TY - JOUR
AU - Cunniff,, Brian
AB - Abstract Mitochondria are not passive bystanders aimlessly floating throughout our cell’s cytoplasm. Instead, mitochondria actively move, anchor, divide, fuse, self-destruct and transfer between cells in a coordinated fashion, all to ensure proper structure and position supporting cell function. The existence of the mitochondria in our cells has long been appreciated, but their dynamic nature and interaction with other subcellular compartments has only recently been fully realized with the advancement of high-resolution live-cell microscopy and improved fractionization techniques. The how and why that dictates positioning of mitochondria to specific subcellular sites is an ever-expanding research area. Furthermore, the advent of new and improved functional probes, sensitive to changes in subcellular metabolite levels has increased our understanding of local mitochondrial populations. In this review, we will address the evidence for intentional mitochondrial positioning in supporting subcellular mitochondrial metabolite levels, including calcium, adenosine triphosphate and reactive oxygen species and the role mitochondrial metabolites play in dictating cell outcomes. metabolite gradients, mitochondrial contacts, mitochondrial dynamics, reactive oxygen species Mitochondrial networks are not identical between two cells of the same origin and certainly not across cell types/tissue origin. Although this is outside the scope of this review, it is nonetheless important to establish a baseline of observations regarding mitochondrial networks in cells. Traditionally, mitochondria have been observed in mammalian cells using electron microscopy and fluorescent microscopy techniques. Using these methods, mitochondria are observed as ‘tube-like’ structures that are densely located in the perinuclear space and emanate outward towards the cell periphery, often as individual mitochondria or a branched network of tubules (Fig. 1). More complex organization has been observed in migrating fibroblasts (1–3), rolling lymphocytes (4), neurons (5), leukocytes (6) and polarized cells such as intestinal epithelial cells (7), proximal kidney cells (7), the lung epithelium (8) and pancreatic acinar cells (9). As techniques to visualize mitochondrial networks in more physiologically accurate models evolves, investigation into the role of particular mitochondrial presentations will aid in our understanding of their localized function. Fig. 1 Open in new tabDownload slide (A) Schematic of mitochondrial positioning in multiple cell types. Intentional mitochondrial positioning supports fibroblast cell migration, Ca2+ signalling in epithelial cells, lymphocyte attachment to endothelial cells, neuron heath and function and T-cell uropod formation. (B) Mitochondria (red) of a mouse embryonic fibroblast (MEF). Dotted line indicates cell outline. Fig. 1 Open in new tabDownload slide (A) Schematic of mitochondrial positioning in multiple cell types. Intentional mitochondrial positioning supports fibroblast cell migration, Ca2+ signalling in epithelial cells, lymphocyte attachment to endothelial cells, neuron heath and function and T-cell uropod formation. (B) Mitochondria (red) of a mouse embryonic fibroblast (MEF). Dotted line indicates cell outline. In this review, we will describe the molecular basis underlying mitochondrial dynamics to provide a framework for the role of subcellular mitochondrial positioning in dictating local metabolite levels. The intricacies controlling the signalling for recruitment, docking and local production of mitochondrial metabolites is still beyond our understanding, but nonetheless a clearer picture is forming regarding the dynamic control of mitochondrial populations supporting cell function. Mitochondrial structure Mitochondria are composed of an outer mitochondrial membrane (OMM) and an inner mitochondrial membrane (IMM) that gives rise to the intermembrane space (IMS) and matrix space (10). The components of these membranes and relevance of the compartmentalization have been extensively reviewed elsewhere and therefore will not be elaborated on within (11). Briefly, numerous proteins, RNA and metabolites are specifically localized to the OMM, IMS, IMM and matrix. Primary examples of this is the electron transport chain (ETC) complexes that are partially embedded and specifically oriented in the IMM to produce energy through oxidative phosphorylation (11). The electrochemical gradient across the IMM fuels oxygen consumption and adenosine triphosphate (ATP) synthesis on the matrix side of the IMM. Additionally, ∼1,000 nuclear encoded proteins are imported into the mitochondria through a series of membrane translocation complexes that recognize a N-terminal mitochondrial localization sequence that destines nuclear encoded proteins to reside in the various mitochondrial compartments (12). The selective compartmentalization between mitochondrial membranes and space leads to specific protein, RNA and metabolite composition in the relative spaces. Control of mitochondrial structure Mitochondria undergo structural transitions, splitting one mitochondrion into two through fission and bringing two mitochondria together into one through fusion (Fig. 2). The factors involved in mitochondrial fission/fusion have been reviewed elsewhere and therefore will not be covered in detail (13). In short, fission of the OMM is initialized through mitochondrial contact with the endoplasmic reticulum and the actin cytoskeleton to physically constrict mitochondrial tubules (14). In mammals, the ER-localized formin INF2 has been proposed to stimulate actin polymerization, facilitating mitochondrial division by providing a force-generating system to drive the initial constriction of mitochondria and by enhancing ER–mitochondria contact and calcium transfer to constrict the IMM (15). The ER was found to be present at over half of the mitochondrial fusion events observed and the duration of fusion events from initial contact to completion of fusion was shorter for fusion events associated with the ER in comparison to those not associated with the ER (16). Membrane constriction facilitates the assembly of dynamin-related protein 1 (DRP1) by OMM receptors (MiD49, MiD51, Mff, Fis1) to further constrict and pinch the OMM (17). Mitochondrial fusion is facilitated by the action of mitofusion proteins (Mfn1 and Mfn2), proteins that physically bring mitochondrial membranes into close contact and drive membrane fusion in a GTP-dependent manner (18). Mfn2 has also been shown to contain redox sensitive cysteines on the IMS facing C-terminus that support reactive oxygen species (ROS)-induced Mfn2 dimerization (19). Mitochondrial fusion supports increased respiratory function, ATP production and escape from stress insults. Mitochondrial fission/fusion dynamics support numerous cell processes including mitochondrial redistribution, coordinated degradation (mitophagy) and equal distribution of mitochondria during cell division (20, 21). Defects in both mitochondrial fission and fusion compromise mitochondrial function and contribute to numerous disease states (22). Fig. 2 Open in new tabDownload slide (A) (1) Mitochondrial networks exist as individual components and elongated networks. (2) Mitochondrial fission is initiated by contact with the endoplasmic reticulum and actin structures followed by assembly of pro-fission molecules (DRP1, DNM2) on mitochondrial membranes through interaction with resident adapter proteins (Mid) to facilitate membrane constriction. (3) Mitochondrial fission promotes the separation of mitochondrial components and the redistribution of mitochondrial structures. (4) Mitochondrial fusion brings individual mitochondria together as a stress response and to increase metabolic capacity. Mitochondrial-derived vesicles or compartments pinch off from mitochondria to facilitate removal of damaged proteins and act as signalling structures. (B) Mitochondria couple to microtubule motors through the action of adapter proteins Miro1 and Trak1/2. Signal input in the form of metabolic and/or second messenger signals controls mitochondrial movement. Mitochondrial movement promotes localized production of mitochondrial metabolites such as ATP and ROS. Fig. 2 Open in new tabDownload slide (A) (1) Mitochondrial networks exist as individual components and elongated networks. (2) Mitochondrial fission is initiated by contact with the endoplasmic reticulum and actin structures followed by assembly of pro-fission molecules (DRP1, DNM2) on mitochondrial membranes through interaction with resident adapter proteins (Mid) to facilitate membrane constriction. (3) Mitochondrial fission promotes the separation of mitochondrial components and the redistribution of mitochondrial structures. (4) Mitochondrial fusion brings individual mitochondria together as a stress response and to increase metabolic capacity. Mitochondrial-derived vesicles or compartments pinch off from mitochondria to facilitate removal of damaged proteins and act as signalling structures. (B) Mitochondria couple to microtubule motors through the action of adapter proteins Miro1 and Trak1/2. Signal input in the form of metabolic and/or second messenger signals controls mitochondrial movement. Mitochondrial movement promotes localized production of mitochondrial metabolites such as ATP and ROS. Portions of the mitochondria can be eliminated by mitochondrial-derived vesicles (MDVs), which deliver oxidized mitochondrial proteins to the lysosome or late endosome for degradation (23). MDVs are likely the mitochondria’s first round of defence to dispose of damaged proteins before they can interrupt mitochondrial function and cause breakdown (24). Studies have found that MDVs are enriched with oxidized proteins. Proteins from the outer membrane, inner membrane or the matrix are selectively incorporated into MDVs based on the type of mitochondrial stress they experience (24). Selective removal of proteins to protect the mitochondria in times of stress is also achieved by mitochondrial-derived compartments (MDCs), which have been investigated in yeast. Membrane proteins lacking defined mitochondrial-targeting sequences are often sequestered to MDCs. Formation of MDCs requires the import receptors Tom70/71. MDCs are then engulfed by autophagosomes and delivered to the vacuole for degradation by autophagy (23). The purpose of the MDC pathway may also be to maintain regulated nutrient influx into the mitochondria by sequestering nutrient transporters in response to cytoplasmic nutrient overload (23). Mitochondrial-associated membranes for metabolite transfer Mitochondria associate with other cellular organelles to facilitate the exchange of cargo and information. These associations are critical to organelle and cell homeostasis, information transfer through signalling cascades and regulated cellular processes such as apoptosis (25). The mitochondrial-associated ER membrane (MAM) is composed of a unique set of proteins that interact with mitochondrial proteins which facilitates lipid exchange between the ER and mitochondria to promote mitochondrial membrane integrity as well as inter-organelle trafficking of lipid intermediates destined for metabolism or other biosynthetic pathways (26). Mitochondria–ER contacts are also involved in bioenergetics, intracellular signalling, inflammation, autophagy and apoptosis (27). Calcium ion signalling requires close interaction between the ER and the mitochondria (28). Calcium microdomains exist at contact points between mitochondria and the ER that facilitate Ca2+ transfer from the ER to the mitochondria. Mitochondria can store calcium transiently, making them efficient buffers for cytosolic calcium. Once in the mitochondrial matrix, Ca2+ stimulates mitochondrial ATP synthesis by activating the TCA cycle. When the Ca2+ transfer is excessive and sustained, mitochondrial Ca2+ overload induces apoptosis (28). Many regulatory proteins can be found at MAMs to maintain an optimal distance between the ER and mitochondria and coordinate their Ca2+ transporters/channels (28). For example, the endoplasmic reticulum stress transducer IRE1α plays a role in the structure of MAMs by operating as a scaffold and controls mitochondrial calcium uptake. IRE1α deficiency results in altered mitochondrial physiology and energy metabolism (29). Additionally, in yeast Gem1, the orthologue of Miro, is enriched at ER–mitochondria contact sites and contains multiple Ca2+-binding domains that allow for regulation (30). In addition to the ER, mitochondria contact the Golgi, vacuoles, lysosomes, endosomes, peroxisomes, lipid droplets and the plasma membrane (31). Mitochondrial interaction with the Golgi apparatus provides ATP and helps establish calcium gradients across the Golgi. Mitochondria also play a role in segregating the Golgi from the plasma membrane and nucleus (32). In yeast, mitochondria interact with vacuoles for lipid metabolism and this contact is mediated by vCLAMP (33). Additionally, Mcp1 recruits Vps13 to the mitochondria to support transport between the mitochondria and vacuole (31). Mitochondrial contact with lysosomes has also been linked to mitochondrial division. Lysosomal GTP-bound RAB7 has been shown to be involved in the formation and stabilization of mitochondria–lysosome contacts (34). Mitochondria also come in contact with endosomes (containing transferrin) for iron transfer. The molecular basis behind this is unclear, but the duration of this contact is known to be regulated by iron concentration. Halotransferrin facilitates disassociation once the mitochondria have taken up the iron (35). Mitochondria interact with peroxisomes for the transfer of β-oxidation products from peroxisomes to mitochondria. In yeast, Pex34 and Fzo1 have been identified as components of the tether between mitochondria and peroxisomes (36). In yeast, the mitochondria–ER–cortex anchor (MECA) tethers mitochondria to the plasma membrane. The core protein component of MECA, Num1, interacts directly with both the mitochondrial membrane and plasma membrane. The tethering activity of Num1 is required for proper mitochondrial distribution and inheritance during budding yeast mitosis (37). Control of mitochondrial movement Mitochondria do not stay fixed to the perinuclear space but are dynamically transported throughout the entirety of the cell (38). The movement of mitochondria is primarily controlled by the action of microtubule motors (kinesin and dynein) and mitochondrial adapter proteins providing the tether to motors (Fig. 2) (39–41). Additional factors such as actin, actin motors, intermediate filaments, ER/mitochondrial contacts and microtubule binding proteins have also been shown to mediate the movement and docking of mitochondria. Mitochondria tether to kinesin and dynein through the action of the OMM adapter proteins Miro1/2 and the cytosolic adapters Trak1/2. Miro1/2 are embedded in the OMM at their C-terminus and form complexes with cytosolic Trak1/2 to connect mitochondria to microtubule motors. Additional factors including syntaphilin have been implicated in the activity of this complex and regulation of mitochondrial movement in neurons and tumour cells (42, 43). The dense perinuclear bundle of mitochondria can be localized to either the anterior or posterior of the cell, depending on cell type (44). Furthermore, the position of the perinuclear bundle is not fixed in all cells and can reposition based on cellular context (44). Early studies in rat intestinal cells showed that changes in diet caused dramatic redistribution of mitochondrial populations (45, 46). The majority of mitochondria were present at the basal surface of intestinal cells in fed animals, upon fasting mitochondria populations were reoriented towards the apical cell surface. Additionally, lipid feeding led to a basal orientation of mitochondrial networks while glucose fed animals showed apical polarization of mitochondria, indicating that the specific nutrient supply had pronounced effects on mitochondrial position (45, 46). Endothelial cells actively move their mitochondria towards the nucleus in response to hypoxia, concentrating mitochondrial reactive oxygen species (mROS) to the perinuclear space, inducing oxidative modifications on target genes to modulate their expression (47). Immune cells move the primary mitochondrial bundle towards the uropod to support cell–cell contacts required for cell clearance (6). Lymphocytes rolling and attaching along the endothelial layer reposition their mitochondrial networks to support cell attachment (4). This is largely mediated by the primary mitochondrial adapter protein Miro1. In larger cells such as neurons, mitochondria are transported over long distances on microtubule tracks to be deposited at axon terminals (48). Mitochondria pause at sites of increased calcium, partially induced through the action of Ca2+ binding EF-hand motifs in Miro1, and act as localized sinks to buffer excess calcium levels (5). In more recent studies, ROS have been shown to slow mitochondrial movement in neurons, presumably through modifications to the Miro/Trak adapter complex (49). In smaller cells, such as migrating mouse embryonic fibroblasts (MEFs), single mitochondria are actively transported anterograde towards the polarized leading edge. Mitochondria do not actively move towards the trailing edge, but rather are slowly moved back towards the perinuclear space by an unidentified mechanism (1). Miro1, versus Miro2, is primarily responsible for this movement as MEFs lacking Miro1 fail to move mitochondria into leading edge protrusions (3), while MEFs lacking Miro2 appear to have normal mitochondrial networks (50). Fibroblasts lacking Miro1 harbour perinuclear restricted mitochondria and fail to migrate as efficiently as wild-type MEFs (3). The perinuclear mitochondrial bundle of Miro1−/− MEFs is less polarized compared to Miro1+/+ MEFs (B. Cunniff, unpublished results), providing further evidence in the role of Miro1 in polarizing the dense perinuclear mitochondrial bundle. Local activation of the energy sensing enzyme AMPK, specifically in the leading edge of migrating cells, induces directed mitochondrial movement into leading edge structures, supporting actin ruffling, a readout of active cell migration (1). Conversely, pharmacologic inactivation of AMPK slows mitochondrial movement and compromises cell migration (1). These studies shed light on the dynamic control of mitochondrial positioning across diverse cell types. Mitochondrial positioning in dictating subcellular energy levels Mitochondria are the primary source of ATP in most cells. Although still requiring much investigation, cells appearing to be primarily glycolytic, including cancer cells, still require mitochondrial function to grow and survive (51). Several studies in the early 1980s described the existence of separate ATP pools within the cytosol of mammalian cells without any indication of physical compartmentalization of these pools (52). Aw and Jones (52) went on to investigate the hypothesis that multiple pools of ATP reflect intracellular ATP gradients as a result of ATP production and consumption in different subcellular regions. Results from this work showed that ATP levels are not homogenous throughout the cell cytoplasm as enzymes localized to the cell membrane (peripheral) are more sensitive to ATP depletion than soluble enzymes (52). The flux of ATP from mitochondrial clusters to peripheral sites was hypothesized to account for these observations. Additional work indicated that ATP levels were highest at mitochondrial membranes in yeast when the activity of recombinant luciferase tethered to the mitochondrial membrane was compared to soluble luciferase in the presence of endogenously added hexokinase (53). In the presence of excess hexokinase, the mitochondrial tethered luciferase retained ATP-dependent activity while soluble luciferase activity was significantly reduced. Thus, ATP accumulates near the mitochondrial membranes where it is produced (53). Additional evidence for subcellular ATP gradients came from studies with Xenopus laevis egg extracts to model metabolic gradients (54). Using this system, it was observed that an energy demand signalling gradient existed around ATP-consuming beads and this was sensitive to oxygen availability (54). Together, these studies provide substantial evidence for the microheterogeneity of ATP in the cell cytoplasm and support the hypothesis that local mitochondrial populations dictate subcellular metabolite gradients (Fig. 3). Fig. 3 Open in new tabDownload slide Schematic describing the existence of intracellular metabolite gradients dependent on subcellular mitochondrial positioning. As mitochondria are dispersed throughout the cell, mitochondrial metabolites become homogenous and gradients are reduced. The interplay between healthy and damaged mitochondria may dictate subcellular mitochondrial metabolite gradients. Fig. 3 Open in new tabDownload slide Schematic describing the existence of intracellular metabolite gradients dependent on subcellular mitochondrial positioning. As mitochondria are dispersed throughout the cell, mitochondrial metabolites become homogenous and gradients are reduced. The interplay between healthy and damaged mitochondria may dictate subcellular mitochondrial metabolite gradients. With the advent of high spatial and temporal resolution microscopy, our ability to visualize dynamic changes to subcellular organelles in real time has drastically increased our understanding of mitochondrial dynamics (16). The ability to visualize mitochondria has elucidated a dynamic network of tubules reorganizing throughout the cytoplasm to sites of high energy demand (1, 3). Numerous groups have investigated the role of mitochondrial positioning in dictating subcellular metabolite levels and uncovered the importance of local mitochondrial populations regulating dynamic cell processes (55). Although some studies have conclusively shown the necessity for local mitochondrial populations for governing their cell response of interest, direct measurement of local metabolites contributing to this process is still lacking. Early observations in neurons indicated mitochondrial density was increased at sites of high energy demand including synapses and in active growth cones and branches (56). Nerve growth factor was found to stimulate mitochondrial recruitment to axon shafts and induce outgrowth, once removed mitochondrial density was reduced at these sites (57). Using a cell body (CB)/pseudopod (PD) assay in which leading edge PDs can be fractionated away from nuclear and perinuclear CB compartments, was observed that PD compartments with lower mitochondrial density had reduced ATP:ADP levels compared to CB compartments containing high levels of mitochondria (1). Although ATP levels were observed to be higher in PDs compared to CBs, higher ADP levels, presumably from high ATP turnover rate, significantly reduced the ATP:ADP ratio in PD compartments. Treatment of PDs with rotenone, a drug that inhibits complex I of the mitochondiral ETC significantly reduced ATP levels, providing evidence that mitochondria were the primary source of energy in PDs. Furthermore, stimulating PDs with the AMP analogue AICAR induced mitochondrial recruitment to PDs and leading edge structures and significantly increased ATP levels in PDs (1). These observations provided clear evidence for the subcellular positioning of mitochondria in dictating subcellular energy levels (Fig. 4). Fig. 4 Open in new tabDownload slide (A) The subcellular ATP/ADP ratio in CB and PD compartments is modulated by local mitochondrial density. AMPK-dependent mitochondrial recruitment to PDS increases local ATP/ADP in PDs and supports cell migration (1). (B) We hypothesize local protein redox status (red = reduced thiol, OX = oxidized thiol) is modulated by mitochondrial recruitment to subcellular sites. Fig. 4 Open in new tabDownload slide (A) The subcellular ATP/ADP ratio in CB and PD compartments is modulated by local mitochondrial density. AMPK-dependent mitochondrial recruitment to PDS increases local ATP/ADP in PDs and supports cell migration (1). (B) We hypothesize local protein redox status (red = reduced thiol, OX = oxidized thiol) is modulated by mitochondrial recruitment to subcellular sites. Subcellular energy gradients have now been visualized in MEFs using the ratiometric ATP:ADP Biosensor Perceval HR (3). Using Perceval HR, it was observed that cells lacking Miro1, which have perinuclear restricted mitochondrial networks (58), have a steeper drop in ATP:ADP in peripheral sites lacking mitochondria compared to cells in which mitochondrial networks were unperturbed (3). Additionally, in line with earlier observations in yeast (53), the highest ATP:ADP ratio was found at sites of highest mitochondrial density in all three-dimensions (3). ATP:ADP gradients were completely abolished following incubation with rotenone, indicating mitochondria are the primary source of energy in cultured MEFs (3). Compartmentalized redox systems In addition to being the ‘powerhouse of the cell’, mitochondria are a primary source of ROS (59). mROS is produced as a by-product of mitochondrial metabolism during normal electron flow through the ETC. Approximately 0.15–2% of electrons leak from the ETC and react with molecular oxygen to form superoxide (⁠ O2− ⁠) (59). O2− is spontaneously or enzymatically dismutated to H2O2 through the action of superoxide dismutase enzymes. H2O2 can oxidize a subset of solvent exposed cysteine residues in target proteins leading to alterations in protein structure and function (60). This process, termed redox-dependent signal transduction, is a method of cell signalling that is intimately linked to cellular metabolism and regulates myriad cell process in both normal and disease states. mROS-dependent signalling requires precise localization of both source and target as H2O2 is rapidly metabolized at sites distant to ROS sources (60, 61). ROS are produced from additional cellular sources including enzyme specific production of ROS by NADPH oxidases (NOXs and DUOXs). Tissue scale gradients of H2O2 have been observed in zebrafish (62). Upon tail fin tip amputation, a 150–300-µm wide H2O2 gradient is established extending from the wound margin into the tissue (62). The Nox family member DUOX2 was found to be primarily responsible for the production of this H2O2 gradient. In these experiments, it was determined that the H2O2 gradient was required for recruitment of distant leukocytes to the wound area. Although a tissue scale H2O2 gradient of 150–300 µm was observed, further investigation with refined approaches showed that H2O2 was inactivated by resident antioxidants, including peroxiredoxins, within 30 µm (61). Additionally, a gradient of H2O2 can exist across membranes when a bolus of peroxide is added to cells. It was experimentally determined that the gradient between extracellular and intracellular levels of exogenously added H2O2 is approximately a 390-fold difference (63). Although the H2O2 may be consumed by resident antioxidant proteins within a fixed volume, the possibility of a redox signalling cascade through thiol exchange reactions may extend redox-dependent signalling events far beyond initial local oxidation. Additional evidence supporting localized redox modulation in support of cell migration comes from studies showing that SIRT3, which promotes energy homeostasis, is down-regulated specifically at the leading edge of migrating cells. This is hypothesized to balance leading edge ROS levels in support of tumour cell metastasis (64). Increased evidence is accumulating describing the cross-talk between NOXs and mitochondria to regulate ROS production from either source. Both NOX-dependent regulation of mitochondria and mitochondrial regulation of NOX function have been described (65). For NOX4, which can reside within the mitochondrial membrane (66, 67), it was shown that mitochondrial ATP directly regulates the activity of this enzyme and subsequent NOX4-dependent ROS production (68). For other NOXs that do not reside in the mitochondria, it is reasonable to speculate that these entities must be in close proximity to the mitochondria to elicit regulatory control over the other. The subcellular positioning of mitochondria in regulating NOX activity has not been investigated but is deserving of attention to further elucidate the mechanisms controlling NOX–mitochondria cross-talk. Concluding Remarks The fluid reorganization of intracellular components supports spatial and temporal regulation of dynamic cell processes. Mitochondria, which supply cellular energy (ATP), buffer Ca2+ and produce ROS are a primary hub for regulating cell growth, division, movement and communication. To ensure mitochondrial metabolites exert their effects on local targets, mitochondria must be strategically positioned near target organelles and to subcellular sites. Control of mitochondrial movement is dynamic and few factors governing precise mitochondrial positioning have been uncovered. Therefore, new approaches and techniques to determine factors modulating mitochondrial movement, anchoring, dynamics and metabolic output in the context of supporting downstream cellular processes are imperative. Investigation into these factors may provide valuable insight into new pathways governing normal and disease states and unidentified targetable features of mitochondrial dynamics for therapeutic intervention. Acknowledgements We apologize for not mentioning relevant studies due to space constraints. H.A. and M.H. were recipients of Summer Undergraduate Research Fellowships from the University of Vermont. B.C. receives funding from Department of Pathology, Paredox Therapeutics, LLC, The UVM Larner College of Medicine and the UVM Cancer Center. Conflict of Interest None declared. References 1 Cunniff B. , McKenzie A.J. , Heintz N.H. , Howe A.K. ( 2016 ) AMPK activity regulates trafficking of mitochondria to the leading edge during cell migration and matrix invasion . Mol. Biol. Cell 27 , 2662 – 2674 Google Scholar Crossref Search ADS PubMed WorldCat 2 Cunniff B. , Newick K. , Nelson K.J. , Wozniak A.N. , Beuschel S. , Leavitt B. , Bhave A. , Butnor K. , Koenig A. , Chouchani E.T. , James A.M. , Haynes A.C. , Lowther W.T. , Murphy M.P. , Shukla A. , Heintz N.H. ( 2015 ) Disabling mitochondrial peroxide metabolism via combinatorial targeting of peroxiredoxin 3 as an effective therapeutic approach for malignant mesothelioma . PLoS One 10 , e0127310 Google Scholar Crossref Search ADS PubMed WorldCat 3 Schuler M.H. , Lewandowska A. , Caprio G.D. , Skillern W. , Upadhyayula S. , Kirchhausen T. , Shaw J.M. , Cunniff B. ( 2017 ) Miro1-mediated mitochondrial positioning shapes intracellular energy gradients required for cell migration . Mol. Biol. Cell 28 , 2159 – 2169 Google Scholar Crossref Search ADS PubMed WorldCat 4 Morlino G. , Barreiro O. , Baixauli F. , Robles-Valero J. , Gonzalez-Granado J.M. , Villa-Bellosta R. , Cuenca J. , Sanchez-Sorzano C.O. , Veiga E. , Martin-Cofreces N.B. , Sanchez-Madrid F. ( 2014 ) Miro-1 links mitochondria and microtubule Dynein motors to control lymphocyte migration and polarity . Mol. Cell. Biol . 34 , 1412 – 1426 Google Scholar Crossref Search ADS PubMed WorldCat 5 Schwarz T.L. ( 2013 ) Mitochondrial trafficking in neurons . Cold Spring Harb. Perspect. Biol . 5 , a011304 Google Scholar Crossref Search ADS PubMed WorldCat 6 Campello S. , Lacalle R.A. , Bettella M. , Mañes S. , Scorrano L. , Viola A. ( 2006 ) Orchestration of lymphocyte chemotaxis by mitochondrial dynamics . J. Exp. Med. 203 , 2879 – 2886 Google Scholar Crossref Search ADS PubMed WorldCat 7 Aw T.Y. ( 2000 ) Intracellular compartmentation of organelles and gradients of low molecular weight species . Int. Rev. Cytol . 192 , 223 – 253 Google Scholar Crossref Search ADS PubMed WorldCat 8 Ribeiro C.M. , Paradiso A.M. , Livraghi A. , Boucher R.C. ( 2003 ) The mitochondrial barriers segregate agonist-induced calcium-dependent functions in human airway epithelia . J. Gen. Physiol. 122 , 377 – 387 Google Scholar Crossref Search ADS PubMed WorldCat 9 Tinel H. , Cancela J.M. , Mogami H. , Gerasimenko J.V. , Gerasimenko O.V. , Tepikin A.V. , Petersen O.H. ( 1999 ) Active mitochondria surrounding the pancreatic acinar granule region prevent spreading of inositol trisphosphate-evoked local cytosolic Ca(2+) signals . EMBO J . 18 , 4999 – 5008 Google Scholar Crossref Search ADS PubMed WorldCat 10 Frey T.G. , Mannella C.A. ( 2000 ) The internal structure of mitochondria . Trends Biochem. Sci . 25 , 319 – 324 Google Scholar Crossref Search ADS PubMed WorldCat 11 Kuhlbrandt W. ( 2015 ) Structure and function of mitochondrial membrane protein complexes . BMC Biol . 13 , 89 Google Scholar Crossref Search ADS PubMed WorldCat 12 Wiedemann N. , Pfanner N. ( 2017 ) Mitochondrial machineries for protein import and assembly . Annu. Rev. Biochem. 86 , 685 – 714 Google Scholar Crossref Search ADS PubMed WorldCat 13 Scott I. , Youle R.J. ( 2010 ) Mitochondrial fission and fusion . Essays Biochem. 47 , 85 – 98 Google Scholar Crossref Search ADS PubMed WorldCat 14 Friedman J.R. , Lackner L.L. , West M. , DiBenedetto J.R. , Nunnari J. , Voeltz G.K. ( 2011 ) ER tubules mark sites of mitochondrial division . Science 334 , 358 – 362 Google Scholar Crossref Search ADS PubMed WorldCat 15 Korobova F. , Ramabhadran V. , Higgs H.N. ( 2013 ) An actin-dependent step in mitochondrial fission mediated by the ER-associated formin INF2 . Science 339 , 464 – 467 Google Scholar Crossref Search ADS PubMed WorldCat 16 Guo Y. , Li D. , Zhang S. , Yang Y. , Liu J.J. , Wang X. , Liu C. , Milkie D.E. , Moore R.P. , Tulu U.S. , Kiehart D.P. , Hu J. , Lippincott-Schwartz J. , Betzig E. , Li D. ( 2018 ) Visualizing intracellular organelle and cytoskeletal interactions at nanoscale resolution on millisecond timescales . Cell 175 , 1430 – 1442.e17 Google Scholar Crossref Search ADS PubMed WorldCat 17 Kraus F. , Ryan M.T. ( 2017 ) The constriction and scission machineries involved in mitochondrial fission . J. Cell Sci. 130 , 2953 – 2960 Google Scholar Crossref Search ADS PubMed WorldCat 18 Zorzano A. , Liesa M. , Sebastián D. , Segalés J. , Palacín M. ( 2010 ) Mitochondrial fusion proteins: dual regulators of morphology and metabolism . Semin. Cell Dev. Biol . 21 , 566 – 574 Google Scholar Crossref Search ADS PubMed WorldCat 19 Mattie S. , Riemer J. , Wideman J.G. , McBride H.M. ( 2018 ) A new mitofusin topology places the redox-regulated C terminus in the mitochondrial intermembrane space . J. Cell Biol. 217 , 507 – 515 Google Scholar Crossref Search ADS PubMed WorldCat 20 Chen H. , Chan D.C. ( 2009 ) Mitochondrial dynamics–fusion, fission, movement, and mitophagy–in neurodegenerative diseases . Hum. Mol. Genet . 18 , R169 – R176 Google Scholar Crossref Search ADS PubMed WorldCat 21 Youle R.J. , van der Bliek A.M. ( 2012 ) Mitochondrial fission, fusion, and stress . Science 337 , 1062 – 1065 Google Scholar Crossref Search ADS PubMed WorldCat 22 Archer S.L. ( 2013 ) Mitochondrial dynamics–mitochondrial fission and fusion in human diseases . N Engl. J. Med. 369 , 2236 – 2251 Google Scholar Crossref Search ADS PubMed WorldCat 23 Hughes A.L. , Hughes C.E. , Henderson K.A. , Yazvenko N. , Gottschling D.E. ( 2016 ) Selective sorting and destruction of mitochondrial membrane proteins in aged yeast . Elife 5 , e13943 Google Scholar Crossref Search ADS PubMed WorldCat 24 Sugiura A. , McLelland G.L. , Fon E.A. , McBride H.M. ( 2014 ) A new pathway for mitochondrial quality control: mitochondrial-derived vesicles . EMBO J . 33 , 2142 – 2156 Google Scholar Crossref Search ADS PubMed WorldCat 25 Szabadkai G. , Simoni A.M. , Chami M. , Wieckowski M.R. , Youle R.J. , Rizzuto R. ( 2004 ) Drp-1-dependent division of the mitochondrial network blocks intraorganellar Ca2+ waves and protects against Ca2+-mediated apoptosis . Mol. Cell 16 , 59 – 68 Google Scholar Crossref Search ADS PubMed WorldCat 26 Tagaya M. , Arasaki K. ( 2017 ) Regulation of mitochondrial dynamics and autophagy by the mitochondria-associated membrane . Adv. Exp. Med. Biol . 997 , 33 – 47 Google Scholar Crossref Search ADS PubMed WorldCat 27 Herrera-Cruz M.S. , Simmen T. ( 2017 ) Of yeast, mice and men: mAMs come in two flavors . Biol. Direct 12 , 3 Google Scholar Crossref Search ADS PubMed WorldCat 28 Marchi S. , Patergnani S. , Missiroli S. , Morciano G. , Rimessi A. , Wieckowski M.R. , Giorgi C. , Pinton P. ( 2018 ) Mitochondrial and endoplasmic reticulum calcium homeostasis and cell death . Cell Calcium 69 , 62 – 72 Google Scholar Crossref Search ADS PubMed WorldCat 29 Carreras-Sureda A. , Jaña F. , Urra H. , Durand S. , Mortenson D.E. , Sagredo A. , Bustos G. , Hazari Y. , Ramos-Fernández E. , Sassano M.L. , Pihán P. , van Vliet A.R. , González-Quiroz M. , Torres A.K. , Tapia-Rojas C. , Kerkhofs M. , Vicente R. , Kaufman R.J. , Inestrosa N.C. , Gonzalez-Billault C. , Wiseman R.L. , Agostinis P. , Bultynck G. , Court F.A. , Kroemer G. , Cárdenas J.C. , Hetz C. ( 2019 ) Non-canonical function of IRE1alpha determines mitochondria-associated endoplasmic reticulum composition to control calcium transfer and bioenergetics . Nat. Cell Biol. 21 , 755 – 767 Google Scholar Crossref Search ADS PubMed WorldCat 30 Kornmann B. , Osman C. , Walter P. ( 2011 ) The conserved GTPase Gem1 regulates endoplasmic reticulum-mitochondria connections . Proc. Natl. Acad. Sci. USA 108 , 14151 – 14156 Google Scholar Crossref Search ADS WorldCat 31 Lackner L.L. ( 2019 ) The expanding and unexpected functions of mitochondria contact sites . Trends Cell Biol . 29 , 580 – 590 Google Scholar Crossref Search ADS PubMed WorldCat 32 Dolman N.J. , Gerasimenko J.V. , Gerasimenko O.V. , Voronina S.G. , Petersen O.H. , Tepikin A.V. ( 2005 ) Stable Golgi-mitochondria complexes and formation of Golgi Ca(2+) gradients in pancreatic acinar cells . J. Biol. Chem. 280 , 15794 – 15799 Google Scholar Crossref Search ADS PubMed WorldCat 33 Soto-Heredero G. , Baixauli F. , Mittelbrunn M. ( 2017 ) Interorganelle communication between mitochondria and the endolysosomal system . Front. Cell Dev. Biol . 5 , 95 Google Scholar Crossref Search ADS PubMed WorldCat 34 Wong Y.C. , Ysselstein D. , Krainc D. ( 2018 ) Mitochondria-lysosome contacts regulate mitochondrial fission via RAB7 GTP hydrolysis . Nature 554 , 382 – 386 Google Scholar Crossref Search ADS PubMed WorldCat 35 Hamdi A. , Roshan T.M. , Kahawita T.M. , Mason A.B. , Sheftel A.D. , Ponka P. ( 2016 ) Erythroid cell mitochondria receive endosomal iron by a “kiss-and-run” mechanism . Biochim. Biophys. Acta 1863 , 2859 – 2867 Google Scholar Crossref Search ADS PubMed WorldCat 36 Shai N. , Yifrach E. , van Roermund C.W.T. , Cohen N. , Bibi C. , IJlst L. , Cavellini L. , Meurisse J. , Schuster R. , Zada L. , Mari M.C. , Reggiori F.M. , Hughes A.L. , Escobar-Henriques M. , Cohen M.M. , Waterham H.R. , Wanders R.J.A. , Schuldiner M. , Zalckvar E. ( 2018 ) Systematic mapping of contact sites reveals tethers and a function for the peroxisome-mitochondria contact . Nat. Commun . 9 , 1761 Google Scholar Crossref Search ADS PubMed WorldCat 37 Lackner L.L. , Ping H. , Graef M. , Murley A. , Nunnari J. ( 2013 ) Endoplasmic reticulum-associated mitochondria-cortex tether functions in the distribution and inheritance of mitochondria . Proc. Natl. Acad. Sci. USA 110 , E458 – E467 Google Scholar Crossref Search ADS WorldCat 38 McBride H.M. , Neuspiel M. , Wasiak S. ( 2006 ) Mitochondria: more than just a powerhouse . Curr. Biol . 16 , R551 – R560 Google Scholar Crossref Search ADS PubMed WorldCat 39 Pilling A.D. , Horiuchi D. , Lively C.M. , Saxton W.M. ( 2006 ) Kinesin-1 and Dynein are the primary motors for fast transport of mitochondria in Drosophila motor axons . Mol. Biol. Cell 17 , 2057 – 2068 Google Scholar Crossref Search ADS PubMed WorldCat 40 Fransson Å , Ruusala A. , Aspenström P. ( 2006 ) The atypical Rho GTPases Miro-1 and Miro-2 have essential roles in mitochondrial trafficking . Biochem. Biophys. Res. Commun . 344 , 500 – 510 Google Scholar Crossref Search ADS PubMed WorldCat 41 Glater E.E. , Megeath L.J. , Stowers R.S. , Schwarz T.L. ( 2006 ) Axonal transport of mitochondria requires Milton to recruit kinesin heavy chain and is light chain independent . J. Cell Biol. 173 , 545 – 557 Google Scholar Crossref Search ADS PubMed WorldCat 42 Seo J.H. , Agarwal E. , Bryant K.G. , Caino M.C. , Kim E.T. , Kossenkov A.V. , Tang H.Y. , Languino L.R. , Gabrilovich D.I. , Cohen A.R. , Speicher D.W. , Altieri D.C. ( 2018 ) Syntaphilin ubiquitination regulates mitochondrial dynamics and tumor cell movements . Cancer Res. 78 , 4215 – 4228 Google Scholar Crossref Search ADS PubMed WorldCat 43 Kang J.S. , Tian J.H. , Pan P.Y. , Zald P. , Li C. , Deng C. , Sheng Z.H. ( 2008 ) Docking of axonal mitochondria by syntaphilin controls their mobility and affects short-term facilitation . Cell 132 , 137 – 148 Google Scholar Crossref Search ADS PubMed WorldCat 44 Soubannier V. , McBride H.M. ( 2009 ) Positioning mitochondrial plasticity within cellular signaling cascades . Biochim. Biophys. Acta 1793 , 154 – 170 Google Scholar Crossref Search ADS PubMed WorldCat 45 Gonzalez-Licea A. ( 1970 ) Polarization of mitochondria in the absorptive cells of the small intestine of suckling rats. Light and electron microscopy study . Lab Invest . 23 , 163 – 167 Google Scholar PubMed WorldCat 46 Gonzalez-Licea A. ( 1971 ) An ultrastructural study of intestinal mitochondrial morphology during the absorption of various nutrients and water in suckling rats . Lab Invest . 24 , 273 – 278 Google Scholar PubMed WorldCat 47 Al-Mehdi A.B. , Pastukh V.M. , Swiger B.M. , Reed D.J. , Patel M.R. , Bardwell G.C. , Pastukh V.V. , Alexeyev M.F. , Gillespie M.N. ( 2012 ) Perinuclear mitochondrial clustering creates an oxidant-rich nuclear domain required for hypoxia-induced transcription . Sci. Signal . 5 , ra47 Google Scholar Crossref Search ADS PubMed WorldCat 48 Saxton W.M. , Hollenbeck P.J. ( 2012 ) The axonal transport of mitochondria . J. Cell Sci . 125 , 2095 – 2104 Google Scholar Crossref Search ADS PubMed WorldCat 49 Debattisti V. , Gerencser A.A. , Saotome M. , Das S. , Hajnóczky G. ( 2017 ) ROS control mitochondrial motility through p38 and the motor adaptor Miro/Trak . Cell Rep . 21 , 1667 – 1680 Google Scholar Crossref Search ADS PubMed WorldCat 50 Lopez-Domenech G. , Covill-Cooke C. , Ivankovic D. , Halff E.F. , Sheehan D.F. , Norkett R. , Birsa N. , Kittler J.T. ( 2018 ) Miro proteins coordinate microtubule- and actin-dependent mitochondrial transport and distribution . EMBO J. 37 , 321 – 336 Google Scholar Crossref Search ADS PubMed WorldCat 51 Porporato P.E. , Filigheddu N. , Pedro J.M.B. , Kroemer G. , Galluzzi L. ( 2018 ) Mitochondrial metabolism and cancer . Cell Res. 28 , 265 – 280 Google Scholar Crossref Search ADS PubMed WorldCat 52 Aw T.Y. , Jones D.P. , ( 1985 ) ATP concentration gradients in cytosol of liver cells during hypoxia . Am. J. Physiol . 249 , C385 – C392 Google Scholar Crossref Search ADS PubMed WorldCat 53 Aflalo C. , DeLuca M. ( 1987 ) Continuous monitoring of adenosine 5′-triphosphate in the microenvironment of immobilized enzymes by firefly luciferase . Biochemistry 26 , 3913 – 3920 Google Scholar Crossref Search ADS PubMed WorldCat 54 Niethammer P. , Kueh H.Y. , Mitchison T.J. ( 2008 ) Spatial patterning of metabolism by mitochondria, oxygen, and energy sinks in a model cytoplasm . Curr. Biol . 18 , 586 – 591 Google Scholar Crossref Search ADS PubMed WorldCat 55 Jayashankar V. , Rafelski S.M. ( 2014 ) Integrating mitochondrial organization and dynamics with cellular architecture . Curr. Opin. Cell Biol . 26 , 34 – 40 Google Scholar Crossref Search ADS PubMed WorldCat 56 Hollenbeck P.J. ( 1996 ) The pattern and mechanism of mitochondrial transport in axons . Front. Biosci . 1 , d91 – d102 Google Scholar Crossref Search ADS PubMed WorldCat 57 Chada S.R. , Hollenbeck P.J. ( 2004 ) Nerve growth factor signaling regulates motility and docking of axonal mitochondria . Curr. Biol . 14 , 1272 – 1276 Google Scholar Crossref Search ADS PubMed WorldCat 58 Nguyen T.T. , Oh S.S. , Weaver D. , Lewandowska A. , Maxfield D. , Schuler M.H. , Smith N.K. , Macfarlane J. , Saunders G. , Palmer C.A. , Debattisti V. , Koshiba T. , Pulst S. , Feldman E.L. , Hajnoczky G. , Shaw J.M. ( 2014 ) Loss of Miro1-directed mitochondrial movement results in a novel murine model for neuron disease . Proc. Natl. Acad. Sci. USA 111 , E3631 – E3640 Google Scholar Crossref Search ADS WorldCat 59 Liu Y. , Fiskum G. , Schubert D. ( 2002 ) Generation of reactive oxygen species by the mitochondrial electron transport chain . J. Neurochem. 80 , 780 – 787 Google Scholar Crossref Search ADS PubMed WorldCat 60 Jones D.P. , ( 2010 ) Redox sensing: orthogonal control in cell cycle and apoptosis signalling . J. Intern. Med . 268 , 432 – 448 Google Scholar Crossref Search ADS PubMed WorldCat 61 Jelcic M. , Enyedi B. , Xavier J.B. , Niethammer P. ( 2017 ) Image-based measurement of H2O2 reaction-diffusion in wounded zebrafish larvae . Biophys. J . 112 , 2011 – 2018 Google Scholar Crossref Search ADS PubMed WorldCat 62 Niethammer P. , Grabher C. , Look A.T. , Mitchison T.J. ( 2009 ) A tissue-scale gradient of hydrogen peroxide mediates rapid wound detection in zebrafish . Nature 459 , 996 – 999 Google Scholar Crossref Search ADS PubMed WorldCat 63 Lyublinskaya O. , Antunes F. ( 2019 ) Measuring intracellular concentration of hydrogen peroxide with the use of genetically encoded H2O2 biosensor HyPer . Redox Biol . 24 , 101200 Google Scholar Crossref Search ADS PubMed WorldCat 64 Lee J.J. , van de Ven R.A.H. , Zaganjor E. , Ng M.R. , Barakat A. , Demmers J. , Finley L.W.S. , Gonzalez Herrera K.N. , Hung Y.P. , Harris I.S. , Jeong S.M. , Danuser G. , McAllister S.S. , Haigis M.C. ( 2018 ) Inhibition of epithelial cell migration and Src/FAK signaling by SIRT3 . Proc. Natl. Acad. Sci. USA 115 , 7057 – 7062 Google Scholar Crossref Search ADS WorldCat 65 Daiber A. ( 2010 ) Redox signaling (cross-talk) from and to mitochondria involves mitochondrial pores and reactive oxygen species . Biochim. Biophys. Acta 1797 , 897 – 906 Google Scholar Crossref Search ADS PubMed WorldCat 66 Cunniff B. , Benson K. , Stumpff J. , Newick K. , Held P. , Taatjes D. , Joseph J. , Kalyanaraman B. , Heintz N.H. ( 2013 ) Mitochondrial-targeted nitroxides disrupt mitochondrial architecture and inhibit expression of peroxiredoxin 3 and FOXM1 in malignant mesothelioma cells . J. Cell. Physiol. 228 , 835 – 845 Google Scholar Crossref Search ADS PubMed WorldCat 67 Kuroda J. , Ago T. , Matsushima S. , Zhai P. , Schneider M.D. , Sadoshima J. ( 2010 ) NADPH oxidase 4 (Nox4) is a major source of oxidative stress in the failing heart . Proc. Natl. Acad. Sci. USA 107 , 15565 – 15570 Google Scholar Crossref Search ADS WorldCat 68 Shanmugasundaram K. , Nayak B.K. , Friedrichs W.E. , Kaushik D. , Rodriguez R. , Block K. ( 2017 ) NOX4 functions as a mitochondrial energetic sensor coupling cancer metabolic reprogramming to drug resistance . Nat. Commun . 8 , 997 Google Scholar Crossref Search ADS PubMed WorldCat Abbreviations Abbreviations DRP1 dynamin related protein 1 ETC electron transport chain IMM inner mitochondrial membrane MAM mitochondrial-associated ER membrane MDCs mitochondrial-derived compartments MDVs mitochondrial-derived vesicles MECA mitochondria–ER–cortex anchor MEFs mouse embryonic fibroblasts mROS mitochondrial reactive oxygen species OMM outer mitochondrial membrane © The Author(s) 2019. Published by Oxford University Press on behalf of the Japanese Biochemical Society. 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TI - Dynamic regulation of subcellular mitochondrial position for localized metabolite levels
JF - The Journal of Biochemistry
DO - 10.1093/jb/mvz058
DA - 2020-02-01
UR - https://www.deepdyve.com/lp/oxford-university-press/dynamic-regulation-of-subcellular-mitochondrial-position-for-localized-8o0VXi8NnC
SP - 109
VL - 167
IS - 2
DP - DeepDyve
ER -