels to favor Ca²⁺/CaM binding, then endows their N-lobe CDI with local Ca²⁺ selectivity (Dick et al., 2008). In this manner, the presence of NSCaTE can tune the spatial Ca²⁺ selectivity of N-lobe–mediated channel regulation. This overall arrangement of CaM interactions with a target molecule could endow numerous other regulatory systems with spatial Ca²⁺ selectivity.

Molecular basis of calmodulation

Elucidating the arrangement of apoCaM and Ca²⁺/CaM on Ca²⁺ channels is fundamental for the field, given the biological influence of calmodulation, and the importance of this system as an ion-channel regulatory prototype (Dunlap, 2007). This critical task, however, remains an ongoing challenge, given the >2,000 amino acids comprising the main pore-forming α₁ subunit alone, the ability of multiple peptide segments of the channel to bind CaM in vitro, and the formidable nature of obtaining atomic structures for these channels.The stoichiometry of CaM interaction with channels has been debated. Crystal structures of Ca²⁺/CaM complexed with portions of the carboxy tail CI region of Ca_V1.2 channels (Fallon et al., 2009; Kim et al., 2010) suggest that multiple CaM molecules may interact to produce the full spectrum of Ca²⁺ regulatory functions. Moreover, CaM can interact with multiple peptide segments of the channel (Ivanina et al., 2000; Tang et al., 2003; Zhou et al., 2005; Dick et al., 2008; Ben Johny et al., 2013). In contrast, covalent fusion of single CaM molecules to Ca_V1.2 channels suggests that only one CaM per channel suffices to elicit CDI (Mori et al., 2004). Moreover, live-cell FRET studies of the interactions between CaM and holochannels (Ca_V1.2) also point to a 1:1 CaM/channel ratio (Ben Johny et al., 2012). In the holochannel, the various CaM binding segments may be arranged in close proximity so as to allow the two lobes of CaM to preferentially interact with the distinct channel segments during Ca²⁺ regulation (Dick et al., 2008; Ben Johny et al., 2013). Recent biochemical studies of the NSCaTE element show that this segment preferentially interacts with a single lobe of CaM (N-lobe) at a time (Liu and Vogel, 2012). Furthermore, the NSCaTE element can interact also with Ca²⁺/CaM prebound to an IQ domain peptide (Taiakina et al., 2013). Thus, we favor the interpretation that only one CaM interacts within holochannels, although peptide fragments of Ca²⁺ channels may bind to multiple CaM molecules.The IQ domain (Fig. 3 A, blue circle) is critical for calmodulation. Mutations within this domain markedly modulate the Ca²⁺ regulation of Ca_V1.2 channels (Zühlke and Reuter, 1998; Qin et al., 1999; Zühlke et al., 1999, 2000; Erickson et al., 2003), Ca_V1.3 channels (Yang et al., 2006; Bazzazi et al., 2013; Ben Johny et al., 2013), Ca_V2.1 channels (DeMaria et al., 2001; Lee et al., 2003; Kim et al., 2008; Mori et al., 2008), and Ca_V2.2 and Ca_V2.3 channels (Liang et al., 2003). Furthermore, apoCaM preassociation with Ca_V1.2/1.3 channels requires the IQ domain (Erickson et al., 2001; Pitt et al., 2001; Erickson et al., 2003; Bazzazi et al., 2013; Ben Johny et al., 2013). Finally, Ca²⁺/CaM binds well to IQ-domain peptides of many Ca_V channels (Peterson et al., 1999; Zühlke et al., 1999; DeMaria et al., 2001; Pitt et al., 2001; Liang et al., 2003; Bazzazi et al., 2013; Ben Johny et al., 2013). All these results gave rise to the IQ-centric hypothesis shown in Fig. 3 A. Here, the IQ domain is important for both apoCaM preassociation (left) and Ca²⁺/CaM binding (effector configuration shown on the right). This IQ-centric paradigm has motivated efforts to resolve crystal structures of Ca²⁺/CaM complexed with IQ-domain peptides of various Ca_V1-2 channels (Fallon et al., 2005; Van Petegem et al., 2005; Kim et al., 2008; Mori et al., 2008), as shown in Fig. 3 (B and C). Throughout, the IQ peptide segment appears as an α-helical entity with N and C termini as labeled.Open in a separate windowFigure 3.Toward an atomic-level understanding of CDI. (A) IQ-centric view of calmodulation of Ca_V channels. In this mechanistic scheme, ApoCaM is preassociated with the IQ domain (blue). Upon Ca²⁺ binding, CaM rebinds the same IQ domain with a higher affinity, and subtle conformational rearrangements are presumed to trigger CaM-mediated channel regulation. (B, left) Crystal structure of Ca_V1.2 IQ domain peptide in complex with Ca²⁺/CaM (Protein Data Bank accession no. 2BE6). The IQ domain is colored blue. CaM is show in cyan (N-lobe, pale cyan; C-lobe, cyan). Ca²⁺ ions are depicted as yellow spheres. The key isoleucine residue (red) serves as a hydrophobic anchor for CaM. Ca²⁺/CaM adopts a parallel arrangement with the IQ domain in which the N-lobe binds closer to the amino terminus of the IQ domain and the C-lobe binds further downstream. (B, right) Crystal structure of Ca²⁺/CaM bound to mutant Ca_V1.2 IQ domain with alanines substituted for the key isoleucine-glutamine residues (accession no. 2F3Z). This double mutation abolishes CDI. Structurally, however, Ca²⁺/CaM hugs the mutant IQ domain in a similar conformation as to its interaction with the wild-type (left). (C) Crystal structure of Ca²⁺/CaM bound to Ca_V2.1 IQ domain (left, accession no. 3BXK; right, accession no. 3DVM). The IQ domain is shaded green, CaM in cyan. Ca²⁺/CaM was reported to bind to the Ca_V2.1 IQ domain in both parallel (left) and antiparallel (right) arrangements. The antiparallel arrangement in which the C-lobe of CaM binds upstream of IQ domain has led to speculation that CDF may result from this inverted polarity of Ca²⁺/CaM association. (D) Simplified configurations of the CaM/channel complex relevant for calmodulation (E, A, I_C, I_N, and I_CN). In configuration E, channels lack preassociated CaM and therefore cannot undergo CDI. Configuration A corresponds to channels bound to apoCaM, and thus capable of undergoing robust CDI. Ca²⁺ binding to the C-lobe and N-lobe of CaM leads to the inactivated configurations I_C and I_N, respectively. Ca²⁺ binding to both lobes of CaM then yields configuration I_CN, with both C and N lobes of CaM engaged toward CDI. This final transition may exhibit positive cooperativity as specified by the constant λ >> 1. Under endogenous levels of CaM, channels may reside in any of the five configurations. Strong coexpression of wild-type or mutant CaM restricts accessibility to various states. Reproduced with permission from Ben Johny et al., 2013.However, this viewpoint remains problematic in three regards. (1) The atomic structures of Ca²⁺/CaM bound to wild-type and mutant IQ peptides of Ca_V1.2 (Fig. 3 B) show that a central isoleucine (side chains explicitly shown) in the IQ element is deeply buried within the C-lobe of Ca²⁺/CaM (Van Petegem et al., 2005), and that alanine substitution at this site hardly changes structure (Fallon et al., 2005). Additionally, Ca²⁺/CaM dissociation constants for corresponding wild-type and mutant IQ peptides are nearly the same (Zühlke et al., 2000; Bazzazi et al., 2013). Why then would alanine substitution at this well-ensconced site influence the rest of the channel to blunt regulation (Fallon et al., 2005)? (2) For Ca_V1.2/1.3 channels, the N-lobe of Ca²⁺/CaM effector site appears to be an NSCaTE element (Fig. 3 A, oval) of the channel amino terminus (Dick et al., 2008; Tadross et al., 2008; Liu and Vogel, 2012), separate from the IQ element. (3) Crystal structures of Ca²⁺/CaM in complex with the IQ peptide of Ca_V2.1 channels show that CaM can adopt both a parallel (Mori et al., 2008) and an antiparallel configuration (Kim et al., 2008; Fig. 3 C, left and right, respectively). The apparent inversion in configuration of CaM binding to IQ domain between Ca_V1.2 and Ca_V2.1 has been proposed as a mechanism for the opposing polarity of Ca²⁺ regulation observed in the two channels (Kim et al., 2008; Minor and Findeisen, 2010). However, detailed structural analysis along with functional systematic alanine scanning mutagenesis and chimeric channel analysis argues that the C-lobe effector site resides at a site beyond the IQ module (Mori et al., 2008).A major concern with older IQ domain analyses is that the regulatory system was not considered conceptually as a whole, as shown in Fig. 3 D (drawn with specific reference to Ca_V1.3 channels; Ben Johny et al., 2013). Configuration E portrays channels lacking apoCaM. Such channels can open normally, but do not manifest CDI because Ca²⁺/CaM from bulk solution cannot efficiently access a channel in configuration E to induce CDI (Mori et al., 2004; Yang, P.S., M.X. Mori, E.A. Antony, M.R. Tadross, and D.T. Yue. 2007. Biophysical Journal abstracts issue. 1669-Plat; Liu et al., 2010; Findeisen et al., 2011). ApoCaM binding with configuration E gives rise to configuration A, where opening can also occur normally, but CDI can now take place. For CDI, Ca²⁺ binding to both lobes of CaM yields configuration I_CN, which underlies a fully inactivated channel with strongly reduced opening. For intermediate configurations, Ca²⁺ binding only to the C-lobe brings about configuration I_C, equivalent to a C-lobe inactivated channel; Ca²⁺ binding only to the N-lobe yields the N-lobe–inactivated arrangement (I_N). Of particular importance, ensuing entry into I_CN likely involves positively cooperative interactions specified by cooperativity factor λ >> 1.This scheme emphasizes several challenges for older analyses of the IQ domain, where CDI was mostly assessed with only endogenous CaM present. Results thus obtained would be ambiguous, because IQ-domain mutations could alter calmodulation via changes at multiple steps depicted in Fig. 3 D, whereas other interpretations largely attributed the effects to altered Ca²⁺/CaM binding with an IQ effector site. In contrast, IQ mutations could well weaken apoCaM preassociation and reduce CDI by favoring configuration E. Moreover, functional deficits caused by mutations that do attenuate interaction with one lobe of Ca²⁺/CaM may be masked by positively cooperative steps (λ in Fig. 3 D).To minimize these issues, CDI of Ca_V1.3 channels was recently characterized during strong coexpression with various mutant CaM molecules (Bazzazi et al., 2013; Ben Johny et al., 2013) as shown in Fig. 4 (A–C, top). For orientation, CDI of channels expressed with only endogenous CaM present is shown in the upper portion of Fig. 4 A. Strong CDI produces a rapid decay of whole-cell Ca²⁺ current (red trace), compared with the negligible decline of Ba²⁺ current (black trace). Because Ba²⁺ binds CaM poorly (Chao et al., 1984), the decline of Ca²⁺ versus Ba²⁺ current after 300 ms of depolarization quantifies CDI (CDI parameter plotted below in bar graphs). One can isolate the diamond-shaped subsystem lacking configuration E (Fig. 4 B, top) by leveraging mass action with strong coexpression of wild-type CaM (CaM_WT). Further simplification of CDI was obtained by strongly coexpressing channels with CaM₁₂ (Fig. 4 C, top), which empties configuration E by mass action, and forbids configurations I_N and I_CN. Thus, the isolated C-lobe component of CDI can be studied, with the signature rapid time course of current decay shown near the top of Fig. 4 C. Critically, this layout avoids interplay with cooperative λ steps in Fig. 3 D. Likewise, strongly coexpressing CaM₃₄ focuses on the slower N-lobe form of CDI, with attendant simplifications.Open in a separate windowFigure 4.Residue-level roadmap for calmodulation of Ca_V1.3 channels. Systematic alanine scanning mutagenesis of the entire CI domain of Ca_V1.3 channels. (A) CDI of wild-type and mutant Ca_V1.3 channels recombinantly expressed in HEK293 cells was measured with only endogenous CaM present. CDI observed here reflects properties of the entire system (stick figure) diagrammed in Fig. 3 D. Exemplar whole-cell current shows robust CDI for wild-type (WT) Ca_V1.3, reflecting their high affinity for apoCaM. Here and throughout, the vertical bar pertains to 0.2 nA of Ca²⁺ current (black); the Ba²⁺ current (gray) has been scaled ∼3-fold downward to aid comparison of decay kinetics. Horizontal bar, 100 ms. CDI is measured as the fractional reduction of Ca²⁺ current in comparison to Ba²⁺ at 300 ms (Ben Johny et al., 2013). Bottom, bar graph summary of CDI for alanine substitutions for indicated residues. CDI for WT channels, blue-green vertical line. Alanine substitutions in both EF hand regions and IQ domain resulted in diminished CDI. (B) Overexpression of CaM_WT isolates the behavior of the diamond-shaped subsystem. Such overexpression of CaM_WT should rescue CDI for mutations that weakened apoCaM preassociation. For WT Ca_V1.3, exemplar current shows that CDI is unaltered, as expected for a channel with high affinity for apoCaM. Bottom, bar graph summary of CDI for various mutant channels with CaM_WT overexpressed. Format is as in A. CDI is rescued for several mutations in the EF hand region (EF2) and the IQ domain, reflecting the preassociation of N-lobe and C-lobe of apoCaM. (C) Overexpression of CaM₁₂ isolates C-lobe CDI. Exemplar currents for WT channels show the fast C-lobe form of CDI. Bottom, bar graph summary shows the footprint of C-lobe CDI. Mutants in both the first EF hand and the IQ domain disrupted the C-lobe CDI (see the LGF loci in the EF hand regions and TFL and IQD loci in the IQ domain). The format is as in A. (D) Overexpression of CaM₃₄ isolates N-lobe CDI. N-lobe CDI was largely preserved by mutations in the CI region. Mutations that weakened N-lobe apoCaM preassociation exhibited enhanced N-lobe CDI (see EF2 segment). Reassuringly, alanine scanning mutagenesis of NSCaTE element resulted in specific disruption of N-lobe CDI of Ca_V1.3 channels (Tadross et al., 2008). Adapted with permission from Ben Johny et al. (2013) and Bazzazi et al. (2013).Accordingly, a new framework of CaM/channel configurations that underlie calmodulation could be deduced, as diagrammed in Fig. 5 (Ben Johny et al., 2013). For convenience, Fig. 4 compiles results from a systematic alanine scan covering the entire CI domain of the carboxyl tail of Ca_V1.3 channels (Bazzazi et al., 2013; Ben Johny et al., 2013), where substitution positions are denoted to the left of the bar graph in Fig. 4 A. The N-terminal end of the CI segment corresponds to the top of the graph, purple and blue-green regions indicate EF1 and EF2, and the lavender zone denotes the IQ domain. Electrophysiological characterization for each of the substitutions was performed for all the various subsystems (Fig. 4, A–D, top), and bar graphs plot the strength of CDI for the corresponding subsystems in each of the panels. The blue-green vertical lines reference the profile for wild-type channels. The results thus obtained were combined with CaM binding assays (not depicted) to identify meaningful structure–function correlations.Open in a separate windowFigure 5.Next-generation blueprint for CaM/CaBP regulation of Ca_V channels. (A) De novo molecular model of Ca_V1.3 CI region docked to apoCaM. The N-lobe of apoCaM is thought to preassociate with the PCI region (green), whereas the C-lobe binds to the IQ domain (blue). The NSCaTE segment (tan) is unoccupied. This model corresponds to configuration A in Fig. 3 D, where the channels are charged with an apoCaM and ready to undergo CDI. (B) Proposed model for the Ca²⁺ inactivated state of the channel. The N-lobe of Ca²⁺/CaM is shown binding to the NSCaTE segment based on a recent NMR structure (Protein Data Bank accession no. 2LQC). This configuration is believed to result in N-lobe CDI. The de novo molecular model shows C-lobe of Ca²⁺/CaM, the EF hand segment, and the IQ domain forming a tripartite complex resulting in C-lobe CDI. Overall, this model corresponds to the inactive configuration I_CN in Fig. 3 D. (C) Proposed allosteric mechanism of CaBP action. CaBP and apoCaM may dually bind the Ca_V1 channel. However, CaBP may allosterically inhibit CDI of Ca_V1 channels by interacting with the NT, III-IV loop, or PCI segment, thereby preventing Ca²⁺/CaM from reaching its effector configuration. (A–C) Adapted with permission from Yang et al. (2014).Under the regimen in Fig. 4 A, where all configurations are potentially accessible, diminished CDI by alanine substitutions occurred not only in the IQ domain, but upstream in the EF hand regions (between LGF and TLF residues). The latter effects strongly suggested that something outside the IQ domain was essential. Some of these deficits in CDI could be attributed to weakening of apoCaM preassociation with channels, because overexpressing wild-type CaM to depopulate configuration E (in Fig. 4 B) substantially recovered many of these CDI deficits. Binding assays of apoCaM with various CI segments confirmed this interpretation (Bazzazi et al., 2013; Ben Johny et al., 2013).Turning to potential deficits relating to diminished Ca²⁺/CaM action through channel effector sites, Fig. 4 (C and D) separately interrogates the C-lobe and N-lobe components of CDI. Importantly, isolating the C- and N-lobe components minimizes masking of mutational effects by positive cooperativity, allowing CDI deficits to be more sensitively observed in this regimen. That said, Fig. 4 C displays C-lobe CDI deficits in the EF hand region (strongest for LGF substitution), as well as in the IQ domain (with the strongest effect upon substitution at the central isoleucine). These outcomes support a hypothesis where the Ca²⁺/CaM effector configuration for the C-lobe component of CDI involves both of these functional hotspots, as developed two paragraphs below. For the N-lobe component of CDI (Fig. 4 D), no appreciable deficits were observed, as would be expected if interaction with the channel NSCaTE element (in the channel amino terminus) serves as the effector element.Fig. 5 A displays a proposed configuration A for apoCaM interaction with the channel. This includes a homology model of the apoCaM C-lobe complexed with the IQ domain (blue), based on analogous Na_V structures (Chagot and Chazin, 2011; Feldkamp et al., 2011). The portrayal of the apoCaM N-lobe incorporates ab initio structural prediction of the CI domain, with two vestigial EF hands (EF), and a protruding helix (preIQ subelement). The EF hand module (EF) resembles the structure of a homologous Na_V segment (Chagot et al., 2009; Wang et al., 2012), and the preIQ helix resembles a helical segment observed in crystal structures of analogous Ca_V1.2 peptides (Fallon et al., 2009; Kim et al., 2010). The atomic structure of the apoCaM N-lobe (1CFD) was interfaced with shape-complementarity docking algorithms. Regarding the proposed interaction interface of the N-lobe with the channel EF domain, we note that alanine substitution therein produced a curious enhancement of N-lobe CDI seen with certain alanine substitutions, especially in the EF region (Fig. 4 D). This feature is interesting because weakening of channel interaction with the N-lobe of apoCaM would be predicted to strengthen CDI produced by the N-lobe of Ca²⁺/CaM (Tadross et al., 2008).Fig. 5 B displays a proposed arrangement for the Ca²⁺/CaM-bound configuration I_CN. The N-lobe complex with NSCaTE is an NMR structure (Liu and Vogel, 2012). An alternative ab initio model of the PCI is computationally docked with the C-lobe of Ca²⁺/CaM (Protein Data Bank accession no. 3BXL) and IQ module, which together form a ternary complex. This ternary arrangement is consistent with the importance of both IQ and EF domains for C-lobe CDI (Fig. 4 C). Intriguingly the C-lobe configuration resembles a rather canonical CaM–peptide complex, where the channel EF module contributes a surrogate lobe of CaM. Importantly, assays of Ca²⁺/CaM binding to the IQ segment alone would correlate poorly with functional effects relating to the ternary complex, as experimental studies observe (Bazzazi et al., 2013; Ben Johny et al., 2013). Overall, the proposed framework in Fig. 5 (A and B) may aid future structural biology and structure–function work. In addition, this scheme of CaM exchange within its target molecule may generalize beyond the Ca²⁺ channel family.

Biological consequences and prospects for new disease therapies

The biological consequences of Ca²⁺ regulation by CaM promise to be wide ranging and immense. In the heart, elimination of Ca_V1.2 CDI by means of dominant-negative CaM elicits marked prolongation of ventricular action potential duration (APD), implicating CDI as a dominant control factor in specifying APD (Alseikhan et al., 2002; Mahajan et al., 2008). As APD is one of the main determinants of electrical stability and arrhythmias in the heart, pharmacological manipulation of such regulation looms as a future antiarrhythmic strategy (Mahajan et al., 2008; Anderson and Mohler, 2009). Most recently, genome-wide linkage analysis in humans has uncovered heritable and de novo CaM mutations as the probable cause of several cases of catecholaminergic polymorphic ventricular tachycardia (CPVT), with altered CaM-ryanodine receptor function implicated as a major contributing factor (Nyegaard et al., 2012). Whole-exome sequencing has revealed an association between three de novo missense CaM mutations and severe long-QT (LQT) syndrome with recurrent cardiac arrest (Crotti et al., 2013), quite plausibly acting via disruption of Ca_V1.2/1.3 channel CDI (Limpitikul et al., 2014).In brain, state-of-the-art analysis of genome-wide single-nucleotide polymorphisms (SNPs) has identified Ca_V channels as a major risk factor for several psychiatric disorders (Cross-Disorder Group of the Psychiatric Genomics Consortium, 2013). More specifically to calmodulation, Ca_V1.3 channels constitute a prominent Ca²⁺ entry portal into pacemaking and oscillatory neurons (Bean, 2007), owing to the more negative voltages required to open these ion channels. These channels are subject to extensive alternative splicing (Hui et al., 1991; Xu and Lipscombe, 2001; Shen et al., 2006; Bock et al., 2011; Tan et al., 2011) and RNA editing (Huang et al., 2012) in the carboxy tail, in ways that strongly modulate the strength of CDI (Shen et al., 2006; Liu et al., 2010; Huang et al., 2012). This fine tuning of CDI appears to be important for circadian rhythms (Huang et al., 2012). From the specific perspective of disease, Ca_V1.3 channels contribute a substantial portion of Ca²⁺ entry into substantia nigral neurons (Bean, 2007; Chan et al., 2007; Puopolo et al., 2007; Guzman et al., 2009), which exhibit high-frequency pacemaking (Chan et al., 2007) that drives dopamine release important for movement control. Notably, loss of these neurons is intimately related to Parkinson’s disease, and Ca²⁺ disturbances and overload are critical to this neurodegeneration (Bezprozvanny, 2009; Surmeier and Sulzer, 2013). Accordingly, an attractive therapeutic possibility for Parkinson’s disease involves discovery of small molecules that selectively down-regulate the opening of Ca_V1.3 versus other closely related Ca²⁺ channels (Kang et al., 2012). Thus, understanding the mechanisms underlying the modulation of Ca_V1.3 CDI is crucial, particularly to furnish specific molecular interfaces as targets of rational screens for small-molecule modulators.The fundamental mechanism of action of a natural modulator of Ca_V1.3 CDI may be especially relevant. In particular, recent discoveries identify Ca²⁺-binding proteins (CaBPs), a family of CaM-like brain molecules (Haeseleer et al., 2000) that may also bind Ca_V channels and other targets (Yang et al., 2002; Kasri et al., 2004). Like CaM, CaBPs are bilobed, with each lobe containing two EF hand Ca²⁺ binding motifs (Haeseleer et al., 2000), and a recent crystal structure shows overall similarity to CaM (Findeisen and Minor, 2010). However, whereas all four EF hands bind Ca²⁺ in CaM, one lobe is nonfunctional in CaBPs. Coexpressing CaBPs with Ca_V1 channels eliminates their CDI, thereby potentially influencing diverse biological processes (Zhou et al., 2004; Yang et al., 2006). Some have argued that CaBPs may simply compete for apoCaM on channels (Lee et al., 2002; Findeisen et al., 2013; Oz et al., 2013). More recent data argue for a different mechanism (Fig. 5 C), in which CaBP and apoCaM can both preassociate with the channel (Yang et al., 2014). In particular, by binding to channel regions that overlap Ca²⁺/CaM effector loci, CaBPs may preemptively retard the ability of Ca²⁺/CaM to reach its effector configuration (Fig. 5 B), allowing low concentrations of CaBP molecules in the CNS to still exert functional effects in the presence of much higher CaM levels (Yang et al., 2014). If this scheme is correct, the basic mechanisms of CaBP action may have implications for drug discovery, in that small molecules that target channel interaction surfaces for CaBP and/or Ca²⁺/CaM may exert potent modulatory actions.In conclusion, this overview of the calmodulation of voltage-gated Ca²⁺ channels highlights an impressive synergy among elegant molecular regulatory mechanisms, vital biological functions, and pathogenesis and potential therapy. As such, this field promises considerable mystery, enrichment, and enlightenment in the years ahead.     

Focus Issue on Calcium Signaling: Analyses of a Gravistimulation-Specific Ca2+ Signature in Arabidopsis using Parabolic Flights

Gravity is a critical environmental factor affecting the morphology and functions of organisms on the Earth. Plants sense changes in the gravity vector (gravistimulation) and regulate their growth direction accordingly. In Arabidopsis (Arabidopsis thaliana) seedlings, gravistimulation, achieved by rotating the specimens under the ambient 1g of the Earth, is known to induce a biphasic (transient and sustained) increase in cytoplasmic calcium concentration ([Ca2+]_c). However, the [Ca2+]_c increase genuinely caused by gravistimulation has not been identified because gravistimulation is generally accompanied by rotation of specimens on the ground (1g), adding an additional mechanical signal to the treatment. Here, we demonstrate a gravistimulation-specific Ca²⁺ response in Arabidopsis seedlings by separating rotation from gravistimulation by using the microgravity (less than 10⁻⁴g) conditions provided by parabolic flights. Gravistimulation without rotating the specimen caused a sustained [Ca2+]_c increase, which corresponds closely to the second sustained [Ca2+]_c increase observed in ground experiments. The [Ca2+]_c increases were analyzed under a variety of gravity intensities (e.g. 0.5g, 1.5g, or 2g) combined with rapid switching between hypergravity and microgravity, demonstrating that Arabidopsis seedlings possess a very rapid gravity-sensing mechanism linearly transducing a wide range of gravitational changes (0.5g–2g) into Ca²⁺ signals on a subsecond time scale.Calcium ion (Ca²⁺) functions as an intracellular second messenger in many signaling pathways in plants (White and Broadley, 2003; Hetherington and Brownlee, 2004; McAinsh and Pittman, 2009; Spalding and Harper, 2011). Endogenous and exogenous signals are spatiotemporally encoded by changing the free cytoplasmic concentration of Ca²⁺ ([Ca2+]_c), which in turn triggers [Ca2+]_c-dependent downstream signaling (Sanders et al., 2002; Dodd et al., 2010). A variety of [Ca2+]_c increases induced by diverse environmental and developmental stimuli are reported, such as phytohormones (Allen et al., 2000), temperature (Plieth et al., 1999; Dodd et al., 2006), and touch (Knight et al., 1991; Monshausen et al., 2009). The [Ca2+]_c increase couples each stimulus and appropriate physiological responses. In the Ca²⁺ signaling pathways, the stimulus-specific [Ca2+]_c pattern (e.g. amplitude and oscillation) provide the critical information for cellular signaling (Scrase-Field and Knight, 2003; Dodd et al., 2010). Therefore, identification of the stimulus-specific [Ca2+]_c signature is crucial for an understanding of the intracellular signaling pathways and physiological responses triggered by each stimulus, as shown in the case of cold acclimation (Knight et al., 1996; Knight and Knight, 2000).Plants often exhibit biphasic [Ca2+]_c increases in response to environmental stimuli. Thus, slow cooling causes a fast [Ca2+]_c transient followed by a second, extended [Ca2+]_c increase in Arabidopsis (Arabidopsis thaliana; Plieth et al., 1999; Knight and Knight, 2000). The Ca²⁺ channel blocker lanthanum (La³⁺) attenuated the fast transient but not the following increase (Knight and Knight, 2000), suggesting that these two [Ca2+]_c peaks have different origins. Similarly, hypoosmotic shock caused a biphasic [Ca2+]_c increase in tobacco (Nicotiana tabacum) suspension-culture cells (Takahashi et al., 1997; Cessna et al., 1998). The first [Ca2+]_c peak was inhibited by gadolinium (Gd³⁺), La³⁺, and the Ca²⁺ chelator EGTA (Takahashi et al., 1997; Cessna et al., 1998), whereas the second [Ca2+]_c increase was inhibited by the intracellular Ca²⁺ store-depleting agent caffeine but not by EGTA (Cessna et al., 1998). The amplitude of the first [Ca2+]_c peak affected the amplitude of the second increase and vice versa (Cessna et al., 1998). These results suggest that even though the two [Ca2+]_c peaks originate from different Ca²⁺ fluxes (e.g. Ca²⁺ influx through the plasma membrane and Ca²⁺ release from subcellular stores, respectively), they are closely interrelated, showing the importance of the kinetic and pharmacological analyses of these [Ca2+]_c increases.Changes in the gravity vector (gravistimulation) could work as crucial environmental stimuli in plants and are generally achieved by rotating the specimens (e.g. +180°) in ground experiments. Use of Arabidopsis seedlings expressing apoaequorin, a Ca²⁺-reporting photoprotein (Plieth and Trewavas, 2002; Toyota et al., 2008a), has revealed that gravistimulation induces a biphasic [Ca2+]_c increase that may be involved in the sensory pathway for gravity perception/response (Pickard, 2007; Toyota and Gilroy, 2013) and the intracellular distribution of auxin transporters (Benjamins et al., 2003; Zhang et al., 2011). These two Ca²⁺ changes have different characteristics. The first transient [Ca2+]_c increase depends on the rotational velocity but not angle, whereas the second sustained [Ca2+]_c increase depends on the rotational angle but not velocity. The first [Ca2+]_c transient was inhibited by Gd³⁺, La³⁺, and the Ca²⁺ chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N′,N′-tetraacetic acid but not by ruthenium red (RR), whereas the second sustained [Ca2+]_c increase was inhibited by all these chemicals. These results suggest that the first transient and second sustained [Ca2+]_c increases are related to the rotational stimulation and the gravistimulation, respectively, and are mediated by distinct molecular mechanisms (Toyota et al., 2008a). However, it has not been demonstrated directly that the second sustained [Ca2+]_c increase is induced solely by gravistimulation; it could be influenced by other factors, such as an interaction with the first transient [Ca2+]_c increase (Cessna et al., 1998), vibration, and/or deformation of plants during the rotation.To elucidate the genuine Ca²⁺ signature in response to gravistimulation in plants, we separated rotation and gravistimulation under microgravity (μg; less than 10⁻⁴g) conditions provided by parabolic flight (PF). Using this approach, we were able to apply rotation and gravistimulation to plants separately (Fig. 1). When Arabidopsis seedlings were rotated +180° under μg conditions, the [Ca2+]_c response to the rotation was transient and almost totally attenuated in a few seconds. Gravistimulation (transition from μg to 1.5g) was then applied to these prerotated specimens at the terminating phase of the PF. This gravistimulation without simultaneous rotation induced a sustained [Ca2+]_c increase. The kinetic properties of this sustained [Ca2+]_c increase were examined under different gravity intensities (0.5g–2g) and sequences of gravity intensity changes (Fig. 2A). This analysis revealed that gravistimulation-specific Ca²⁺ response has an almost linear dependency on gravitational acceleration (0.5g–2g) and an extremely rapid responsiveness of less than 1 s.Open in a separate windowFigure 1.Diagram of the experimental procedures for applying separately rotation and gravistimulation to Arabidopsis seedlings. Rotatory stimulation (green arrow) was applied by rotating the seedlings 180° under μg conditions, and 1.5g 180° rotation gravistimulation (blue arrow) was applied to the prerotated seedlings after μg.Open in a separate windowFigure 2.Acceleration, temperature, humidity, and pressure in an aircraft during flight experiments. A, Accelerations along x, y, and z axes in the aircraft during PF. The direction of flight (FWD) and coordinates (x, y, and z) are indicated in the bottom graph. The inset shows an enlargement of the acceleration along the z axis (gravitational acceleration) during μg conditions lasting for approximately 20 s. B, Temperature, humidity, and pressure in the aircraft during PF. Shaded areas in graphs denote the μg condition.     

Bridging the gap: Membrane contact sites in signaling,metabolism, and organelle dynamics

Regions of close apposition between two organelles, often referred to as membrane contact sites (MCSs), mostly form between the endoplasmic reticulum and a second organelle, although contacts between mitochondria and other organelles have also begun to be characterized. Although these contact sites have been noted since cells first began to be visualized with electron microscopy, the functions of most of these domains long remained unclear. The last few years have witnessed a dramatic increase in our understanding of MCSs, revealing the critical roles they play in intracellular signaling, metabolism, the trafficking of metabolites, and organelle inheritance, division, and transport.

Introduction

The compartmentalization of cells allows the segregation and regulation of the myriad reactions that occur within them. The tremendous benefits of intracellular compartmentalization also come at a price; to function optimally, cells must transmit signals and exchange material between compartments. Numerous mechanisms have evolved to facilitate these exchanges. One that has not been well appreciated until the last few years is the transmission of signals and molecules between organelles that occurs at regions where the organelles are closely apposed, often called membrane contact sites (MCSs). These sites were first characterized because of their critical roles in the intracellular exchange of lipids and calcium, which can be directly channeled between organelles via MCSs. More recently, it has also become apparent that MCSs are important sites for intracellular signaling, organelle trafficking, and inheritance, and that MCSs are specialized regions where regulatory complexes are assembled (English and Voeltz, 2013; Helle et al., 2013).A hallmark of MCSs is that membranes from two organelles (or compartments of the same organelle) are tethered to one another, but not all instances in which membranes interact with or are tethered to one another are considered MCSs. True MCSs have four properties: (1) membranes from two intracellular compartments are tethered in close apposition, typically within 30 nm, (2) the membranes do not fuse (though they may transiently hemi-fuse), (3) specific proteins and/or lipids are enriched at the MCS, and (4) MCS formation affects the function or composition of at least one of the two organelles in the MCS.This review will discuss what we know about proteins that tether organelles, the exchange of small molecules at MCSs, and other emerging functions of MCSs.

MCS tethers

An MCS tether is a protein or complex of proteins (Fig. 1) that simultaneously binds the two apposing membranes at an organelle contact site and plays a role in maintaining the site (English and Voeltz, 2013; Helle et al., 2013). In many cases it is not yet clear if these proteins and complexes are genuine tethers, which are necessary to maintain MCSs, or function at MCSs but are not necessary to sustain contacts. Distinguishing between these possibilities is an important challenge for the field, especially when more than one protein or complex of proteins independently hold together the membranes at an MCS.Open in a separate windowFigure 1.Proteins proposed to mediate tethering at MCSs. Mammalian proteins are shown on a yellow background, yeast proteins on a blue background, and proteins found in both mammals and yeast are on a green background. Tethering complexes not described in the text are indicated with red numbers: (1) StARD3-VAPs (Alpy et al., 2013), (2) NPC1-ORP5 (Du et al., 2011), (3) Psd2-Pdr17 (Riekhof et al., 2014), (4) Vac8-Nvj1 (Pan et al., 2000), (5) Nvj2 (Toulmay and Prinz, 2012), (6) PTPIP51-VAPs (De Vos et al., 2012), (7) Orai1-STIM1 (Nunes et al., 2012), (8) DGAT2-FATP1 (Xu et al., 2012), and (9) IncD-CERT-VAPs (Derré et al., 2011; Elwell et al., 2011).As a growing number of potential tethers are identified, three trends are emerging. First, most MCSs are maintained by several tethers. One of the best-characterized examples of this is the junction of the ER and plasma membrane (PM) in Saccharomyces cerevisiae. Recent work showed that it was necessary to eliminate six ER resident proteins to dramatically reduce the normally extensive interactions between the ER and PM (Manford et al., 2012; Stefan et al., 2013). This suggests that these six proteins mediate tethering independently of each other. Four of the six proteins (three calcium and lipid-binding domain proteins 1–3, also called Tcb1–3, and Ist2) are integral ER membrane proteins that have cytosolic domains that bind the plasma membranes (Fischer et al., 2009; Toulmay and Prinz, 2012). The other two proteins, Scs2 and Scs22 (Scs, suppressor of Ca²⁺ sensitivity), are homologues of mammalian VAPs (vesicle-associated membrane protein–associated proteins). VAPs are integral membrane tail-anchored proteins in the ER that bind proteins containing FFAT (phenylalanines in an acid tract) motifs (Loewen et al., 2003). A number of proteins that contain these motifs also have domains that bind lipids and proteins in the PM, allowing them to simultaneously bind and tether the ER and PM. For example, some oxysterol-binding protein (OSBP)–related proteins (ORPs) have FFAT motifs and pleckstrin homology (PH) domains that bind phosphoinositides (PIPs) in the plasma membrane (Levine and Munro, 1998; Weber-Boyvat et al., 2013). Thus, ORPs and other FFAT motif-containing proteins can mediate ER–PM tethering via VAPs. It should be noted that VAPs and proteins bound by VAPs also mediate tethering between the ER and organelles in addition to the PM. These are shown in Fig. 1.A second emerging trend is that tethering seems to be a dynamic, regulated process, and we are beginning to understand the mechanisms of dynamic apposition of membranes at MCSs by tethers. One example is ER–PM tethering mediated by proteins called extended synaptotagmins (E-Syts), which are homologues of the yeast Tcb tethers. The tethering of the ER and PM by E-Syts is regulated by Ca²⁺ and the PM-enriched lipid PI(4,5)P₂ (Chang et al., 2013; Giordano et al., 2013). Binding of these molecules by E-Syts may control both the extent of ER–PM contact and the distance between these organelles at MCSs. A second example of regulated MCS formation is provided by a recent study on OSBP. This protein and other FFAT motif-containing proteins have been thought to mediate ER–Golgi tethering by simultaneously binding VAPs in the ER and PIPs in the Golgi complex (Kawano et al., 2006; Peretti et al., 2008). In an elegant set of experiments, Mesmin et al. (2013) showed that OSBP regulates its own ability to mediate ER–Golgi tethering by modulating PI4P levels in the Golgi complex. When PI4P levels in the Golgi complex are high, OSBP tethers the ER and Golgi complex and also transports PI4P from the Golgi to the ER. When the PI4P reaches the ER, it is hydrolyzed by the phosphatase Sac1, preventing it from being transferred back to the Golgi. The reduction in Golgi complex PI4P levels by OSBP causes OSBP to dissociate from the Golgi, decreasing ER–Golgi tethering. Thus, OSBP negatively regulates its own tethering of the ER and Golgi membranes. Lipid transport by OSBP and similar proteins will be discussed in more detail in the section on lipid transport at MCSs.The third important feature of many MCS tethering complexes is that most have functions in addition to tethering. This is well illustrated by complexes proposed to mediate ER–mitochondria tethering in mammalian cells, where four such complexes have been described (Fig. 1). For example, Mfn2 (mitofusin-2) acts as a tether (de Brito and Scorrano, 2008), but the primary function of this dynamin-like protein is to mediate mitochondrial fusion. Although Mfn2 is largely in the outer mitochondrial membrane (OMM), a small fraction also resides the ER, and it has been proposed that the interaction of Mfn2 in the ER with Mfn2 in the OMM tethers the ER and mitochondria (de Brito and Scorrano, 2008). The other ER–mitochondria tethering complexes proposed in mammals (Fig. 1) also have additional functions—either Ca²⁺ signaling or apoptotic signaling between these organelles.

Tethers within organelles

MCSs may form not only between organelles but also between compartments of the same organelle. In two cases, proteins necessary for these intra-organelle contacts are known. The Golgi complex is divided into a number of cisternae that remain closely apposed in some cell types, forming stacked compartments. Two tethering proteins maintain connections between Golgi cisternae. Golgi reassembly stacking protein 65 (GRASP65) forms contacts between cis- and medial-Golgi cisternae and GRASP55 mediates medial- to trans-cisternal interactions (Fig. 1; Barr et al., 1997; Shorter et al., 1999). The Golgi stack disassembles when both GRASPs are depleted, indicating that they are the primary or sole tethers (Xiang and Wang, 2010). Tethering by these proteins is regulated by kinases to allow Golgi cisternal disassembly during the cell cycle. Whether the inter-Golgi contacts formed by GRASPs mediate signaling or lipid exchange between cisternae is not yet known (Tang and Wang, 2013).MCSs also form inside organelles with internal membranes: mitochondria, chloroplasts, and multivesicular bodies. These MCSs may form between membranes within these organelles or between internal membranes and the outer membrane of the organelle. Recently, three groups discovered a tethering complex involved in forming contacts between mitochondrial cisternae and between cisternae and the mitochondrial outer membrane (Harner et al., 2011; Hoppins et al., 2011; von der Malsburg et al., 2011). This complex, called the mitochondrial contact site and cristae organizing system (MICOS), is conserved from yeast to humans and contains at least six proteins (Fig. 1). It is necessary to maintain inner membrane organization and also interacts with protein complexes in the outer membrane, including the translocase of the outer membrane (TOM) complex and the sorting and assembly machinery (SAM) complex (van der Laan et al., 2012; Zerbes et al., 2012).

Lipid exchange at MCSs

Lipid exchange between organelles at MCSs may serve a number of important functions. One is that it allows cells to rapidly modulate the lipid composition of an organelle independently of vesicular trafficking. In addition, some organelles, such as mitochondria and chloroplasts, must obtain most of the lipids they require for membrane biogenesis by nonvesicular lipid trafficking that almost certainly occurs at MCSs (Osman et al., 2011; Wang and Benning, 2012; Horvath and Daum, 2013). Finally, and perhaps most importantly, lipid transfer at MCSs may play an important role in lipid metabolism by channeling lipids to or away from enzymes in different compartments.Some lipid exchange at MCSs is facilitated by soluble lipid transport proteins (LTPs), which can shuttle lipid monomers between membranes (Fig. 2 A). In other cases, known LTPs do not seem to be required and lipids may be exchanged at MCSs by other mechanisms (Fig. 2, B and C), which will be discussed next.Open in a separate windowFigure 2.Possible mechanisms of lipid exchange at MCSs. (A) Transfer by LTPs using CERT as an example. The targeting PH domain (pink) and FFAT motif (blue) are shown. CERT could shuttle between membranes (left) or transfer while binding both membranes (right). (B) Some transfer could occur through hydrophobic channels or tunnels (in green) bridging the two membranes at a MCS. (C) Lipid exchange between hemifused membranes. Hemifusion could be promoted and regulated by proteins (red).Most LTPs fall into at least five superfamilies that differ structurally but that all have a hydrophobic pocket or groove that can bind a lipid monomer, and often have a lid domain that shields the bound lipid from the aqueous phase (D’Angelo et al., 2008; Lev, 2010). This allows LTPs to shuttle lipid monomers between membranes. LTPs probably transfer lipids between organelles in cells most efficiently at MCSs, where they have only a short distance to diffuse between membranes. LTPs that may transfer lipids at contact sites are: OSBP, ceramide transport protein (CERT), the yeast OSBP homologues Osh6 and Osh7, protein tyrosine kinase 2 N-terminal domain–interacting receptor 2 (Nir2), and Ups1 (Hanada, 2010; Connerth et al., 2012; Chang et al., 2013; Maeda et al., 2013; Mesmin et al., 2013).LTPs could function by shuttling between membranes at MCSs or while simultaneously bound to both membranes (Fig. 2 A). Many LTPs have domains that target them to the two membranes at an MCS. For example, OSBP and CERT have FFAT motifs, which bind ER resident VAPs, and PH domains that bind PIPs in the Golgi complex or PM.Another important emerging aspect of lipid exchange by some LTPs is that it may be driven by their ability to exchange one lipid for another. For example, OSBP can transfer both cholesterol and PI4P. At ER–Golgi MCSs, OSBP may facilitate the net movement of cholesterol from the ER to the Golgi and PI4P in the opposite direction (Mesmin et al., 2013). The difference in the PI4P concentrations in the ER and Golgi (lower in the ER than in the Golgi) may drive the net transfer of cholesterol to the Golgi. The ability to exchange one lipid for another has been found for other LTPs (Schaaf et al., 2008; de Saint-Jean et al., 2011; Kono et al., 2013) and may be critical for driving directional lipid exchange at MCSs.Some lipid exchange at MCSs does not seem to be facilitated by LTPs. The best evidence for this comes from studies on lipid transfer between the ER and mitochondria. It has long been known that lipids are exchanged between these two organelles; mitochondria must acquire most of the lipid it requires for membrane biogenesis from the rest of the cell. Lipid exchange at ER–mitochondria MCSs occurs by a mechanism that does not require energy, at least in vitro, and does not require any cytosolic factors (Osman et al., 2011; Vance, 2014).How this lipid transfer occurs is not known, and two possible types of mechanism are shown in Fig. 2, B and C. One is that some MCS proteins form a hydrophobic channel that allows lipids to move between membranes. Such a channel would be similar to an LTP, but whereas lipids enter and exit LTPs by the same opening, they enter and exit channels by different openings. This difference could allow lipid exchange by a channel to be regulated and, if the channel could bind two different lipids simultaneously, it might couple the transfer of the lipids. A domain that may form channels at MCSs has been identified. Called the synaptotagmin-like mitochondrial lipid-binding protein (SMP) domain, it has been predicted to be part of a superfamily of proteins that includes cholesterol ester transfer protein (CETP; Kopec et al., 2010). CETP has a tubular lipid-binding domain that transfers lipids between high-density and low-density lipoproteins, probably while simultaneously bound to both (Qiu et al., 2007; Zhang et al., 2012). SMP domains could transfer lipids between membranes by a similar mechanism. Consistent with this possibility, all SMP-containing proteins in budding yeast localize to MCSs and many mammalian SMP-containing proteins do as well (Toulmay and Prinz, 2012). Interestingly, SMP domains are present in three of the five proteins in a yeast ER–mitochondria tethering complex called ERMES (Kornmann et al., 2009). Whether ERMES facilitates lipid exchange between the ER and mitochondria is not yet clear. Mitochondria derived from cells missing ERMES have altered lipid composition (Osman et al., 2009; Tamura et al., 2012; Tan et al., 2013), indicating that lipid exchange between the ER and mitochondria could be altered in these strains. On the other hand, little or no defect in the rates of phospholipid exchange between ER and mitochondria were found in ERMES mutants (Kornmann et al., 2009; Nguyen et al., 2012; Voss et al., 2012). Thus, whether proteins that contain SMP domains actually facilitate lipid exchange remains to be determined.As second possible mechanism of lipid transfer at MCSs that does not require LTPs is membrane hemifusion (Fig. 2 C), which could allow rapid exchange of large amounts of lipids between compartments. Recent indirect evidence suggests that hemifusion may occur between the ER and chloroplasts (Mehrshahi et al., 2013). This is consistent with an earlier study using optical tweezers that found the ER and chloroplasts remained attached to one another even when a stretching force of 400 pN was applied (Andersson et al., 2007). Whether hemifusion occurs at MCSs in animal cells remains to be determined.

Calcium signaling at MCSs

MCSs between the ER and PM and the ER and mitochondria play central roles in intracellular Ca²⁺ storage, homeostasis, and signaling in mammalian cells. MCSs between the ER and lysosomes may also be important, though they are less well understood (Helle et al., 2013; Lam and Galione, 2013).One of the best-characterized MCSs is the one formed between the PM and ER in muscle cells. In both cardiac and skeletal muscle cells, deep invaginations of the PM, called T (transverse)-tubules, allow it to form extensive contacts with the ER, called the sarcoplasmic reticulum (SR) in muscle cells. These contacts are essential for coupling excitation and contraction. Before excitation, Ca²⁺ levels in the cytoplasm of muscle cells are low, whereas the Ca²⁺ concentrations in the SR and outside muscle cells are high. During muscle excitation, Ca²⁺ rapidly flows into the cytosol through channels in the PM and the SR (Fig. 3 A). The channels in the PM, called dihydropyridine receptors (DHPRs), and those in the SR, known ryanodine receptors RyRs, directly interact with each other where the SR and PM are closely apposed, allowing the opening of both types of channels to be coordinated (Fabiato, 1983; Bannister, 2007; Beam and Bannister, 2010; Rebbeck et al., 2011).Open in a separate windowFigure 3.Ca²⁺ trafficking at ER–PM MCSs. (A) In muscle cells, the interaction of the RyR in the SR and with DHPR in the PM allows the coordinated release of Ca²⁺ during muscle excitation and contraction. See text for details. (B) When STIM1 senses low Ca²⁺ concentration in the ER, it undergoes a conformational change that allows it to oligomerize and bind to the PM, to the protein Orai1, and to accumulate at ER–PM MCSs. Ca²⁺ influx at these sites facilitates Ca²⁺ import into the ER by sarco/endoplasmic reticulum Ca²⁺-ATPase (SERCA). (C) Calcium channeling from the ER lumen to the mitochondrial matrix. Calcium exits the ER through the inositol trisphosphate receptor (IP3R) channel, enters mitochondria via VDAC, and then uses the mitochondrial Ca²⁺ uniporter (MCU) to move into the mitochondrial matrix.The extensive contacts between the SR and PM in muscle cells are largely maintained by tethering proteins called junctophilins, which have a single transmembrane domain in the SR and a large cytosolic domain that interacts with the PM. Expression of junctophilins in cells lacking them induces ER–PM contacts (Takeshima et al., 2000) and cells lacking junctophilins have abnormal SR–PM MCSs and defects in Ca²⁺ signaling (Ito et al., 2001; Komazaki et al., 2002; Hirata et al., 2006). Thus, junctophilins are both necessary and sufficient for generating functional SR–PM contacts. However, cells lacking junctophilins still maintain some SR–PM contacts, indicating that other proteins also tether the SR and the PM. Some of this residual tethering probably comes from the interaction of DHPRs and RyRs.ER–PM contacts also play a role in regulating intracellular Ca²⁺ levels in non-excitable cells. When the Ca²⁺ concentration in the ER lumen is low it triggers Ca²⁺ entry into the cytosol and ER from outside cells (Fig. 3 B), a process known as store-operated Ca²⁺ entry (SOCE). The PM channel responsible for Ca²⁺ entry is Orai1, and the sensor of Ca²⁺ concentration in the ER lumen is the integral membrane protein stromal interaction molecule-1 (STIM1). When STIM1 senses that the Ca²⁺ concentration in the ER is low, it oligomerizes and undergoes a conformational change that exposes a basic cluster of amino acids in its C terminus that binds PIPs in the PM (Stathopulos et al., 2006, 2008; Liou et al., 2007; Muik et al., 2011). STIM1 also binds to Orai1 in the PM and activates it (Kawasaki et al., 2009; Muik et al., 2009; Park et al., 2009; Wang et al., 2009). Activation of STIM1 causes it to shift from being relatively evenly distributed on the ER to forming a number of puncta, which are regions were the ER and PM are closely apposed. It seems likely that STIM1 accumulates at and expands preexisting ER–PM MCSs and may also drive the formation of new MCSs (Wu et al., 2006; Lur et al., 2009; Orci et al., 2009).The interaction of STIM1 and Orai1 at ER–PM contacts during SOCE is an elegant mechanism for channeling both signals and small molecules at an MCS. The signal that ER luminal Ca²⁺ concentration is low is transmitted directly from STIM1 in the ER to Orai1 in the PM. The close contact of PM and ER also allows Ca²⁺ to move from outside the cell into the lumen of the ER without significantly increasing cytosolic Ca²⁺ levels (Jousset et al., 2007). During SOCE, ER Ca²⁺ levels are restored by the sarco/endoplasmic reticulum Ca²⁺-ATPase (SERCA) pump (Sampieri et al., 2009; Manjarrés et al., 2011). This pump is enriched in ER–PM contacts with STIM1 and may interact directly with it, suggesting how Ca²⁺ can be effectively channeled from outside cells directly into the ER lumen at ER–PM MCSs (Fig. 3 B).Interestingly, it has become clear that proteins that are not part of the SOCE pathway also facilitate ER–PM connections during Ca²⁺ signaling. The E-Syts have multiple domains that probably bind Ca²⁺. They have been shown to regulate both the number of the ER–PM contacts and the distance between the ER and PM at MCSs during Ca²⁺ signaling (Chang et al., 2013; Giordano et al., 2013).MCSs between the ER and mitochondria similarly facilitate Ca²⁺ movement from the ER lumen to mitochondria (Rizzuto et al., 1998; Csordás et al., 2006). Ca²⁺ channels in the ER and OMM interact with each other at MCSs (Fig. 3 C). The channel in the ER is called the inositol trisphosphate receptor (IP3R), while the voltage-dependent anion channel (VDAC) in the outer mitochondrial membrane is a nonspecific pore that allows Ca²⁺ entry into mitochondria. These proteins, together with the cytosolic chaperone Grp75, form a complex that links the ER and mitochondria and facilitates Ca²⁺ exchange (Szabadkai et al., 2006).More evidence that Ca²⁺ transfer from the ER to mitochondria occurs at MCSs came from studies on the channel that allows Ca²⁺ to move across the inner mitochondrial membrane, called the mitochondrial Ca²⁺ uniporter (MCU). Surprisingly, this channel has an affinity for Ca²⁺ that is lower than the typical Ca²⁺ concentration in the cytosol (Kirichok et al., 2004). However, Ca²⁺ release by the ER at ER–mitochondrial MCSs suggests a solution to this puzzle; the local Ca²⁺ concentration at these MCSs is probably high enough for MCU to function (Csordás et al., 2010). Close contacts between the ER and mitochondria are therefore essential for channeling Ca²⁺ from the ER lumen to the mitochondrial matrix.It is thought that MCSs between the ER (or SR) and lysosomes regulate Ca²⁺ release by lysosomes, but the mechanism is not yet understood (Kinnear et al., 2004, 2008; Galione et al., 2011; Morgan et al., 2011).

Enzymes working in trans and signaling at MCSs

MCSs allow rapid and efficient signaling between intracellular compartments. We are still just beginning to understand the mechanisms and functions of this signaling. One way that signals are transmitted between the two compartments at an MCS is for an enzyme in one compartment to modify substrates in the second; that is, for the enzyme to work in trans. Although there are currently only a few examples of this, which are discussed here, it seems likely that many more will be uncovered.The protein tyrosine phosphatase PTP1B regulates a number of receptor tyrosine kinases. PTP1B resides on the surface of the ER with its active site in the cytosol, and yet the receptor tyrosine kinases it modifies are in the PM. Although this was initially puzzling, it was found that PTP1B probably encounters its substrates at MCSs, either at ER–PM junctions or at contacts between the ER and endocytic recycling compartments (Haj et al., 2002; Boute et al., 2003; Anderie et al., 2007; Eden et al., 2010; Nievergall et al., 2010). Interestingly, in some cases the interaction of PTP1B with substrates in the PM occurs on portions of the PM that are part of cell–cell contacts (Haj et al., 2012), suggesting that ER–PM contacts could play a role in signaling, not only between the ER and PM but between cells as well. Dephosphorylation of receptor tyrosine kinases by PTP1B at contact sites probably allows their kinase activity to be regulated in response to changes in the ER or changes in cellular architecture that alter MCSs. For example, the dephosphorylation of epidermal growth factor receptor (EGFR) by PTP1B occurs at regions of close contact between the ER and multivesicular bodies, causing EGFR to become sequestered with multivesicular bodies (Eden et al., 2010). This may provide a mechanism for cells to regulate EGFR levels on the PM in response to signals in the ER.Lipid metabolism enzymes can also work in trans at MCSs. In two cases, both in yeast, enzymes that reside in the ER have been found to modify lipids in the PM at MCSs. In one instance, the phosphatase Sac1, which is on the surface of the ER, can dephosphorylate PIPs in the PM (Stefan et al., 2011). In the second, the ER enzyme Opi3 methylates phosphatidylethanolamine in the PM, a reaction that is required for the conversion of phosphatidylethanolamine to phosphatidylcholine (Tavassoli et al., 2013). Remarkably, the PIP-binding protein Osh3 (Tong et al., 2013) regulates both reactions, suggesting that lipid metabolism at ER–PM junctions is regulated by PIPs. It seems likely that ER–PM junctions play important roles in integrating lipid metabolism in both organelles.

MCSs and organelle trafficking and inheritance

In addition to being sites at which signals and small molecules are exchanged between cellular compartments, there is growing evidence that MCS formation also regulates organelle trafficking and inheritance.In budding yeast, organelle transport is polarized from the mother cell to the growing bud and is required for proper organelle inheritance. The transport of peroxisomes and mitochondria to the bud is regulated by their association with the ER or PM.Knoblach et al. (2013) found that tethering of the ER to peroxisomes requires Pex3, an integral membrane protein that resides in both compartments, and Inp1, a cytosolic protein that binds to Pex3. This tether keeps peroxisomes in mother cells. When peroxisomes divide they are transferred to the bud by the myosin V motor Myo2 and become attached to the ER in the bud. In cells lacking the ER–peroxisome tether, peroxisomes accumulate in daughter cells. Thus, tethering plays a critical role in ensuring that some peroxisomes are retained in mother cells and that both cells inherit peroxisomes.Mitochondrial inheritance in yeast is regulated by close contacts with both the ER and PM. Mitochondria–PM contacts mediated by a complex containing Num1 and Mdm36 ensure that mitochondria are properly distributed between mother and daughter cells and seem to be particularly important for retaining mitochondria in the mother cells (Klecker et al., 2013; Lackner et al., 2013). Interestingly, Num1–Mdm36-mediated contacts also associate with the ER (Lackner et al., 2013), suggesting that three membranes may somehow associate at these MCSs. An ER–mitochondria tether containing the protein Mmr1, which anchors mitochondria to bud tips, also plays a role in mitochondrial inheritance (Swayne et al., 2011). Thus, the Num1-tethering complex and Mmr1-tethering complex seem to play antagonistic roles in mitochondrial distribution; the Num1 complex promotes mitochondrial retention in the mother, whereas the Mmr1 complex favors retention in the bud.MCSs also play a role in endosomal trafficking in mammalian cells. One of the complexes that tethers the ER to endosomes contains VAPs and ORP1L, which is an OSBP homologue that can bind cholesterol (Fig. 1). ORPlL can also binds the p150^Glued subunit of the dynein–dynactin motor that participates in endosome transport along microtubules (Johansson et al., 2007). When cellular cholesterol levels are high, ORP1L associates with p150^Glued but not VAPs and endosomes are transported on microtubules. However, when cholesterol levels decrease, ORP1L undergoes a conformation change that dissociates it from p150^Glued and allows it to bind to VAPs on the surface of the ER, thus forming a tether between endosomes and the ER (Rocha et al., 2009). Under these conditions, endosome transport on microtubules is blocked. ORPlL is therefore a cholesterol sensor that regulates a switch between the association of endosomes with either motors or the ER.

MCSs and organelle division

A groundbreaking study revealed a new and unexpected role for MCSs between the ER and mitochondria: the ER regulates mitochondrial fission (Friedman et al., 2011). Although a mechanistic understanding of how ER participates in mitochondrial fission is not yet available, the sequence of events is beginning to come into focus (Fig. 4). The ER encircles mitochondria at sites where scission will occur. The ERMES complex is present at these sites (Murley et al., 2013). Because mammalian cells lack ERMES, another tethering complex must perform the same function in higher eukaryotes. Mitochondrial division requires membrane scission by the dynamin-like protein Dnm1/Drp1, which multimerizes on the outer mitochondria membrane. Close contacts between the ER and mitochondria occur before Dnm1/Drp1 assembly, suggesting that these contacts promote or regulate the association of Dnm1/Drp1 with mitochondria and hence mitochondrial division. It is possible that when the ER encircles mitochondria it causes mitochondria to constrict to a diameter that allows Dnm1/Drp1 to assemble. The force necessary to drive constriction may come from actin polymerization. A recent study found that the ER protein, inverted formin-2, probably drives actin polymerization at these sites and is necessary for mitochondria fusion (Korobova et al., 2013).Open in a separate windowFigure 4.Model of ER-mediated regulation of mitochondrial fission at sites of contact. (A) The ER and mitochondria are tethered by ERMES in yeast (other tethers are used in higher eukaryotes). (B) The ER encircles mitochondria at sites where division will occur. (C) Actin polymerization facilitated by formin 2 may cause mitochondrial constriction. (D) The dynamin-like protein Drp1 is recruited to the mitochondrial surface, where it multimerizes and causes mitochondrial scission. (E) After fission, the ER remains associated with the mitochondrion that retains the ERMES complex.Understanding the assembly and regulation of the mitochondrial division machinery at ER–mitochondria MCSs and how this is linked to mitochondrial and perhaps ER function remain fascinating questions for the future. Another interesting question is whether other MCSs play roles in the fission of other organelles.

Proposed functions of ER–mitochondrial MCSs

A growing number of studies have suggested that ER–mitochondria MCSs play critical roles in autophagy, apoptosis, inflammation, reactive oxygen species signaling, and metabolic signaling. ER–mitochondria MCSs have also been implicated in Alzheimer’s disease, Parkinson’s disease, and some viral infections. These topics have been recently reviewed (Eisner et al., 2013; Raturi and Simmen, 2013; Marchi et al., 2014; Vance, 2014) and will not be discussed in detail here.One issue with most of the studies on the functions of ER–mitochondria junctions is that they rely, at least in part, on density gradient purification of the ER that associates with mitochondria. These operationally defined membranes, often called mitochondrial-associated membranes (MAMs), remain poorly defined. In fact, a significant number of proteins that are enriched in MAMs do not seem to be enriched at ER–mitochondria junctions when their localization is determined by other methods (Helle et al., 2013; Vance, 2014). Therefore, it remains unclear why some proteins and lipids are enriched in MAMs.Here, two interesting findings will be discussed that suggest the importance of ER–mitochondrial junctions in signaling in addition to their well-known role in Ca²⁺ signalling.The induction of apoptosis requires signal transmission between the ER and mitochondria. Part of this signaling process occurs through an interaction between the ER protein Bap31 and the mitochondrial fission protein Fission 1 homologue (Fis1; Iwasawa et al., 2011). This interaction occurs at ER–mitochondria MCSs and results in the cleavage of Bap31 by caspase-8 to form p20Bap31, which is pro-apoptotic. Both Bap31 and Fis1 are parts of larger complexes that are still being characterized. Interestingly, it has recently been found that a protein called cell death–involved p53 target-1 (CDIP1) binds to Bap31 during ER stress and promotes apoptotic signaling from the ER to mitochondria (Namba et al., 2013), suggesting how ER stress signals are transmitted from the ER to mitochondria through MCSs.Another important connection between ER–mitochondrial MCSs and signaling has to do with the target of rapamycin (TOR) kinase complexes, which are critical regulators of growth and metabolism. The mammalian TOR complex 2 (mTORC2) was found to interact with the IP3R–Grp75–VDAC complex that tethers the ER and mitochondria (Betz et al., 2013). Remarkably, this study presents evidence that mTORC regulates both the formation of ER–mitochondrial MCSs and mitochondrial function, suggesting an interesting new mechanism for how metabolic signaling can impact mitochondrial function via MCSs.

Conclusions and perspectives

The potential of MCSs to facilitate Ca²⁺ signaling and channel lipids between organelles was recognized some time ago (Levine and Loewen, 2006), but it has only been in the last few years that we have finally begun to have some mechanistic insight into how these processes occur and how MCSs are formed. Many fundamental questions remain to be addressed. How lipid exchange at MCSs that does not require soluble LTPs occurs or whether transient hemifusion of membranes at MCS ever occurs remain open questions. Another is the mechanisms by which Ca²⁺ regulates MCS formation between the ER and other organelles. One major challenge for the field will be devising better methods to visualize MCSs and identify proteins and lipids enriched at these sites. It is particularly important to better understand what the MAM fraction is and what it means for proteins and lipids to be enriched in this fraction.One of the most exciting developments in the study of MCSs in the last few years has been the discovery of the role of MCSs in organelle trafficking, inheritance, and dynamics. These studies have revealed that MCSs not only play critical roles in signaling and metabolism, but also modulate the intracellular distribution of organelles and organelle architecture. Understanding how MCSs perform these functions will probably shed light on the connection between the still murky relationship between organelle structure and function as well as the role of the ER as a regulator of other organelles. Given the current pace of discovery, it seems likely that in the next few years our knowledge of the functions of MCSs will grow dramatically.     

Mitochondrial trafficking and anchoring in neurons: New insight and implications

Mitochondria are essential organelles for neuronal growth, survival, and function. Neurons use specialized mechanisms to drive mitochondria transport and to anchor them in axons and at synapses. Stationary mitochondria buffer intracellular Ca²⁺ and serve as a local energy source by supplying ATP. The balance between motile and stationary mitochondria responds quickly to changes in axonal and synaptic physiology. Defects in mitochondrial transport are implicated in the pathogenesis of several major neurological disorders. Recent work has provided new insight in the regulation of microtubule-based mitochondrial trafficking and anchoring, and on how mitochondrial motility influences neuron growth, synaptic function, and mitophagy.

Introduction

Mitochondria are cellular power plants that convert chemicals into ATP through coupled electron transport chain and oxidative phosphorylation. A cortical neuron in human brain utilizes up to ∼4.7 billion ATP molecules per second to power various biological functions (Zhu et al., 2012). Constant ATP supply is essential for nerve cell survival and function (Nicholls and Budd, 2000). Mitochondrial ATP production supports synapse assembly (Lee and Peng, 2008), generation of action potentials (Attwell and Laughlin, 2001), and synaptic transmission (Verstreken et al., 2005). Synaptic mitochondria maintain and regulate neurotransmission by buffering Ca²⁺ (Medler and Gleason, 2002; David and Barrett, 2003). In addition, mitochondria sequester presynaptic Ca²⁺ transients elicited by trains of action potentials and release Ca²⁺ after stimulation, thus inducing certain forms of short-term synaptic plasticity (Werth and Thayer, 1994; Tang and Zucker, 1997; Billups and Forsythe, 2002; Levy et al., 2003; Kang et al., 2008). Removing mitochondria from axon terminals results in aberrant synaptic transmission likely due to insufficient ATP supply or altered Ca²⁺ transients (Stowers et al., 2002; Guo et al., 2005; Ma et al., 2009).Neurons are polarized cells consisting of a relatively small cell body, dendrites with multiple branches and elaborate arbors, and a thin axon that can extend up to a meter long in some peripheral nerves. Due to these extremely varied morphological features, neurons face exceptional challenges to maintain energy homeostasis. Neurons require specialized mechanisms to efficiently distribute mitochondria to far distal areas where energy is in high demand, such as synaptic terminals, active growth cones, and axonal branches (Fig. 1; Morris and Hollenbeck, 1993; Ruthel and Hollenbeck, 2003). Axonal branches and synapses undergo dynamic remodeling during neuronal development and in response to synaptic activity, thereby changing mitochondrial trafficking and distribution. Neurons are postmitotic cells surviving for the lifetime of the organism. A mitochondrion needs to be removed when it becomes aged or dysfunctional. Mitochondria also alter their motility and distribution under certain stress conditions or when their integrity is impaired (Miller and Sheetz, 2004; Chang and Reynolds, 2006; Cai et al., 2012). Therefore, efficient regulation of mitochondrial trafficking and anchoring is essential to: (1) recruit and redistribute mitochondria to meet altered metabolic requirements; and (2) remove aged and damaged mitochondria and replenish healthy ones at distal terminals. Research into neuronal regulation of mitochondrial trafficking and anchoring is thus a very important frontier in neurobiology. This review article focuses on new mechanistic insight into the regulation of microtubule (MT)-based mitochondrial trafficking and anchoring and provides an updated overview of how mitochondrial motility influences neuronal growth, synaptic function, and mitochondrial quality control. Additional insight and overviews from different perspectives can be found in other in-depth reviews (Frederick and Shaw, 2007; Morfini et al., 2009; Hirokawa et al., 2010; MacAskill and Kittler, 2010; Schon and Przedborski, 2011; Court and Coleman, 2012; Saxton and Hollenbeck, 2012; Sheng and Cai, 2012; Birsa et al., 2013; Lovas and Wang, 2013; Schwarz, 2013).Open in a separate windowFigure 1.Mitochondrial trafficking and anchoring in neurons. Due to complex structural features, neurons require specialized mechanisms trafficking mitochondria to their distal destinations and anchoring them in regions where metabolic and calcium homeostatic capacity is in a high demand. The figure highlights transporting mitochondria to a presynaptic bouton (A) and an axonal terminal (B). MT-based long-distance mitochondrial transport relies on MT polarity. In axons, the MT’s plus ends (+) are oriented toward axonal terminals whereas minus ends (−) are directed toward the soma. Thus, KIF5 motors are responsible for anterograde transport to distal synaptic terminals whereas dynein motors return mitochondria to the soma. The motor adaptor Trak proteins can mediate both KIF5- and dynein-driven bi-directional transport of axonal mitochondria (van Spronsen et al., 2013). Myosin motors likely drive short-range mitochondrial movement at presynaptic terminals where enriched actin filaments constitute cytoskeletal architecture. Motile mitochondria can be recruited into stationary pools via dynamic anchoring interactions between syntaphilin and MTs (Kang et al., 2008) or via an unidentified actin-based anchoring receptor (Chada and Hollenbeck, 2004). Such anchoring mechanisms ensure neuronal mitochondria are adequately distributed along axons and at synapses, where constant energy supply is crucial (figure courtesy of Qian Cai, Department of Cell Biology and Neuroscience, Rutgers University, Piscataway, NJ).

Neuronal mitochondria display complex motility patterns

The complex motility patterns of mitochondrial transport in neurons can be visualized by applying time-lapse imaging in live primary neurons. Mitochondria are labeled by expressing DsRed-Mito, a mitochondria-targeted fluorescent protein, or by loading fluorescent dye MitoTracker Green or MitoTracker Red CMXRos, which stains mitochondria in live cells dependent upon membrane potential. Mitochondria move bi-directionally over long distances along processes, pause briefly, and move again, frequently changing direction. Motile mitochondria can become stationary, and stationary ones can be remobilized and redistributed in response to changes in metabolic status and synaptic activity. Given frequent pauses and bi-directional movements, the mean velocities of neuronal mitochondrial transport are highly variable, ranging from 0.32 to 0.91 µm/s (Morris and Hollenbeck, 1995; MacAskill et al., 2009a). In mature neurons, ∼20–30% of axonal mitochondria are motile, some of which pass by presynaptic terminals while the remaining two thirds are stationary (Kang et al., 2008). Using dual-channel imaging of mitochondria and synaptic terminals in hippocampal neurons, a recent study identified five motility patterns of axonal mitochondria including: stationary mitochondria sitting out of synapses (54.07 ± 2.53%) or docking at synapses (16.29 ± 1.66%); motile mitochondria passing through synapses (14.77 ± 1.58%); or pausing at synapses for a short (<200 s, 7.01 ± 1.29%) or a longer period of time (>200 s, 8.30 ± 1.52%; Sun et al., 2013). These findings are consistent with a previous study in primary cortical neurons (Chang et al., 2006).Altered mitochondrial transport is one of the pathogenic changes in major adult-onset neurodegenerative diseases (Sheng and Cai, 2012). Although in vitro live imaging in primary neurons prepared from embryonic or newborn mice provides an important tool to study mechanisms regulating mitochondrial transport, it has obvious limitations for investigating adult-onset disease mechanisms. For example, reduced transport of axonal mitochondria was consistently reported in amyotrophic lateral sclerosis–linked hSOD1^G93A mice (De Vos et al., 2007; Magrané and Manfredi, 2009; Bilsland et al., 2010; Zhu and Sheng, 2011; Marinkovic et al., 2012; Song et al., 2013). Therefore, it is critical to establish spinal motor neuron cultures directly isolated from adult mice, which will provide more reliable models to investigate mitochondrial transport defects underlying adult-onset pathogenesis in motor neurons. Given that the morphology and conditions of neurons grown in culture dishes differ from that of neurons in vivo, an in vivo transport study would better reflect physiological environments. An ex vivo study of mitochondrial transport in acute nerve explants was developed in transgenic mice, in which neuronal mitochondria are labeled with mito-CFP or mito-YFP (Misgeld et al., 2007).

Moving mitochondria along MTs

Mitochondrial transport between the soma and distal processes or synapses depends upon MT-based motors, which drive their cargos via mechanisms requiring ATP hydrolysis (Martin et al., 1999; Hirokawa et al., 2010). MTs are uniformly arranged in axons: their plus ends are oriented distally and minus ends are directed toward the soma. Such a uniform polarity has made axons particularly useful for exploring mechanisms regulating bi-directional transport: dynein motors drive retrograde movement, whereas kinesin motors mediate anterograde transport. Of the 45 kinesin motor genes identified, the kinesin-1 family (KIF5) is the key motor driving mitochondrial transport in neurons (Hurd and Saxton, 1996; Tanaka et al., 1998; Górska-Andrzejak et al., 2003; Cai et al., 2005; Pilling et al., 2006). KIF5 heavy chain (KHC) contains a motor domain with ATPase at the N terminus and a C-terminal tail for binding cargo directly or indirectly via a cargo adaptor. Three isoforms (KIF5A, KIF5B, and KIF5C) of the KIF5 family are found in mammals, of which KIF5B is expressed ubiquitously whereas KIF5A and KIF5C are expressed selectively in neurons (Kanai et al., 2000).KIF5 motors attach to mitochondria through adaptor proteins. The motor–adaptor coupling ensures targeted trafficking and effective regulation of mitochondrial transport. Drosophila protein Milton acts as a KIF5 motor adaptor by binding the KIF5 cargo-binding domain and mitochondrial outer membrane receptor Miro (Stowers et al., 2002; Glater et al., 2006). The Drosophila mutation in Milton reduces mitochondrial trafficking to synapses. Milton appears specific for mitochondrial trafficking because the trafficking of other cargoes such as synaptic vesicles is not affected. Two Milton orthologues, Trak1 and Trak2, are found in mammals and are required for mitochondrial trafficking (Smith et al., 2006; MacAskill et al., 2009b; Koutsopoulos et al., 2010). Elevated Trak2 expression in hippocampal neurons robustly enhances axonal mitochondrial motility (Chen and Sheng, 2013). Conversely, depleting Trak1 or expressing its mutants results in impaired mitochondrial motility in axons (Brickley and Stephenson, 2011). A recent study revealed that mammalian Trak1 and 2 contain one N-terminal KIF5B-binding site and two dynein/dynactin-binding sites, one at the N-terminal domain and the other at the C-terminal domain (van Spronsen et al., 2013). Thus, the Trak proteins can mediate both KIF5- and dynein-driving bi-directional transport of axonal mitochondria (Fig. 1). In contrast to the specific role of Drosophila Milton in mitochondrial transport, mammalian Trak proteins are also recruited to endosomal compartments and associate with GABAA (γ-aminobutyric acid A) receptors and K⁺ channels, highlighting their role in neuronal cargo transport (Grishin et al., 2006; Kirk et al., 2006; Webber et al., 2008).The mitochondria outer membrane receptor Miro is a Rho-GTPase with two Ca²⁺-binding EF-hand motifs and two GTPase domains (Frederick et al., 2004; Fransson et al., 2006). Mutation of the Drosophila miro gene impairs mitochondrial anterograde transport, thus depleting mitochondria at distal synaptic terminals (Guo et al., 2005). The first crystal structures of Drosophila melanogaster Miro comprising the tandem EF-hands and GTPase domains were recently resolved (Klosowiak et al., 2013). From these structures, two additional hidden EF-hands were identified, each one pairing with a canonical EF-hand. These structures will help our understanding of the conformational changes in Miro upon Ca²⁺ binding and nucleotide sensing in regulating mitochondrial motility, dynamics, and function. There are two mammalian Miro isoforms, Miro1 and Miro2. The Miro1–Trak2 adaptor complex regulates mitochondrial transport in hippocampal neurons (MacAskill et al., 2009b). Elevated Miro1 expression increases mitochondrial transport, likely by recruiting more Trak2 and motors to the mitochondria (Chen and Sheng, 2013). Thus, KIF5, Milton (Trak), and Miro assemble into the transport machinery that drives mitochondrial anterograde movement (Fig. 2 A). An alternative KIF5 adaptor for neuronal mitochondria is syntabulin (Fig. 2 B; Cai et al., 2005). Depleting syntabulin or blocking the KIF5–syntabulin coupling reduces mitochondrial anterograde transport in axons. Several other proteins are also suggested as candidate KIF5 motor adaptors for mitochondrial transport, including FEZ1 (fasciculation and elongation protein zeta-1) and RanBP2 (Ran-binding protein 2; Cho et al., 2007; Fujita et al., 2007), although their role in neuronal mitochondrial transport requires further investigation. The existence of multiple motor adaptors may highlight the complex regulation of mitochondrial motility in response to various physiological and pathological signals.Open in a separate windowFigure 2.Activity-dependent regulation of mitochondrial transport. (A) The Miro–Milton (or Miro–Trak) adaptor complex mediates KIF5-driven mitochondrial transport. (B) Syntabulin, FEZ1, and RanBP2 serve as an alternative KIF5 motor adaptor in driving mitochondrial anterograde transport. (C and D) Miro-Ca²⁺ models in regulating mitochondrial motility. Miro contains Ca²⁺-binding EF-hand motifs. The C-terminal cargo-binding domain of KIF5 motors binds to the Miro–Trak adaptor complex. (C) Ca²⁺ binding to Miro’s EF-hands induces the motor domain to disconnect with MTs and thus prevents motor–MT engagement (Wang and Schwarz, 2009). (D) Alternatively, Ca²⁺ binding releases KIF5 motors from mitochondria (MacAskill et al., 2009a). Thus, Ca²⁺ influx after synaptic activity arrests motile mitochondria at activated synapses. (E) Syntaphilin-mediated “engine-switch and brake” model. A Miro-Ca²⁺–sensing pathway triggers the binding switch of KIF5 motors from the Miro–Trak adaptor complex to anchoring protein syntaphilin, which immobilizes axonal mitochondria via inhibiting motor ATPase activity. Thus, syntaphilin turns off the “KIF5 motor engine” by sensing a “stop sign” (elevated Ca²⁺) and putting a brake on mitochondria, thereby anchoring them in place on MTs. When in their stationary status, KIF5 motor–loaded mitochondria remain associated with the MT tracks while KIF5 ATPase is in an inactive state (Chen and Sheng, 2013; Figure courtesy of Qian Cai).Dynein motors are composed of multiple chains including heavy chains (DHC) as the motor domain, intermediate (DIC), light intermediate (DLIC), and light chains (DLC), which function by associating with cargos and regulating motility. Dynein associates with Drosophila mitochondria, and mutations of DHC alter velocity and run length of mitochondrial retrograde transport in axons (Pilling et al., 2006). Miro may serve as an adaptor for both KIF5 and dynein motors in Drosophila (Guo et al., 2005; Russo et al., 2009), as loss of dMiro impairs both kinesin- and dynein-driven transport and overexpressing dMiro also alters mitochondrial transport in both directions. These findings support an attractive model in which the relative activity of the opposite-moving motors is regulated by their cargo–adaptor proteins.

Mitochondrial motility and membrane dynamics

Mitochondrial trafficking, anchoring, and membrane dynamics are coordinated, thus controlling their shape, integrity, and distribution in neurons. It is expected that the size of a mitochondrion may have a direct impact on its mobility. Indeed, interconnected mitochondrial tubules found in fission-deficient neurons are less efficiently transported to distal synapses. Hippocampal neurons expressing defective Drp1, a mitochondrial fission protein, display accumulated mitochondria within the soma and reduced mitochondrial density in dendrites (Li et al., 2004). Drosophila Drp1 is required for delivering mitochondria to neuromuscular junctions (Verstreken et al., 2005). Conversely, mutation of Miro results in mitochondrial fragmentation in addition to impaired motility, whereas overexpressing Miro not only enhances mitochondrial transport but also induces their interconnection (Fransson et al., 2006; Saotome et al., 2008; Liu and Hajnóczky, 2009; MacAskill et al., 2009b). Additionally, yeast Miro, Gem1p, regulates mitochondrial morphology; cells lacking Gem1p contain collapsed, globular, or grape-like mitochondria (Frederick et al., 2004).Mitochondrial fusion events require two organelles to move closer. A “kiss-and-run” interplay is proposed for driving mitochondrial fusion (Liu et al., 2009). Thus, motile mitochondria, particularly in distal axons and at synapses, undergo fusion events at a higher rate than anchored stationary mitochondria. Recent studies highlight the cross talk between the mitochondrial fusion protein Mfn2 and a motor adaptor complex. Mfn2 associates with Miro2 and regulates axonal mitochondrial transport by affecting kinesin motor processivity or by coordinating the motor switch between kinesin and dynein (Misko et al., 2010). Neurons defective in Mfn2 display reduced mitochondrial motility and altered mitochondrial distribution (Chen et al., 2003, 2007; Baloh et al., 2007). Therefore, regulating mitochondrial trafficking directly affects their morphology. Conversely, changing mitochondrial fusion–fission dynamics also impacts their transport and distribution.

Anchoring mitochondria at MTs

Proper axonal and synaptic function requires stationary mitochondria docking in regions where energy is in high demand. The diffusion capacity of intracellular ATP is rather limited (Hubley et al., 1996), particularly within long axonal processes (Sun et al., 2013). Therefore, docked mitochondria ideally serve as stationary power plants for stable and continuous ATP supply necessary to maintain the activity of Na⁺–K⁺ ATPase, fast spike propagation, and synaptic transmission. In addition, mitochondria maintain calcium homeostasis at synapses by sequestering intracellular [Ca²⁺] (Billups and Forsythe, 2002; Kang et al., 2008). Given the fact that two thirds of the mitochondria are stationary for an extended period along axons, neurons require mechanisms dissociating mitochondria from motor proteins or anchoring mitochondria to the cytoskeleton. One intriguing mitochondrial-anchoring protein is syntaphilin, which is a “static anchor” for immobilizing mitochondria specifically in axons (Kang et al., 2008; Chen et al., 2009). Syntaphilin targets axonal mitochondria through its C-terminal mitochondria-targeting domain and axon-sorting sequence. Syntaphilin immobilizes axonal mitochondria by anchoring to MTs. Deleting syntaphilin results in a robust increase of axonal mitochondria in motile pools. Conversely, overexpressing syntaphilin abolishes axonal mitochondrial transport. These findings provide new mechanistic insight that motile mitochondria can be recruited into stationary pools via anchoring interactions between mitochondria-targeted syntaphilin and MTs. Thus, syntaphilin serves as an attractive molecular target for investigations into mechanisms recruiting motile mitochondria into activated synapses. The syntaphilin knockout mouse is an ideal genetic model to examine the impact of enhanced motility of axonal mitochondria on presynaptic function and mitochondrial quality control in distal axons.

Synaptic activity–dependent regulation

Mitochondrial transport and distribution in axons and at synapses are correlated with synaptic activity. Mitochondria are recruited to synapses in response to elevated intracellular Ca²⁺ during sustained synaptic activity. Elevated Ca²⁺ influx, either by activating voltage-dependent calcium channels or NMDA receptors, arrests mitochondrial movement (Rintoul et al., 2003; Yi et al., 2004; Chang et al., 2006; Szabadkai et al., 2006; Ohno et al., 2011). These studies raise a fundamental question: how are mitochondria recruited to synapses in response to Ca²⁺ influx? Recent work from three groups identified Miro as a Ca²⁺ sensor in regulating mitochondrial motility (Saotome et al., 2008; MacAskill et al., 2009a; Wang and Schwarz, 2009). Miro has four Ca²⁺-binding EF-hands (Klosowiak et al., 2013). Elevated cytosolic Ca²⁺ levels, by firing action potentials or activating glutamate receptors, arrest mitochondrial transport through a Miro-Ca²⁺–sensing pathway that inactivates or disassembles the KIF5–Miro–Trak transport machineries. By this mechanism, mitochondria are recruited to activated synapses. A Miro-Ca²⁺–sensing model was proposed in which KIF5 is loaded onto mitochondria through physical coupling via the motor adaptor complex Miro–Milton (or Miro–Trak) at resting Ca²⁺ levels. When motile mitochondria pass by an active synaptic terminal, the local elevated intracellular Ca²⁺ disrupts or inactivates the Miro–Trak–kinesin transport complex by binding to the EF-hand and changing Miro’s conformation; thus, mitochondria are immobilized at activated synapses (Fig. 2, C and D; MacAskill et al., 2009a; Wang and Schwarz, 2009). The two studies disagreed, however, as to whether kinesin remained associated with stationary mitochondria or whether the motor was completely decoupled from the organelle upon immobilization. In addition, Miro-Ca²⁺–dependent regulation affects both anterograde and retrograde transport. When KIF5 transport machinery is disrupted by the Miro-Ca²⁺–sensing mechanism, dynein-driven retrograde transport does not necessarily take over. It is unclear how the Miro-Ca²⁺–sensing pathway inactivates dynein transport machineries. Thus, the role of the Miro-Ca²⁺–sensing pathway in immobilizing mitochondria likely occurs through an anchoring mechanism.A recent study provides evidence to support the above-described hypothesis. Activating the Miro-Ca²⁺ pathway fails to arrest axonal mitochondria in syntaphilin-null hippocampal neurons: axonal mitochondria keep moving during neuronal firing while dendritic mitochondria are effectively immobilized (Chen and Sheng, 2013). This study suggests that syntaphilin-mediated anchoring plays a central role in the activity-dependent immobilization of axonal mitochondria. Syntaphilin competes with Trak2 for binding KIF5 in cells and inhibits the KIF5 motor ATPase activity in vitro, thus suggesting that syntaphilin functions to prevent KIF5 from moving along MTs. Synaptic activity and elevated calcium levels favor the anchoring interaction between syntaphilin and KIF5 by disrupting the Miro–Trak–kinesin transport complex. In addition, sustained neuronal activity induces syntaphilin translocation to axonal mitochondria, although the underlying mechanisms remain elusive. Expressing the syntaphilin mutant lacking this axon-sorting domain results in its distribution into all mitochondria, including those in the soma and dendrites (Kang et al., 2008). These studies collectively suggest a possibility that syntaphilin is delivered to axons before its translocation into mitochondria. One interesting question is how syntaphilin-anchored mitochondria are remobilized. Given the fact that syntaphilin is a cellular target of the E3 ubiquitin ligase Cullin 1 (Yen and Elledge, 2008), syntaphilin may be degraded by ubiquitin-mediated pathways. It is necessary to further investigate the mechanisms underlying activity-induced syntaphilin translocation and its subsequent removal from mitochondria.A new “engine-switch and brake” model (Fig. 2 E) was recently proposed (Chen and Sheng 2013). In response to a “stop” sign (elevated Ca²⁺) at active synapses, syntaphilin switches off the kinesin motor and puts a brake on mitochondria, thereby anchoring them in place on MTs. Thus, KIF5-loaded mitochondria remain associated with the axonal MT track while in their stationary phase. This model may help reconcile the standing disagreements regarding how Miro-Ca²⁺ sensing immobilizes mitochondria in neurons. MacAskill et al. (2009a) propose that Ca²⁺ binding to Miro disconnects KIF5 motors from mitochondria in dendrites by activating glutamate receptors (Fig. 2 D), whereas Wang and Schwarz (2009) suggest that KIF5 remains associated with immobilized mitochondria within axons during elevated Ca²⁺ conditions (Fig. 2 C). These discrepant results are likely due to their selective observations of mitochondrial motility in dendrites versus axons. Upon Miro-Ca²⁺ sensing, KIF5 motors remain associated with axonal mitochondria by binding syntaphilin, which is absent from dendritic mitochondria (Fig. 2 E). This difference may help axonal mitochondria be more responsive to changes in synaptic activity. If the Ca²⁺ signal is removed, the cargo-loaded motor proteins can be quickly reactivated to move mitochondria to new active synapses. This model also suggests that motor loading on mitochondria is insufficient for transport. The release of the anchoring interaction is also required. The latter is supported by a study that both KIF5 and dynein motors remain bound on prion protein vesicles regardless of whether they are motile or stationary (Encalada et al., 2011). A newly reported Alex3 protein, organized in a genomic Armcx cluster encoding mitochondria-targeted proteins, regulates mitochondrial trafficking in neurons by interacting with the KIF5–Miro–Trak complex in a Ca²⁺-dependent manner (López-Doménech et al., 2012). Although the underlying mechanism is unknown, the study highlights the complexity in controlling mitochondrial motility in response to various physiological signals in neurons.

Mitochondrial motility and synaptic variability

Mitochondria are commonly found within synaptic terminals (Shepherd and Harris, 1998), where they power neurotransmission by supplying ATP. A stationary mitochondrion within presynaptic terminals provides a stable and continuous ATP supply and maintains energy homeostasis at synapses. Conversely, a motile mitochondrion passing by presynaptic boutons dynamically alters local ATP levels, thus influencing various ATP-dependent functions at synapses such as: (1) establishing the proton gradient necessary for neurotransmitter loading; (2) removing Ca²⁺ from nerve terminals; and (3) driving synaptic vesicle transport from reserve pools to release sites. For example, Drosophila photoreceptors expressing mutant Milton display impaired synaptic transmission due to reduced mitochondrial trafficking to synapses (Stowers et al., 2002). Mutation of Drosophila Drp1 reduces synaptic localization of mitochondria and results in faster depletion of synaptic vesicles during prolonged stimulation (Verstreken et al., 2005). Addition of ATP to terminals partially rescues these defects, indicating that mitochondrial ATP production is required for maintaining sustained synaptic transmission. Syntabulin loss of function reduces mitochondrial density in axonal terminals, which is accompanied by accelerated synaptic depression and slowed synapse recovery during high-frequency firing (Ma et al., 2009).One of the most notable characteristics of synaptic transmission is the wide variation in synaptic strength in response to repeated stimulations. The effects of synaptic variability on neuronal circuit activity are increasingly recognized. Some degree of synaptic variability may be necessary for signal processing in flexible or adaptive systems (Murthy et al., 1997). A long-standing fundamental question is how the variation in synaptic strength arises. There is general agreement that some structural and molecular stochastic alternations at synapses are the basis for heterogeneity in synaptic transmission from neuron to neuron or from synapse to synapse (Atwood and Karunanithi, 2002; Stein et al., 2005; Marder and Goaillard, 2006; Branco and Staras, 2009; Ribrault et al., 2011). However, it is unknown whether mitochondrial motility at axonal terminals contributes to pulse-to-pulse variability of presynaptic strength at single-bouton levels. This is particularly relevant in hippocampal neurons, where approximately one third of axonal mitochondria are highly motile, some of which dynamically pass through presynaptic boutons.We recently solved this puzzle by combining dual-channel imaging in live neurons and electrophysiological analysis in syntaphilin knockout hippocampal neurons and acute hippocampal slices in which axonal mitochondrial motility is selectively manipulated (Sun et al., 2013). Our study revealed that the motility of axonal mitochondria is one of the primary mechanisms underlying the variability of presynaptic strength. Enhanced motility of axonal mitochondria increases the pulse-to-pulse variability of EPSC amplitudes, whereas immobilizing axonal mitochondria effectively diminishes the variability. The data raise two assumptions: (1) EPSC amplitudes are averaged through summation from multiple synapses; and (2) release at individual boutons changes over time when mitochondrial distribution and motility are dynamically altered. Thus, at any given time these boutons display different patterns of mitochondrial distribution and motility; mitochondria move either in or out of boutons or pass through boutons during stimulation, thus contributing to the pulse-to-pulse variability. By applying dual-channel live imaging of mitochondria and synaptic vesicle release at single-bouton levels, the study further shows that mitochondrial movement, either into or out of presynaptic boutons, significantly influences synaptic vesicle release due to fluctuation of synaptic ATP levels. In the absence of a stationary mitochondrion within an axonal terminal, there is no constant on-site ATP supply. A motile mitochondrion passing through could temporally supply ATP, thus changing synaptic energy levels and influencing ATP-dependent processes including synaptic vesicle cycling, mobilization, and replenishment (Fig. 3). This is further supported by a recent study applying a new quantitative optical presynaptic ATP reporter. Synaptic activity drives large ATP consumption at nerve terminals and synaptic vesicle cycling consumes most presynaptic ATP (Rangaraju et al., 2014). It is possible that all of these ATP-dependent processes collectively contribute to presynaptic variability. Therefore, the fluctuation of synaptic ATP levels resulting from mitochondrial movement is one of the primary sources for the wide variability of EPSC amplitudes. This study revealed, for the first time, that the dynamic movement of axonal mitochondria is one of the primary mechanisms underlying the presynaptic variation in the CNS, thus providing new insight into the fundamental properties of the CNS to ensure the plasticity and reliability of synaptic transmission (Sun et al., 2013).Open in a separate windowFigure 3.Motile mitochondria passing by synapses contribute to presynaptic variation. A stationary mitochondrion within presynaptic terminals powers neurotransmission by stable and continuous ATP supply (left). Conversely, in the absence of a mitochondrion within a presynaptic bouton (right), there is no stable on-site ATP supply; a motile mitochondrion passing through this bouton temporally supplies ATP, thus changing synaptic energy levels and influencing ATP-dependent synaptic functions over time when mitochondrial distribution and motility are altered. Therefore, the fluctuation of synaptic ATP levels resulting from mitochondrial movement is one of the primary sources for the wide variability of synaptic vesicle release and amplitudes of excitatory postsynaptic currents (EPSCs; Sun et al., 2013).

Mitochondrial motility, neuronal morphology, and energy consumption

Proper mitochondrial transport into neurites in developing neurons is tightly regulated to ensure that metabolically active areas are adequately supplied with ATP (Morris and Hollenbeck, 1993). A recent study showed the differential functions of two mitochondrial motor adaptors, Trak1 and Trak2, in driving polarized mitochondrial transport (van Spronsen et al., 2013). Trak1 is prominently localized in axons and is required for axonal mitochondria transport by binding to both kinesin-1 and dynein/dynactin, whereas Trak2 is present in dendrites and mainly transports mitochondria into dendrites by interacting with dynein/dynactin. The folded configuration of Trak2 preferentially binds dynein motors rather than kinesin-1. Consistently, depleting Trak1 inhibits axonal outgrowth, whereas suppressing Trak2 impairs dendrite morphology. Thus, this study established a new role of Trak proteins in polarized mitochondrial targeting necessary to maintain neuronal morphology.Anchored stationary mitochondria ideally serve as local energy power plants. Increased intracellular [ADP]_i slows down mitochondrial movement and recruits mitochondria to subcellular regions correlated with local [ATP]_i depletion (Mironov, 2007). However, the underlying mechanisms coordinating mitochondria transport and energy consumption remain elusive. The balance between motile and stationary mitochondria responds quickly to changes in axonal growth status. Two recent studies demonstrated that generating and maintaining axonal branching depend upon stable and continuous ATP supply by stationary mitochondria captured at branching points. Courchet et al. (2013) revealed that LKB1, the serine/threonine liver kinase B1, regulates terminal axon branching of cortical neurons in both in vitro and in vivo systems through activating downstream kinase NUAK1, an AMPK-like kinase. AMPK is an AMP-activated protein kinase, which is a master regulator of cellular energy homeostasis and is activated in response to stresses that deplete cellular ATP supplies. Deleting LKB1 or NUAK1 results in a threefold decrease in stationary pools of axonal mitochondria, whereas overexpressing LKB1 or NUAK1 conversely increases the proportion of immobilized mitochondria along axons, thus establishing a causal correlation between stationary mitochondria and axonal branching. To test the hypothesis, Courchet et al. (2013) altered the motility of axonal mitochondria by changing expression levels of syntaphilin. Intriguingly, suppressing syntaphilin expression in cortical progenitors at E15.5 enhances axonal mitochondrial motility accompanied by decreased axon branching. Conversely, overexpressing syntaphilin immobilizes axonal mitochondria and increases axon branching, thus highlighting a critical role for syntaphilin-mediated mitochondrial anchoring in AMPK-induced axonal branching. It is possible that LKB1 and NUAK1-mediated signaling pathways regulate axon branch formation and stabilization by recruiting mitochondria to branch points. However, this study raises one mechanistic question: does syntaphilin or the mitochondrial transport motor–adaptor complex serve as a downstream effector of this signaling pathway? Syntaphilin is strictly developmentally regulated in mouse brains, with low expression before postnatal day 7 and peaking two weeks after birth; thus, its expression is hardly detectable in early embryonic stages (Kang et al., 2008). It remains unclear how siRNA application at E15.5 significantly impacts endogenous syntaphilin levels in the brain. Furthermore, it is unknown whether local ATP homeostasis maintained by stationary but not motile mitochondria contributes to the formation and stabilization of axonal branching.As a cellular energy sensor for detecting increased intracellular [AMP]_i or ATP consumption, AMPK activation positively regulates signaling pathways that replenish cellular ATP supplies. Tao et al. (2013) provide further evidence that sustained neuronal depolarization for 20 min depletes cellular ATP and activates AMPK. Activation of AMPK increases anterograde flux of mitochondria into axons and induces axonal branching in regions where mitochondria are docked in an ATP-dependent manner. The mitochondrial uncoupler FCCP blocks axonal branching, even when mitochondria are localized nearby. These studies provide evidence for how the AMPK signaling pathway regulates axon branch formation and stabilization. These studies collectively highlight a critical role for syntaphilin-mediated mitochondrial anchoring in axon branching. One possible mechanism underlying mitochondrial recruiting and anchoring in support of axon branching is through localized intra-axonal protein synthesis (Spillane et al., 2013). Inhibiting mitochondrial respiration and protein synthesis in axons impairs axonal branching. Thus, stationary mitochondria coordinate maturation of axonal filopodia into axon branching likely by supporting active mRNA translation in axonal hot spots. Activity-induced mitochondrial recruitment in axons is further supported by an in vivo study in mouse saphenous nerve axons. Sustained electrical stimulation (50 Hz) enhances anterograde flux of mitochondria into myelinated axons and redistributes them at the peripheral terminals (Sajic et al., 2013).

Mitochondrial motility and mitophagy

Cumulative evidence indicates mitochondrial dysfunction, impaired transport, altered dynamics, and perturbation of mitochondrial turnover are associated with the pathology of major neurodegenerative disorders (Chen and Chan, 2009; Sheng and Cai, 2012). Defective mitochondrial transport accompanied by mitochondrial pathology is one of the most notable pathological changes in several aging-associated neurodegenerative diseases, including Alzheimer’s disease (Rui et al., 2006; Du et al., 2010; Calkins et al., 2011) and amyotrophic lateral sclerosis (De Vos et al., 2007; Magrané and Manfredi, 2009; Bilsland et al., 2010; Shi et al., 2010; Martin, 2011; Zhu and Sheng, 2011; Marinkovic et al., 2012). Dysfunctional mitochondria not only supply less ATP and maintain Ca²⁺ buffering capacity less efficiently, but also release harmful reactive oxygen species (ROS). Defects in mitochondrial transport reduce the delivery of healthy mitochondria to distal processes and also impair removal of damaged mitochondria from synapses, thus causing energy depletion and altered Ca²⁺ transient at synapses. In addition, toxic ROS may further trigger synaptic stress, thereby contributing to neurodegeneration. Mitochondrial quality control involves multiple levels of surveillance to ensure mitochondrial integrity, including repairing dysfunctional mitochondria by fusion with healthy ones (Chen and Chan 2009; Westermann, 2010) and segregating irreversibly damaged mitochondria for mitophagy, a cargo-specific autophagy to remove damaged mitochondria. Effective sequestration of damaged mitochondria into autophagosomes and subsequent clearance within the autophagy–lysosomal system constitute a key cellular pathway in mitochondrial quality control mechanisms. Recent studies indicate that PINK1/Parkin-mediated pathways ensure mitochondrial integrity and function (Clark et al., 2006; Gautier et al., 2008; Narendra et al., 2008). Loss-of-function mutations in PINK1 and Parkin are associated with recessive forms of Parkinson’s disease. When mitochondria are depolarized or their integrity is damaged, PINK1 accumulates on the outer mitochondrial membrane and recruits the E3 ubiquitin ligase Parkin from the cytosol. Parkin ubiquitinates mitochondrial proteins (Geisler et al., 2010; Poole et al., 2010; Ziviani et al., 2010; Yoshii et al., 2011). This process is essential for damaged mitochondria to be engulfed by isolation membranes that then fuse with lysosomes for degradation.Investigating those mechanisms coordinating mitochondrial motility and mitophagy represents an important emerging area for disease-oriented research. Mature acidic lysosomes are mainly located in the soma (Overly and Hollenbeck, 1996; Cai et al., 2010; Lee et al., 2011). Thus, a long-standing question remains: how are aged and damaged mitochondria at distal terminals efficiently eliminated via the autophagy–lysosomal pathway? Cai et al. (2012) recently reported the unique features of spatial and dynamic Parkin-mediated mitophagy in mature cortical neurons, in which mitochondria remain motile after prolonged and chronic dissipation of the mitochondrial membrane potential (Δψ_m) for 24 h. Given the fact that in vitro mitochondrial depolarization will quickly trigger neuron apoptosis, a high-quality culturing system is essential so that the neurons survive long enough to exhibit altered mitochondrial transport and Parkin-mediated mitophagy. Chronically dissipating Δψ_m by prolonged treatment of low concentrations of Δψ_m-uncoupling reagent CCCP (10 µM) results in accumulation of Parkin-targeted mitochondria in the soma and proximal regions. Such compartmental restriction for Parkin-targeted mitochondria is consistently observed in neurons treated with low concentrations of another Δψ_m-dissipating reagent, antimycin A (1 µM), or mitochondrial ATP synthase inhibitor oligomycin (1 µM). This distribution pattern is attributable to altered motility of chronically depolarized mitochondria with reduced anterograde and relatively enhanced retrograde transport, thus reducing anterograde flux of damaged mitochondria into distal processes. Intriguingly, when anchored by syntaphilin before the dissipation of Δψ_m, mitochondria in distal axons also recruit Parkin for mitophagy. Altered motility may be protective under chronic mitochondrial stress conditions; healthy mitochondria remain distally while aged and damaged ones return to the soma for degradation (Fig. 4). This spatial distribution allows neurons to efficiently remove dysfunctional mitochondria distally and then eliminate them via the autophagy–lysosomal pathway in the soma.Open in a separate windowFigure 4.Functional interplay between mitochondrial transport and mitophagy. Chronically dissipating mitochondrial membrane potential (Δψ_m) by prolonged treatment of low concentrations of Δψ_m-uncoupling reagents accumulates Parkin-targeted mitochondria in the soma and proximal regions. Such compartmental restriction is a result of altered motility of depolarized mitochondria with reduced anterograde and relatively enhanced retrograde transport, thus reducing anterograde flux of damaged mitochondria into distal processes (Cai et al., 2012). This spatial process allows neurons to efficiently remove dysfunctional mitochondria from distal axons via the autophagy–lysosomal pathway in the soma, where mature lysosomes are relatively enriched. Damaged mitochondria at axonal terminals can also recruit Parkin for mitophagy once they are anchored by syntaphilin (Cai et al., 2012) or immobilized by turnover of the motor adaptor Miro on the mitochondrial surface (Weihofen et al., 2009; Chan et al., 2011; Wang et al., 2011; Yoshii et al., 2011; Liu et al., 2012; Sarraf et al., 2013). Autophagosomes including those engulfing damaged mitochondria at axonal terminals transport predominantly to the soma for maturation and more efficient degradation of cargoes within acidic lysosomes (Maday et al., 2012).Reduced anterograde transport of depolarized mitochondria is consistent with recent studies showing Parkin-mediated degradation of Miro on the mitochondrial surface upon Δψ_m dissipation–induced mitophagy. In addition to binding to the KIF5–Trak motor complex, the mitochondrial outer membrane protein Miro also interacts with PINK1 and Parkin and is ubiquitinated by Parkin while mitochondria are depolarized (Weihofen et al., 2009; Chan et al., 2011; Wang et al., 2011; Liu et al., 2012; Sarraf et al., 2013). These studies support the current model that altered mitochondrial transport is a critical aspect of the mechanisms by which the PINK1/Parkin pathway governs mitochondrial quality control. Turnover of Miro on the mitochondrial surface may favor their retrograde transport to the soma or immobilize damaged organelles at distal regions for mitophagy (Fig. 4). A recent study reports the predominant retrograde transport of autophagosomes including those engulfing damaged mitochondria from distal axons to the soma (Maday et al., 2012). The retrograde movement is critical for autophagosome maturation and degradation within acidic lysosomes in the proximal regions of neurons. This study highlights the possibility that distal damaged mitochondria, which are either anchored by syntaphilin or immobilized by turning over Miro, may be also degraded in the soma after their retrograde transport in engulfed autophagosomes. Thus, a functional interplay is proposed between mitochondrial motility and mitophagy to ensure proper removal of aged and dysfunctional mitochondria from distal processes. The correlation between mitochondrial Δψ_m and motility is controversial, as demonstrated in two previous studies by acutely treating neurons with high concentrations of Δψ_m-dissipating reagents (Miller and Sheetz, 2004; Verburg and Hollenbeck, 2008). Instead, Cai et al. (2012) examined mitochondrial motility and mitophagy in mature cortical neurons after a 24-h treatment with a much lower concentration of CCCP (10 µM) or antimycin A (1 µM) combined with applying the pan-caspase inhibitor Z-VAD. The majority of neurons survive and mitochondria remain highly motile under such prolonged stress conditions, thus allowing detection of the altered transport event. Therefore, prolonged Δψ_m dissipation in cultured neurons better reflects chronic mitochondria stress under in vivo physiological or pathological conditions.

Moving forward

The recent discoveries of proteins involved in mitochondrial transport and anchoring boosted our understanding of the molecular mechanisms that regulate mitochondrial motility and distribution in response to physiological and pathological signals. However, there are still many mechanistic questions to be addressed in the near future. In particular, how does a Miro-Ca²⁺–sensing pathway arrest both anterograde and retrograde movement of mitochondria at active synapses? How do dynein and KIF5 motors physically or mechanistically interplay to coordinate mitochondrial bi-directional transport? How are stationary mitochondria remobilized, recruited, and redistributed (and vice versa) to sense, integrate, and respond to changes in mitochondrial membrane potential, cellular metabolic status, neuronal growth signals, and pathological stress? Investigating how mitochondrial motility coordinates their function and integrity in neurons represents an important emerging frontier in neurobiology. Mitochondria undergo dynamic membrane fission/fusion and mitophagy to maintain their quality during the neuronal lifetime. Mitochondrial pathology is one of the most notable hallmarks in several major aging-associated neurodegenerative diseases (Chen and Chan, 2009; Schon and Przedborski, 2011; Court and Coleman, 2012; Sheng and Cai, 2012; Itoh et al., 2013). The proper and efficient elimination of damaged mitochondria via mitophagy may serve as an early neuroprotective mechanism. Studying these dynamic processes in live neurons directly isolated from disease-stage adult mice will provide more reliable models for delineating defects in mitochondrial transport and mitophagy underlying adult-onset pathogenesis. Mechanistic insight into these fundamental processes in coordinating mitochondrial trafficking and anchoring, membrane dynamics, and mitophagy will advance our understanding of human neurodegenerative diseases.     

Focus Issue on Calcium Signaling: Identification of Cyclic GMP-Activated Nonselective Ca2+-Permeable Cation Channels and Associated CNGC5 and CNGC6 Genes in Arabidopsis Guard Cells

Cytosolic Ca²⁺ in guard cells plays an important role in stomatal movement responses to environmental stimuli. These cytosolic Ca²⁺ increases result from Ca²⁺ influx through Ca²⁺-permeable channels in the plasma membrane and Ca²⁺ release from intracellular organelles in guard cells. However, the genes encoding defined plasma membrane Ca²⁺-permeable channel activity remain unknown in guard cells and, with some exceptions, largely unknown in higher plant cells. Here, we report the identification of two Arabidopsis (Arabidopsis thaliana) cation channel genes, CNGC5 and CNGC6, that are highly expressed in guard cells. Cytosolic application of cyclic GMP (cGMP) and extracellularly applied membrane-permeable 8-Bromoguanosine 3′,5′-cyclic monophosphate-cGMP both activated hyperpolarization-induced inward-conducting currents in wild-type guard cells using Mg²⁺ as the main charge carrier. The cGMP-activated currents were strongly blocked by lanthanum and gadolinium and also conducted Ba²⁺, Ca²⁺, and Na⁺ ions. cngc5 cngc6 double mutant guard cells exhibited dramatically impaired cGMP-activated currents. In contrast, mutations in CNGC1, CNGC2, and CNGC20 did not disrupt these cGMP-activated currents. The yellow fluorescent protein-CNGC5 and yellow fluorescent protein-CNGC6 proteins localize in the cell periphery. Cyclic AMP activated modest inward currents in both wild-type and cngc5cngc6 mutant guard cells. Moreover, cngc5 cngc6 double mutant guard cells exhibited functional abscisic acid (ABA)-activated hyperpolarization-dependent Ca²⁺-permeable cation channel currents, intact ABA-induced stomatal closing responses, and whole-plant stomatal conductance responses to darkness and changes in CO₂ concentration. Furthermore, cGMP-activated currents remained intact in the growth controlled by abscisic acid2 and abscisic acid insensitive1 mutants. This research demonstrates that the CNGC5 and CNGC6 genes encode unique cGMP-activated nonselective Ca²⁺-permeable cation channels in the plasma membrane of Arabidopsis guard cells.Plants lose water via transpiration and take in CO₂ for photosynthesis through stomatal pores. Each stomatal pore is surrounded by two guard cells, and stomatal movements are driven by the change of turgor pressure in guard cells. The intracellular second messenger Ca²⁺ functions in guard cell signal transduction (Schroeder and Hagiwara, 1989; McAinsh et al., 1990; Webb et al., 1996; Grabov and Blatt, 1998; Allen et al., 1999; MacRobbie, 2000; Mori et al., 2006; Young et al., 2006; Siegel et al., 2009; Chen et al., 2010; Hubbard et al., 2012). Plasma membrane ion channel activity and gene expression in guard cells are finely regulated by the intracellular free calcium concentration ([Ca2+]_cyt; Schroeder and Hagiwara, 1989; Webb et al., 2001; Allen et al., 2002; Siegel et al., 2009; Kim et al., 2010; Stange et al., 2010). Ca²⁺-dependent protein kinases (CPKs) function as targets of the cytosolic Ca²⁺ signal, and several members of the CPK family have been shown to function in stimulus-induced stomatal closing, including the Arabidopsis (Arabidopsis thaliana) CPK3, CPK4, CPK6, CPK10, and CPK11 proteins (Mori et al., 2006; Zhu et al., 2007; Zou et al., 2010; Brandt et al., 2012; Hubbard et al., 2012). Further research found that several CPKs could activate the S-type anion channel SLAC1 in Xenopus laevis oocytes, including CPK21, CPK23, and CPK6 (Geiger et al., 2010; Brandt et al., 2012). At the same time, the Ca²⁺-independent protein kinase Open Stomata1 mediates stomatal closing and activates the S-type anion channel SLAC1 (Mustilli et al., 2002; Yoshida et al., 2002; Geiger et al., 2009; Lee et al., 2009; Xue et al., 2011), indicating that both Ca²⁺-dependent and Ca²⁺-independent pathways function in guard cells.Multiple essential factors of guard cell abscisic acid (ABA) signal transduction function in the regulation of Ca²⁺-permeable channels and [Ca2+]_cyt elevations, including Abscisic Acid Insensitive1 (ABI1), ABI2, Enhanced Response to Abscisic Acid1 (ERA1), the NADPH oxidases AtrbohD and AtrbohF, the Guard Cell Hydrogen Peroxide-Resistant1 (GHR1) receptor kinase, as well as the Ca²⁺-activated CPK6 protein kinase (Pei et al., 1998; Allen et al., 1999, 2002; Kwak et al., 2003; Miao et al., 2006; Mori et al., 2006; Hua et al., 2012). [Ca2+]_cyt increases result from both Ca²⁺ release from intracellular Ca²⁺ stores (McAinsh et al., 1992) and Ca²⁺ influx across the plasma membrane (Hamilton et al., 2000; Pei et al., 2000; Murata et al., 2001; Kwak et al., 2003; Hua et al., 2012). Electrophysiological analyses have characterized nonselective Ca²⁺-permeable channel activity in the plasma membrane of guard cells (Schroeder and Hagiwara, 1990; Hamilton et al., 2000; Pei et al., 2000; Murata et al., 2001; Köhler and Blatt, 2002; Miao et al., 2006; Mori et al., 2006; Suh et al., 2007; Vahisalu et al., 2008; Hua et al., 2012). However, the genetic identities of Ca²⁺-permeable channels in the plasma membrane of guard cells have remained unknown despite over two decades of research on these channel activities.The Arabidopsis genome includes 20 genes encoding cyclic nucleotide-gated channel (CNGC) homologs and 20 genes encoding homologs to animal Glu receptor channels (Lacombe et al., 2001; Kaplan et al., 2007; Ward et al., 2009), which have been proposed to function in plant cells as cation channels (Schuurink et al., 1998; Arazi et al., 1999; Köhler et al., 1999). Recent research has demonstrated functions of specific Glu receptor channels in mediating Ca²⁺ channel activity (Michard et al., 2011; Vincill et al., 2012). Previous studies have shown cAMP activation of nonselective cation currents in guard cells (Lemtiri-Chlieh and Berkowitz, 2004; Ali et al., 2007). However, only a few studies have shown the disappearance of a defined plasma membrane Ca²⁺ channel activity in plants upon mutation of candidate Ca²⁺ channel genes (Ali et al., 2007; Michard et al., 2011; Laohavisit et al., 2012; Vincill et al., 2012). Some CNGCs have been found to be involved in cation nutrient intake, including monovalent cation intake (Guo et al., 2010; Caballero et al., 2012), salt tolerance (Guo et al., 2008; Kugler et al., 2009), programmed cell death and pathogen responses (Clough et al., 2000; Balagué et al., 2003; Urquhart et al., 2007; Abdel-Hamid et al., 2013), thermal sensing (Finka et al., 2012; Gao et al., 2012), and pollen tube growth (Chang et al., 2007; Frietsch et al., 2007; Tunc-Ozdemir et al., 2013a, 2013b). Direct in vivo disappearance of Ca²⁺ channel activity in cngc disruption mutants has been demonstrated in only a few cases thus far (Ali et al., 2007; Gao et al., 2012). In this research, we show that CNGC5 and CNGC6 are required for a cyclic GMP (cGMP)-activated nonselective Ca²⁺-permeable cation channel activity in the plasma membrane of Arabidopsis guard cells.     

Leaf Senescence Signaling: The Ca2+-Conducting Arabidopsis Cyclic Nucleotide Gated Channel2 Acts through Nitric Oxide to Repress Senescence Programming

Ca²⁺ and nitric oxide (NO) are essential components involved in plant senescence signaling cascades. In other signaling pathways, NO generation can be dependent on cytosolic Ca²⁺. The Arabidopsis (Arabidopsis thaliana) mutant dnd1 lacks a plasma membrane-localized cation channel (CNGC2). We recently demonstrated that this channel affects plant response to pathogens through a signaling cascade involving Ca²⁺ modulation of NO generation; the pathogen response phenotype of dnd1 can be complemented by application of a NO donor. At present, the interrelationship between Ca²⁺ and NO generation in plant cells during leaf senescence remains unclear. Here, we use dnd1 plants to present genetic evidence consistent with the hypothesis that Ca²⁺ uptake and NO production play pivotal roles in plant leaf senescence. Leaf Ca²⁺ accumulation is reduced in dnd1 leaves compared to the wild type. Early senescence-associated phenotypes (such as loss of chlorophyll, expression level of senescence-associated genes, H₂O₂ generation, lipid peroxidation, tissue necrosis, and increased salicylic acid levels) were more prominent in dnd1 leaves compared to the wild type. Application of a Ca²⁺ channel blocker hastened senescence of detached wild-type leaves maintained in the dark, increasing the rate of chlorophyll loss, expression of a senescence-associated gene, and lipid peroxidation. Pharmacological manipulation of Ca²⁺ signaling provides evidence consistent with genetic studies of the relationship between Ca²⁺ signaling and senescence with the dnd1 mutant. Basal levels of NO in dnd1 leaf tissue were lower than that in leaves of wild-type plants. Application of a NO donor effectively rescues many dnd1 senescence-related phenotypes. Our work demonstrates that the CNGC2 channel is involved in Ca²⁺ uptake during plant development beyond its role in pathogen defense response signaling. Work presented here suggests that this function of CNGC2 may impact downstream basal NO production in addition to its role (also linked to NO signaling) in pathogen defense responses and that this NO generation acts as a negative regulator during plant leaf senescence signaling.Senescence can be considered as the final stage of a plant’s development. During this process, nutrients will be reallocated from older to younger parts of the plant, such as developing leaves and seeds. Leaf senescence has been characterized as a type of programmed cell death (PCD; Gan and Amasino, 1997; Quirino et al., 2000; Lim et al., 2003). During senescence, organelles such as chloroplasts will break down first. Biochemical changes will also occur in the peroxisome during this process. When the chloroplast disassembles, it is easily observed as a loss of chlorophyll. Mitochondria, the source of energy for cells, will be the last cell organelles to undergo changes during the senescence process (Quirino et al., 2000). At the same time, other catabolic events (e.g. protein and lipid breakdown, etc.) are occurring (Quirino et al., 2000). Hormones may also contribute to this process (Gepstein, 2004). From this information we can infer that leaf senescence is regulated by many signals.Darkness treatment can induce senescence in detached leaves (Poovaiah and Leopold, 1973; Chou and Kao, 1992; Weaver and Amasino, 2001; Chrost et al., 2004; Guo and Crawford, 2005; Ülker et al., 2007). Ca²⁺ can delay the senescence of detached leaves (Poovaiah and Leopold, 1973) and leaf senescence induced by methyl jasmonate (Chou and Kao, 1992); the molecular events that mediate this effect of Ca²⁺ are not well characterized at present.Nitric oxide (NO) is a critical signaling molecule involved in many plant physiological processes. Recently, published evidence supports NO acting as a negative regulator during leaf senescence (Guo and Crawford, 2005; Mishina et al., 2007). Abolishing NO generation in either loss-of-function mutants (Guo and Crawford, 2005) or transgenic Arabidopsis (Arabidopsis thaliana) plants expressing NO degrading dioxygenase (NOD; Mishina et al., 2007) leads to an early senescence phenotype in these plants compared to the wild type. Corpas et al. (2004) showed that endogenous NO is mainly accumulated in vascular tissues of pea (Pisum sativum) leaves. This accumulation is significantly reduced in senescing leaves (Corpas et al., 2004). Corpas et al. (2004) also provided evidence that NO synthase (NOS)-like activity (i.e. generation of NO from l-Arg) is greatly reduced in senescing leaves. Plant NOS activity is regulated by Ca²⁺/calmodulin (CaM; Delledonne et al., 1998; Corpas et al., 2004, 2009; del Río et al., 2004; Valderrama et al., 2007; Ma et al., 2008). These studies suggest a link between Ca²⁺ and NO that could be operating during senescence.In animal cells, all three NOS isoforms require Ca²⁺/CaM as a cofactor (Nathan and Xie, 1994; Stuehr, 1999; Alderton et al., 2001). Notably, animal NOS contains a CaM binding domain (Stuehr, 1999). It is unclear whether Ca²⁺/CaM can directly modulate plant NOS or if Ca²⁺/CaM impacts plant leaf development/senescence through (either direct or indirect) effects on NO generation. However, recent studies from our lab suggest that Ca²⁺/CaM acts as an activator of NOS activity in plant innate immune response signaling (Ali et al., 2007; Ma et al., 2008).Although Arabidopsis NO ASSOCIATED PROTEIN1 (AtNOA1; formerly named AtNOS1) was thought to encode a NOS enzyme, no NOS-encoding gene has yet been identified in plants (Guo et al., 2003; Crawford et al., 2006; Zemojtel et al., 2006). However, the AtNOA1 loss-of-function mutant does display reduced levels of NO generation, and several groups have used the NO donor sodium nitroprusside (SNP) to reverse some low-NO related phenotypes in Atnoa1 plants (Guo et al., 2003; Bright et al., 2006; Zhao et al., 2007). Importantly, plant endogenous NO deficiency (Guo and Crawford, 2005; Mishina et al., 2007) or abscisic acid/methyl jasmonate (Hung and Kao, 2003, 2004) induced early senescence can be successfully rescued by application of exogenous NO. Addition of NO donor can delay GA-elicited PCD in barley (Hordeum vulgare) aleurone layers as well (Beligni et al., 2002).It has been suggested that salicylic acid (SA), a critical pathogen defense metabolite, can be increased in natural (Morris et al., 2000; Mishina et al., 2007) and transgenic NOD-induced senescent Arabidopsis leaves (Mishina et al., 2007). Pathogenesis related gene1 (PR1) expression is up-regulated in transgenic Arabidopsis expressing NOD (Mishina et al., 2007) and in leaves of an early senescence mutant (Ülker et al., 2007).Plant cyclic nucleotide gated channels (CNGCs) have been proposed as candidates to conduct extracellular Ca²⁺ into the cytosol (Sunkar et al., 2000; Talke et al., 2003; Lemtiri-Chlieh and Berkowitz, 2004; Ali et al., 2007; Demidchik and Maathuis, 2007; Frietsch et al., 2007; Kaplan et al., 2007; Ma and Berkowitz, 2007; Urquhart et al., 2007; Ma et al., 2009a, 2009b). Arabidopsis “defense, no death” (dnd1) mutant plants have a null mutation in the gene encoding the plasma membrane-localized Ca²⁺-conducting CNGC2 channel. This mutant also displays no hypersensitive response to infection by some pathogens (Clough et al., 2000; Ali et al., 2007). In addition to involvement in pathogen-mediated Ca²⁺ signaling, CNGC2 has been suggested to participate in the process of leaf development/senescence (Köhler et al., 2001). dnd1 mutant plants have high levels of SA and expression of PR1 (Yu et al., 1998), and spontaneous necrotic lesions appear conditionally in dnd1 leaves (Clough et al., 2000; Jirage et al., 2001). Endogenous H₂O₂ levels in dnd1 mutants are increased from wild-type levels (Mateo et al., 2006). Reactive oxygen species molecules, such as H₂O₂, are critical to the PCD/senescence processes of plants (Navabpour et al., 2003; Overmyer et al., 2003; Hung and Kao, 2004; Guo and Crawford, 2005; Zimmermann et al., 2006). Here, we use the dnd1 mutant to evaluate the relationship between leaf Ca²⁺ uptake during plant growth and leaf senescence. Our results identify NO, as affected by leaf Ca²⁺ level, to be an important negative regulator of leaf senescence initiation. Ca²⁺-mediated NO production during leaf development could control senescence-associated gene (SAG) expression and the production of molecules (such as SA and H₂O₂) that act as signals during the initiation of leaf senescence programs.     

On the move,lysosomal CAX drives Ca2+ transport and motility

Acidic Ca²⁺ stores are important sources of Ca²⁺ during cell signaling but little is known about how Ca²⁺ enters these stores. In this issue, Melchionda et al. (2016. J. Cell Biol. http://dx.doi.org/10.1083/jcb.201510019) identify a Ca²⁺/H⁺ exchanger (CAX) that is required for Ca²⁺ uptake and cell migration in vertebrates.Intracellular Ca²⁺ signaling is of fundamental importance in processes such as cell migration but we do not fully understand the contribution made by different intracellular Ca²⁺ stores to this particular function. Elevation of cytosolic Ca²⁺ by 10- to 100-fold the normal resting levels can occur by entry of external Ca²⁺ across the plasma membrane and release of Ca²⁺ from intracellular organelles such as the ER. Ca²⁺ ions are transported across membranes by ligand-gated ion channels, energy-dependent pumps, and transporters (Berridge et al., 2003; Lloyd-Evans et al., 2010). Intracellular Ca²⁺ levels are regulated in this manner from simple organisms, such as yeast, through to complex multicellular organisms, suggesting a degree of conservation across the taxonomic kingdoms (Patel and Cai, 2015). Recent evidence has indicated that “acidic Ca²⁺ stores” such as lysosomes in mammalian cells are a key intracellular Ca²⁺ signaling store, like the ER (Lloyd-Evans and Platt, 2011; Patel and Muallem, 2011). The Ca²⁺ concentration of the lysosome (500 µM) is similar to the ER (Christensen et al., 2002; Lloyd-Evans et al., 2008) but lysosomes are smaller in volume and their impact on cellular Ca²⁺ signaling seems localized to events that regulate endocytosis, vesicular fusion, and recycling (Ruas et al., 2010; López-Sanjurjo et al., 2013). However, there is a significant amount of evidence emerging that lysosomes are capable of triggering much larger changes in cytosolic Ca²⁺ during signaling via the induction of Ca²⁺ release from the ER. This effect appears to be mediated by the most potent intracellular Ca²⁺–releasing second messenger nicotinic acid adenine dinucleotide phosphate (NAADP), which triggers Ca²⁺ release from lysosomes via two-pore channels (Brailoiu et al., 2009; Calcraft et al., 2009). In addition to two-pore channels, acidic stores also express other Ca²⁺-permeable channels (summarized in Fig. 1).Open in a separate windowFigure 1.Lysosomal Ca²⁺ transporters and channels. Our current understanding of lysosomal Ca²⁺ transport and the proteins that regulate the transport of Ca²⁺ into and out of the lysosome is heavily stacked in favor of Ca²⁺ release channels. To date, voltage-gated (CaV2.1/CACNA1A), ligand-gated (TRPML1 and TRPM2), and nucleotide-gated (TPC1, TPC2, and P2X4) channels have all been identified or implicated in lysosomal Ca²⁺ release (Patel and Cai, 2015). Much less is known about the mechanisms of Ca²⁺ entry into lysosomes. In lower order organisms, CAX mediates lysosomal Ca²⁺ entry against the proton gradient. In this issue, Melchionda et al. (2016) provide the first evidence for a mammalian lysosomal Ca²⁺ uptake mechanism in nonplacental mammals. These findings provide further support for the key role of the lysosome as an intracellular Ca²⁺ store.Despite recent advances in our knowledge of lysosomal Ca²⁺ release channels, we have so far failed to identify the transport proteins that fill the lysosome with Ca²⁺. Ca²⁺ entering the cell by endocytosis is removed by the early endosome after the initiation of endosomal acidification by the vATPase; therefore, it is likely that lysosomes have their own transporters or pumps to take up Ca²⁺ (Gerasimenko et al., 1998; Christensen et al., 2002). Although there have been studies suggesting the presence of ATPases and putative ion exchangers on mammalian cells (Styrt et al., 1988), the identity of the proteins that mediate lysosomal Ca²⁺ uptake remains elusive. In this issue, Melchionda et al. describe the first lysosomal CAX in nonplacental mammals and link lysosomal Ca²⁺ import via CAX to the maintenance of normal cellular migration during development.To identify novel regulators of Ca²⁺ transport in vertebrates, Melchionda et al. (2016) searched gene databases for homologues of the CAX proteins, which are known to use the proton gradient across the vacuole to drive Ca²⁺ uptake in plant and yeast cells (Dunn et al., 1994). They identified putative CAX genes in many species, from sea urchins and frogs to reptiles and birds. The CAX homologues discovered in the genomes of the platypus and Tasmanian devil are the first lysosomal Ca²⁺ exchangers to be identified in any mammalian species. This new work is a significant finding as it suggests that these mechanisms do clearly exist in some mammalian cells and are required for lysosomal Ca²⁺ store filling. To examine the regulation of lysosomal Ca²⁺ uptake by vertebrate CAX transporters, the authors cloned full-length CAX from the frog and found that expression of frog CAX could rescue Ca²⁺ transport in yeast lacking their own CAX. Furthermore, the authors show that the frog CAX channels correctly localize to lysosomes when expressed in human cell lines and that these CAX are capable of manipulating lysosomal and cytosolic Ca²⁺ levels (in a manner perhaps comparable to plasma membrane Ca²⁺ ATPases). The findings reported in Melchionda et al. (2016) also have significance for researchers who are using simpler model organisms to characterize mechanisms regulating acidic store Ca²⁺. A study by Churchill et al. (2002) that used acidic stores purified from sea urchin egg homogenate to monitor acidic store Ca²⁺ entry concluded that vanadate-sensitive Ca²⁺ pumps were absent and suggested instead the presence of a CAX. This now appears to be the case through the reported cloning of sea urchin CAX. The findings of Melchionda et al. (2016) are a step forward in unraveling the molecular mechanisms of Ca²⁺ handling in model animals.Ca²⁺ signaling plays an important role in development, particularly for cellular migration, where localized elevations in intracellular Ca²⁺ drive rearrangement of the cytoskeleton, cellular contraction, and adhesion (Wei et al., 2009; Sumoza-Toledo et al., 2011; Praitis et al., 2013). A concentration gradient of Ca²⁺ exists across the migrating cell, with higher levels at the rear that contribute to cellular detachment and contraction (Praitis et al., 2013). Recent evidence has highlighted the presence of Ca²⁺ flickers at the leading edge of the migrating cell that have been shown to underlie changes in direction (Wei et al., 2009). Despite the clear importance of Ca²⁺ in mediating cellular migration events and the emergent role of lysosomes in maintaining intracellular Ca²⁺ signaling, very little is known about the roles of lysosomal Ca²⁺ stores in cellular migration. ER Ca²⁺ channels including the inositol 1,4,5-trisphosphate receptors and ryanodine receptors as well as the secretory pathway Ca²⁺ ATPase and lysosomal TRPM2 have all been implicated in regulating changes in intracellular Ca²⁺ to mediate cellular migration, but to date no lysosomal transporters have been implicated in this process (Wei et al., 2009; Sumoza-Toledo et al., 2011; Praitis et al., 2013). Melchionda et al. (2016) investigated the migration of neural crest cells during frog development to find out whether or not CAX transporters control cell motility. CAX proteins are expressed in the neural crest of developing frogs and morpholino-mediated knockdown of CAX expression increased cytosolic Ca²⁺ levels and impeded neural crest cell migration. Confocal imaging of neural crest tissue in vitro revealed the dynamic recruitment of CAX-containing vesicles to the protrusions that contain focal adhesion complexes at the leading edge of the migrating neural cells. Loss of CAX protein expression reduced the ability of neural crest cells to form stable focal adhesions and undergo the initial cell spreading required for migration. The work presented by Melchionda et al. (2016) is a significant discovery providing evidence that lysosomal Ca²⁺ uptake is involved in cell migration and that lower organisms are useful model systems to investigate the role of acidic store Ca²⁺ in this critical cellular function during embryo development.Melchionda et al. (2016) have made a significant step forward in our understanding of the mechanisms that regulate lysosomal/vacuolar Ca²⁺ entry. However, we remain in the dark about the identity of the transporters that pump Ca²⁺ into the lysosomes of placental mammals. What led to the loss of CAX genes in these organisms is as much a mystery as the identity of the transporters that have replaced CAX. Evidence from a study using purified mammalian lysosomes to observe Ca²⁺ uptake indicates that the process is ATP-dependent (Styrt et al., 1988). Placental mammals may have completely different ATP-dependent mechanisms governing lysosomal Ca²⁺ uptake compared with lower order organisms and nonplacental mammals. Interestingly, defects in lysosomal Ca²⁺ uptake are associated with two human diseases, Niemann-Pick type C and Chediak-Higashi syndrome (CHS; Styrt et al., 1988; Lloyd-Evans et al., 2008). The lysosomal accumulation of sphingosine, a Ca²⁺ ATPase inhibitor (Lloyd-Evans and Platt, 2011), leads to reduced lysosomal Ca²⁺ levels in Niemann-Pick type C disease cells and defects in NAADP-mediated lysosomal Ca²⁺ release (Lloyd-Evans et al., 2008). In CHS, there have been reports of enhanced lysosomal Ca²⁺ ATPase transporter activity in neutrophils (Styrt et al., 1988). Interestingly, CHS leukocytes show alterations in chemotaxis with a reduced response to chemotactic factors (Clark and Kimball, 1971), which is supportive of the findings of Melchionda et al. (2016). Much remains to be elucidated about the enigma of mammalian lysosomal Ca²⁺ uptake, but the work of Melchionda et al. (2016) begins to pick this mystery apart.     

FERONIA and Her Pals: Functions and Mechanisms

Current research into the FERONIA family of receptor kinases highlights both questions and opportunities for understanding signaling strategies in plant growth and survival.FERONIA and 16 closely related proteins form a distinct clade within the Arabidopsis (Arabidopsis thaliana) superfamily of receptor-like kinases (RLKs), transmembrane proteins with an extracellular domain for signal perception and a cytoplasmic domain that phosphorylates target molecules and induces cellular responses to incoming signals. Several members of this family, such as THESEUS1 and ANXUR1,2, are known to play distinct roles in growth and reproduction; FERONIA is unique in being critically involved in both plant growth and reproduction. The FERONIA family of proteins from Arabidopsis is distinguished from other RLKs by having extracellular protein motifs that share homology with malectin, an animal protein with the capacity to bind dimeric and oligomeric Glc. The possibility that these malectin-like motifs might interact with carbohydrates has generated widespread speculations that these receptor kinases could act as cell-wall sensors, communicating perturbations at the frontline of cell-cell and plant-environment interaction to the cytoplasm to induce responses. Here, we discuss emerging understanding of the functional roles and signaling mechanisms of FERONIA and its related proteins. We also highlight pressing questions, as well as the functional potential of the broader malectin-like domain-containing RLK family that exists across the plant kingdom. We believe FERONIA and her pals provide a rich ground for research with many emerging opportunities for uncovering novel insights into how plants strive for growth and survival.FERONIA/SIRÈNE was first identified genetically more than ten years ago as a key regulator of female fertility in Arabidopsis (Rotman et al., 2003; Huck et al., 2003). It was later determined to be a receptor kinase (Escobar-Restrepo et al., 2007) and one of 17 closely related receptor-like kinases (RLKs) in Arabidopsis (Fig. 1; Hèmaty and Höfte, 2008; Boisson-Dernier et al., 2011; Cheung and Wu, 2011). The name FERONIA (after an Etruscan goddess of fertility) will be used from hereon. Arabidopsis has more than 600 RLKs (Shiu and Bleecker, 2003). Several discoveries made at about the same time led to an extraordinary level of interest in FERONIA and related RLKs. These include: (1) a member of this group, THESEUS1 (named after the Greek mythological figure that slew Procustes the brigand) is a critical regulator of cell growth and appears to function as a surveyor of cell-wall status (Hèmaty et al., 2007); (2) FERONIA functions broadly throughout development and is fundamental to cell and plant growth (Guo et al., 2009; Deslauriers and Larsen, 2010; Duan et al., 2010); and (3) a closely related pair of these RLKs, ANXUR1 and ANXUR2 (named after the consort of FERONIA), is essential for male fertility (Boisson-Dernier et al., 2009; Miyazaki et al., 2009). Last but not least was the report of malectin, a novel protein from animals with the capacity to bind dimeric and oligomeric Glc-binding protein (Schallus et al., 2008) and the realization that FERONIA and related RLKs contain malectin-like motifs (PFAM CL0468) in their extracellular domains (Fig. 1). This led to widespread speculations that FERONIA and related RLKs might interact with carbohydrate moieties and function as sensors of perturbations in the cell wall, communicating conditions at the cell surface to induce appropriate cellular responses (Hèmaty and Höfte, 2008; Boisson-Dernier et al., 2011; Cheung and Wu, 2011; Lindner et al., 2012). FERONIA and related malectin-like domain-containing RLKs are often referred to as the CrRLK1-like RLKs (see e.g. Nibau and Cheung, 2011), after its founding member identified in Catharanthus roseus, CrRLK1 (Schulze-Muth et al., 1996), but for which no functional work has been reported. THESEUS1 was the first member of the group for which a clear functional role was demonstrated, and FERONIA is the most prevalently studied among these RLKs. To provide a functional context for our discussion here, we will refer to the FERONIA-related RLKs in Arabidopsis as the THESEUS1/FERONIA-related RLK family. We update current knowledge about these RLKs from Arabidopsis and highlight pressing questions and emerging opportunities from these and related malectin-like domain-containing RLKs, which are present throughout the plant kingdom (Hèmaty and Höfte, 2008; Antolin-Llovera et al., 2014; Nguyen et al., 2015).Open in a separate windowFigure 1.FERONIA protein domain structure and phylogenetic tree of the Arabidopsis THESEUS1/FERONIA receptor kinase family. A, Deduced FERONIA structural domains. SS, ECD, TM are, respectively, signal peptide, extracellular domain, transmembrane domain. MALA and MALB are tandem malectin-like domains. exJM, extracellular juxtamembrane region. Numbers indicate amino acid residues. B, The THESEUS1/FERONIA protein family.

ADVANCES

• FERONIA and related RLKs have extracellular motifs homologous with the diglucose-binding protein malectin, so potentially interact with cell wall carbohydrates and mediate wall-related activities.
• FERONIA controls growth and female fertility, mediates hormone- and pathogen-induced responses, and is required for a normal cell wall.
• FERONIA is a receptor for RALF1, a peptide regulatory factor, which affects phosphorylation of FERONIA and the key cell growth regulator H⁺-ATPase.
• FERONIA-related THESEUS1 suppresses growth in cellulose-deficient mutants, suggesting a role as surveyor of wall conditions.
• FERONIA homologs ANXUR1 and ANXUR2 ensure pollen tube integrity and male fertility.
• FERONIA, ANXUR1, and ANXUR2 signaling collectively involves a GPI-AP, a MLO protein, the RHO GTPase switch, NADPH oxidases, and a receptor-like cytoplasmic kinase; ROS and Ca²⁺ are key elements in their functions.

Extracellular homology with malectin (Schallus et al., 2008; Fig. 1A) distinguishes the THESEUS1/FERONIA-related RLKs from other members of the Arabidopsis RLK family. Malectin, named after its in vitro ability to bind maltose (Glc α1-4 Glc), is a conserved animal protein located in the lumen of the endoplasmic reticulum where it is involved in protein quality control in the early steps of secretion (Schallus et al., 2008; Qin et al., 2012). A majority of the Arabidopsis THESEUS1/FERONIA family (Fig. 1B) has tandem malectin-like motifs (Fig. 1A; Boisson-Dernier et al., 2011) in their extracellular domains. We focus our discussion on FERONIA, THESEUS1, and three other members of the family, ANXUR1, ANXUR2, and ERULUS/[Ca²⁺]_cyt-associated Protein Kinase1, for which clear biological roles have been demonstrated (Miyazaki et al., 2009; Boisson-Dernier et al., 2009; Bai et al., 2014). A contribution to cell growth and morphogenesis has also been reported for HERCULES1 (Guo et al., 2009) and CURVY (Gachomo et al., 2014), but details regarding their functions remain limited.     

A rise in NAD precursor nicotinamide mononucleotide (NMN) after injury promotes axon degeneration

NAD metabolism regulates diverse biological processes, including ageing, circadian rhythm and axon survival. Axons depend on the activity of the central enzyme in NAD biosynthesis, nicotinamide mononucleotide adenylyltransferase 2 (NMNAT2), for their maintenance and degenerate rapidly when this activity is lost. However, whether axon survival is regulated by the supply of NAD or by another action of this enzyme remains unclear. Here we show that the nucleotide precursor of NAD, nicotinamide mononucleotide (NMN), accumulates after nerve injury and promotes axon degeneration. Inhibitors of NMN-synthesising enzyme NAMPT confer robust morphological and functional protection of injured axons and synapses despite lowering NAD. Exogenous NMN abolishes this protection, suggesting that NMN accumulation within axons after NMNAT2 degradation could promote degeneration. Ectopic expression of NMN deamidase, a bacterial NMN-scavenging enzyme, prolongs survival of injured axons, providing genetic evidence to support such a mechanism. NMN rises prior to degeneration and both the NAMPT inhibitor FK866 and the axon protective protein Wld^S prevent this rise. These data indicate that the mechanism by which NMNAT and the related Wld^S protein promote axon survival is by limiting NMN accumulation. They indicate a novel physiological function for NMN in mammals and reveal an unexpected link between new strategies for cancer chemotherapy and the treatment of axonopathies.Axon degeneration in disease shares features with the progressive breakdown of the distal segment of severed axons as described by Augustus Waller in 1850 and named Wallerian degeneration.¹ The serendipitous discovery of Wallerian degeneration slow (Wld^S) mice, where transected axons survive 10 times longer than in wild types (WTs),² suggested that axon degeneration is a regulated process, akin to apoptosis of the cell bodies but distinct in molecular terms.^3,4 This process appears conserved in rats, flies, zebrafish and humans.^{5, 6, 7, 8} Wld^S blocks axon degeneration in some disease models, indicating a mechanistic similarity.³ Therefore understanding the pathway it influences is an excellent route towards novel therapeutic strategies.Wld^S is a modified nicotinamide mononucleotide adenylyltransferase 1 (NMNAT1) enzyme, whose N-terminal extension partially relocates NMNAT1 from nuclei to axons, conferring gain of function.^9,10 In mammals, three NMNAT isoforms, nuclear NMNAT1, cytoplasmic NMNAT2 and mitochondrial NMNAT3, catalyse nicotinamide adenine dinucleotide (NAD) synthesis from nicotinamide mononucleotide (NMN) and adenosine triphosphate (ATP; Figure 1a).^11,12 Several reports indicate Wld^S protects injured axons by maintaining axonal NMNAT activity.^{13, 14, 15} In WT injured axons, without Wld^S, NMNAT activity falls when the labile, endogenous axonal isoform, NMNAT2, is no longer transported from cell bodies.¹⁶ NMNAT2 is required for axon maintenance¹⁶ and for axon growth in vivo and in vitro,^17,18 and modulation of its stability by palmitoylation¹⁹ or ubiquitin-dependent processes both in mice or when ectopically expressed in Drosophila^{19, 20, 21} has a corresponding effect on axon survival.Open in a separate windowFigure 1FK866 acts within axons to delay degeneration after injury. (a) The salvage pathway of NAD biosynthesis from nicotinamide (Nam) and nicotinic acid (Na). Only NAD biosynthesis from Nam is sensitive to FK866, which potently inhibits NAMPT while having no effect on nicotinic acid phosphoribosyltransferase (NaPRT).²⁹ The reaction catalysed by bacterial NMN deamidase is also shown. (b) SCG explants were treated with 100 nM FK866 for the indicated times, and then the whole explants (top panel) or the cell bodies (bottom left panel) and neurite fractions (bottom right panel) were separately collected. NAD was determined with an HPLC-based method (see Materials and Methods; n=3, mean and S.D. shown). (c) SCG neurites untreated (top panels) or treated with 100 nM FK866 the day before transection (bottom panels) and imaged after transection at the indicated time points. (d) SCG explants were treated with 100 nM FK866 1 day before or at the indicated times after cutting their neurites. Degeneration index was calculated from three fields in 2–4 independent experiments. The effect of treatment is highly significant when the drug is preincubated or added at 0–4 h after cut (mean ±S.E.M., n=6–12, one-way ANOVA followed by Bonferroni''s post-hoc test, *P<0.05, **P<0.01, ***P<0.001, ****P<0.0001, compared with untreated)Wld^S partially colocalizes with mitochondria^14,22 and was shown to increase mitochondria motility and Ca²⁺-buffering capacity.²³ Inhibiting mitochondrial permeability transition pore protects degenerating axons.²⁴ However, Wld^S is protective in axons devoid of mitochondria,⁸ and targeting a cytosolic variant of NMNAT2 to mitochondria abolished its protective effect,¹⁹ suggesting a late mitochondrial involvement in Wallerian degeneration.Despite the importance of NMNAT activity in axon survival and degeneration, the molecular players remain elusive. Although NMNAT activity is required for protection,¹³ the hypothesis that increased NAD levels are responsible^25,26 does not fit some data.^27,28While further investigating the role of NAD, we found that blocking nicotinamide phosphoribosyltransferase (NAMPT, the enzyme preceding NMNAT, Figure 1a), was surprisingly axon-protective despite lowering NAD. NAMPT catalyses the synthesis of NMNAT-substrate NMN, the rate-limiting step in the NAD salvage pathway from nicotinamide (Nam) (Figure 1a). Here, we show that NMN accumulates after axon injury, and we provide genetic and pharmacological evidence supporting a role for this NMN increase in axon degeneration when NMNAT2 is depleted. We reveal an unexpected new direction for research into the degenerative mechanism, a novel class of protective proteins and new players in an axon-degeneration pathway sensitive to drugs under development for cancer.