Cell biology in neuroscience: Architects in neural circuit design: Glia control neuron numbers and connectivity

Glia serve many important functions in the mature nervous system. In addition, these diverse cells have emerged as essential participants in nearly all aspects of neural development. Improved techniques to study neurons in the absence of glia, and to visualize and manipulate glia in vivo, have greatly expanded our knowledge of glial biology and neuron–glia interactions during development. Exciting studies in the last decade have begun to identify the cellular and molecular mechanisms by which glia exert control over neuronal circuit formation. Recent findings illustrate the importance of glial cells in shaping the nervous system by controlling the number and connectivity of neurons.

Introduction

The nervous system is comprised of two main cell types: neurons and glia. Glia represent a heterogeneous population of nervous system cells that associate with neurons and have diverse roles in nervous system development, maintenance, and function. Major glial cell types in vertebrates include astrocytes, microglia, oligodendrocytes, and NG2⁺ oligodendrocyte precursor cells (OPCs) in the central nervous system, and Schwann cells in the peripheral nervous system. Active roles for glia in all major steps of neural development have been described in a variety of model organisms from worms to mammals. Among the first recognized (and perhaps the most well-known) developmental functions of glia is in helping to guide axons to their correct targets. Glia serve as contact-dependent guideposts and sources of numerous attractive and repulsive cues in many circuits (Chotard and Salecker, 2004). It is also now understood that glia serve as stem cells in both embryonic and adult vertebrate nervous systems (Doetsch et al., 1999; Noctor et al., 2001; Seri et al., 2001), in addition to providing important substrates for neuronal migration (Rakic, 1971). In recent years, new functions for glia in synaptogenesis and plasticity have revealed a very active role for glia in neural circuit formation. In this review, we highlight recent insights into active roles for glia during nervous system development, focusing primarily on glial shaping of circuit formation via control of neuron and synapse numbers.

Glia regulate neuron production

A key step in neural development is generating the appropriate number of neurons and glia at the correct time and in the right place. In both invertebrate and vertebrate systems, glial cells have been identified as crucial constituents of neural stem cell niches that modulate neurogenesis by dynamically regulating stem cell proliferation and precursor differentiation in response to a variety of changing developmental time points and functional needs.In Drosophila, neural stem cells called neuroblasts undergo a stereotyped period of quiescence during early larval stages before resuming divisions to give rise to the adult nervous system (Truman and Bate, 1988), and proper transition between these states is crucial for normal nervous system formation (Ebens et al., 1993). As with many other stem cell types, neuroblast exit from quiescence is tied to overall animal development through nutrient signaling. Signals from the larval fat body, which monitors nutrient status in the periphery, are required to activate neuroblasts to reenter the cell cycle (Britton and Edgar, 1998). Two recent studies have shown that glia are required to act as an intermediate to transduce this information to neuroblasts. In response to nutrient-dependent fat body signaling, glia secrete insulin/IGF-like peptides (Drosophila insulin-like peptides [dILPs]). These dILPs activate insulin receptor–PI3K/Akt signaling in neuroblasts, triggering reentry into the cell cycle (Chell and Brand, 2010; Sousa-Nunes et al., 2011). Neuroblast proliferation is significantly delayed in animals lacking dILPs, or when glia are prevented from signaling by inhibiting vesicle trafficking with a mutant form of the fly dynamin gene, shibire. Moreover, glial expression of dILPs is sufficient to induce precocious activation of neuroblasts under normal conditions or bypass the requirement for dietary amino acids in starvation conditions (Chell and Brand, 2010; Sousa-Nunes et al., 2011). Glia, therefore, play a specific role in receiving systemic signals and transmitting them to neural stem cells to modulate their growth. Using glia as an intermediary may allow for more stable and precisely timed regulation of stem cell activation by allowing glia to serve as an integration site for multiple cues, which clearly include nutritional status. Auto-regulation of their own activity also appears to modulate glial control of stem cell activity: there is evidence glia may cell-autonomously modulate their own dILP production through expression of the RNA-binding protein FMRP, and that this regulation is critical for proper timing of neuroblast reactivation (Callan et al., 2012). Thus, glia may not simply relay information, but serve as important integration or filtering sites for multiple signals that impinge on neuroblast biology.Glial cells also play important roles in regulating progenitor cell division and specification in the proliferative zones of embryonic and adult mammalian brains. Microglia derived from myeloid precursors in the yolk sac migrate into the brain as early as E10.5 (Hirasawa et al., 2005; Ginhoux et al., 2010), where they can modulate neural precursor proliferation and neural cell fate specification during embryonic cortical neurogenesis. Evidence from in vitro studies comparing isolated progenitors to those cultured with young microglia suggest that the presence of microglia enhances proliferation and biases progeny toward an astroglial fate (Antony et al., 2011). A novel role for microglia in the negative regulation of neurogenesis has also recently been described. Data from immunostaining of fixed tissue and imaging of microglia in cultured brain slices suggest that young microglia phagocytose cortical precursor cells as cortical neurogenesis nears completion, which may serve as a key mechanism to curb neuron production or reduce neural populations at this time (Cunningham et al., 2013). One intriguing feature of this finding is that microglia appear to engulf cells that lack several standard cell death markers and may even engulf actively dividing progenitors (Cunningham et al., 2013). These observations suggest that microglia may not simply be acting as scavengers, but might be initiating cell death via engulfment. There is evidence that strongly suggests that engulfment by microglia is required to fully execute partially activated developmental cell death programs in post-mitotic Purkinje neurons (Marín-Teva et al., 2004; see next section), and roles for engulfing cells in promoting final execution of apoptosis in target cells have been described in Caenorhabditis elegans (Hoeppner et al., 2001; Reddien et al., 2001). However, the above study goes further, suggesting that microglia somehow decide to engulf and destroy seemingly healthy neural precursors. Defining the molecular mechanisms that govern this choice will be an important focus for future work.Radial glia are a key feature of the developing cortex, long understood to serve as important scaffolds for neuronal migration in the developing embryo. Studies in the last decade have now demonstrated that radial glia cells are in fact neuronal progenitors in the developing brain (Noctor et al., 2001; Tamamaki et al., 2001). This important finding came on the heels of the identification of astrocytes as adult neural stem cells in the mammalian brain (Doetsch et al., 1999). In addition to serving as stem cells, mammalian astrocytes in adult proliferative zones can modulate progenitor division and differentiation through the release of secreted molecules (including FGF2) and contact-dependent mechanisms (such as Eph/ephrin signaling; Morrens et al., 2012). Only astrocytes from proliferative zones are capable of promoting neurogenesis; thus, astrocytes constrain where new neurons are generated in the adult (Song et al., 2002). In addition, astrocytes can also couple neurogenesis to physiological status or injury. For example, IGF-I, normally expressed by neurons, is strongly expressed in astrocytes only after brain injury, and may contribute to injury-induced increases in adult neurogenesis (Yan et al., 2006).These exciting new studies of neural stem cells and neurogenesis in different species and at different developmental stages demonstrate conserved and wide-ranging roles for glia as regulators of neuron production. Through mechanisms that remain to be defined, glia appear to serve as sites of integration that allow dynamic modulation of neurogenesis in response to changing physiological needs during development and after injury or disease in the adult. Understanding the cellular and molecular mechanisms by which glia monitor animal physiology and modulate neurogenesis should have important implications for the understanding and treatment of many neurodevelopmental disorders and neurodegenerative diseases.

Glia in developmental programmed cell death

Overproduction, followed by programmed cell death (PCD) of excess neurons, is a well-described and conserved feature of nervous system development across phyla that is designed to ensure a sufficient number of neurons are initially formed to accomplish neural circuit construction (see Dekkers et al., in this issue). Competition between neurons for limited environmental and target-derived trophic factors results in cell death of extraneous or weakly connected neurons (Levi-Montalcini, 1987). PCD most often occurs via apoptosis followed by phagocytosis of the cellular debris by microglia, the resident immune cells and phagocytes of the central nervous system. A number of recent studies in the rodent have challenged the view that microglia act as passive scavengers of cellular debris. Rather, accumulating evidence now demonstrates an active role for microglia and innate immune mechanisms in promoting developmental PCD (Fig. 1).Open in a separate windowFigure 1.Microglia promote cell death and clearance of neuronal debris in the developing brain. (A) Model of the role of microglia in promoting neuronal programmed cell death in the developing cerebellum and hippocampus. Caspase-3+ neurons that have initiated apoptosis express as-yet-unknown “kill me/eat me” signals that recruit microglia to engulf them. Contact between dying neurons and microglia expressing the integrin CD11b and the immunoreceptor DAP12 triggers production of reactive oxygen species (including O₂^.−) in microglia, which is locally released to promote completion of apoptosis in the dying neuron. CD11b and DAP12 are required in microglia for efficient ROS production and for normal levels of PCD, but it remains unclear whether CD11b/DAP12 act as a receptor for one of the elusive neuronal signal(s), or if other receptors indirectly activate this pathway. (B) Microglia are also required for the clearance of cellular debris generated by PCD. The neuronal signals and specific microglial receptors that mediate corpse recognition and phagocytosis also remain unknown. Proper clearance of corpses is required to prevent inflammation.Close association or partial engulfment of intact but caspase-3–expressing Purkinje cells and microglia is seen in fixed tissue and in cultured cerebellar slices from postembryonic day 3 (P3) mice, a time corresponding to high levels of Purkinje cell PCD in vivo (Marín-Teva et al., 2004). At this stage, microglia were observed to be amoeboid in shape, which is characteristic of phagocytic function. Despite the fact that many of the Purkinje cells at this stage are already positive for activated caspases, depletion of microglia from the slices using clodronate containing liposomes (which target phagocytic cells) dramatically increases the number of surviving Purkinje cells after 3 days in vitro (Marín-Teva et al., 2004). These data fit a model proposed by PCD studies in C. elegans that shows engulfment by phagocytes is not simply required for debris clearance, but is itself required for the completion of cell death, even in cells already expressing activated caspases (Hoeppner et al., 2001; Reddien et al., 2001).In the innate immune response, engulfing cells can release reactive oxygen species (ROS) in a process called “respiratory burst” to aid in killing pathogens (Chanock et al., 1994). Using pharmacology, Marín-Teva et al. (2004) found evidence that microglia produce such respiratory bursts upon engulfment of Purkinje cells that contributed to their death. Subsequent studies demonstrated similar requirements for microglia in mediating PCD in the developing hippocampus and identified two immune molecules expressed in microglia that have been co-opted from the innate immune system for use in developmental PCD (Wakselman et al., 2008). In peripheral immune cells, the cell surface molecules CD11b and DAP12 work together to target pathogens for destruction using ROS (Mócsai et al., 2006). In the developing hippocampus, CD11b and DAP12 are expressed exclusively by microglia, and examination of DAP12 or CD11b mutant mice showed reduced microglial superoxide production along with significantly reduced PCD in the hippocampus (Fig. 1 A; Wakselman et al., 2008). In both the cerebellum and hippocampus, caspase activation is still assumed to be cell-autonomously induced, and although greatly reduced, some PCD is still observed in the absence of microglial signaling. These data suggest that cell-autonomous mechanisms are capable of both inducing PCD and carrying it to completion in some cases, but that microglia play an important role in a majority of cases. Thus, microglia appear to have two important roles in developmental PCD. First, they can act to promote cell death via production of reactive oxygen species (and potentially additional mechanisms) to ensure the appropriately high levels of cell death that are required for normal development (Fig. 1 A). In addition, microglia act as phagocytes to clear the debris PCD generates (Fig. 1 B). It is assumed that dying neurons produce “kill me” and/or “eat me” signals that help recruit microglia and initiate CD11b/DAP12-dependent ROS production and engulfment, but these putative signals and their microglial receptors remain unknown.Whether microglia play such an active role in cell death outside of development is unclear. In uninjured adult hippocampus, for example, microglia lack DAP12 (Wakselman et al., 2008) and have very low levels of CD11b (Sierra et al., 2010), yet there are high levels of cell death among newly born neurons in the subgranular zone of the hippocampus (a site of adult neurogenesis; Sierra et al., 2010). This suggests that microglia may not play the same active role in driving apoptosis of the newly born neurons, but this remains to be tested directly. Interestingly, engulfment of these cells in the mature hippocampus is performed by the processes of ramified (resting) microglia that efficiently phagocytose dying neurons without any morphological signs of activation (Sierra et al., 2010). Engulfment earlier in development is generally performed by amoeboid-like microglia, so this morphological difference may suggest the molecular mechanisms engaged may also be distinct at different stages. On the other hand, microglial activity during PCD at all stages is clearly distinguished from microglial response to injury or infection, which causes inflammation. Rapid engulfment of neurons undergoing PCD is considered immunologically silent—actually preventing inflammation that might be caused by unattended cell corpses.

Glia control connectivity by mediating developmental axon pruning

During development neurons often make inappropriate, excessive, or transient connections that must be eliminated as circuits mature. Though this process sometimes results in developmental cell death, selective pruning of axonal and dendritic branches is an important alternative method of neural circuit refinement and remodeling. Glia play key roles in two of the most well-characterized examples of developmental axon pruning—mammalian neuromuscular junction (NMJ) refinement and fly mushroom body γ-neuron remodeling.In early development, mammalian NMJs are innervated by multiple motor neuron axons. Activity-dependent competition between the axonal branches innervating the same NMJ eventually results in exactly one “winning” axon that maintains connectivity, while all the other “losing” branches are eliminated (Colman et al., 1997; Walsh and Lichtman, 2003). Detailed analysis of branch elimination using confocal and electron microscopy has revealed that losing axons retract from the NMJ by shedding pieces of themselves (termed axosomes) in a distal-to-proximal manner (Bishop et al., 2004; Song et al., 2008). Individual axon branches are ensheathed by perisynaptic Schwann cells—a specialized subtype of glia cell in the peripheral nervous system—before and during retraction. Shed axosomes are engulfed into the ensheathing perisynaptic Schwann cell where they undergo lysosomal degradation in the Schwann cell cytoplasm (Song et al., 2008). Thus, Schwann cells appear to play an essential role in at least the clearance of axonal debris, a feature that appears to be conserved at the fly NMJ (Fuentes-Medel et al., 2009).An unresolved question is whether axosomes are shed cell-autonomously by the retracting axon, or whether the ensheathing Schwann cell may contribute to the formation or “pinching off” of these structures in the losing axon branch. Some imaging evidence reveals thin Schwann cell processes around budding axosomes, suggesting glia might play an active role in axosome formation and/or shedding as they are engulfed (Bishop et al., 2004). Demonstrating such an active role is technically challenging due to the requirement of Schwann cell ensheathment for axon survival (Koirala et al., 2003). Schwann cell ablation does result in loss of nerve terminals, suggesting that retraction can take place in the absence of glia, but whether retraction in this situation is similar to activity-induced pruning is unclear (Reddy et al., 2003)—perhaps in the absence of Schwann cells axon terminals simply degenerate due to lack of support. Experiments in which Schwann cell endocytosis or membrane dynamics can be acutely disrupted during dual-color live imaging will likely be required to definitively resolve this question. Moreover, such an active role in eliminating weak synapses should require that Schwann cells are capable of discriminating and evaluating the relative synaptic strength of multiple inputs so that they target the correct input for elimination. A recent study using simultaneous glial calcium imaging and neuronal electrophysiological recording demonstrates that perisynaptic Schwann cells are capable of “sensing” the relative activity levels of each synapse at a multiply innervated NMJ. An individual PSC contacting two NMJs displays stronger calcium responses to “stronger” than to “weaker” synapses, an effect mediated by purinergic receptors on the glial cells (Darabid et al., 2013). If and how this responsiveness might affect synaptic competition or branch retraction awaits future studies in which glial responses are perturbed and the effects on innervation determined.In Drosophila, axon pruning during metamorphosis is important for the remodeling required to form mature adult circuits. Mushroom body γ-neurons prune their larval axons during early metamorphosis, followed by formation of new adult-specific connections as metamorphosis continues (Fig. 2; Lee et al., 1999). In addition to many cell-autonomous requirements for proper axon pruning, there are multiple essential roles for neighboring glia in mediating axon removal in this circuit. Initiation of pruning is triggered by a pulse of the molting hormone ecdysone just before metamorphosis (Lee et al., 2000). γ-Neuron responsiveness to this critical signal is controlled by surrounding cortex and astrocyte-like glia, which express the TGF-β ligand Myoglianin (Myo) during late larval stages (Awasaki et al., 2011). Glial-derived Myo must activate TGF-β signaling within γ-neurons to up-regulate expression of the ecdysone receptor B1 isoform (EcR-B1) so that γ-neurons can receive the hormonal signal that will trigger pruning (Zheng et al., 2003; Awasaki et al., 2011). Myo therefore represents perhaps the first glial-derived factor required for neurons to become competent to prune their axons (Fig. 2 A).Open in a separate windowFigure 2.Developmental pruning of mushroom body γ-neuron axons requires glia. (A) Before metamorphosis, cortex and astrocyte-like glia secrete the TGF-β ligand Myoglianin, which acts through γ-neuron baboon receptors to up-regulate expression of the ecdysone receptor EcR-B1. This makes γ-neurons competent to respond to the ecdysone pulse that will initiate pruning. (B) As metamorphosis begins, glia infiltrate the mushroom body neuropil. This infiltration is dependent on the glial cell surface receptor Draper. Arrival of glia coincides with axon blebbing and fragmentation, but it remains uncertain whether glia actively promote this fragmentation. (C) Glia are required for clearance of axonal debris. Recognition and phagocytosis of the debris is also mediated by the glial cell surface receptor Draper.γ-Neuron axons are pruned by local degeneration in which specific axonal branches fragment in place and are subsequently cleared by glia cells in a process primarily mediated by the glial cell surface receptor Draper (Fig. 2, B and C; Watts et al., 2003, 2004; Awasaki and Ito, 2004; Awasaki et al., 2006). Although glial clearance and degradation of axonal debris is required for complete pruning, it remains unclear if engulfing glia play active roles in promoting degeneration and fragmentation of pruned axons, or whether they simply respond to and clear cell-autonomously fragmented axons. Preventing glial infiltration by blocking membrane dynamics strongly suppresses axon pruning, providing support for an active role for glia in driving pruning (Awasaki and Ito, 2004). However, these results are inconclusive because the experiments lacked sufficient resolution to distinguish whether axons had been broken down but not cleared, or remained completely intact. Additional evidence in support of an active role is that glia infiltrate the mushroom body lobes before any observable fragmentation and can even infiltrate if fragmentation is suppressed via genetic mutations (Watts et al., 2004). Subsequent work showed that glial infiltration is dependent on the engulfment receptor Draper (Awasaki et al., 2006). Draper is believed to recruit engulfing cells to dying cells by responding to an unidentified “eat me” signal (Zhou et al., 2001; MacDonald et al., 2006). The requirement of Draper for infiltration, therefore, suggests glia are responding to axon-derived signals that may be generated independently of, or before, fragmentation. Microtubule disorganization occurs before fragmentation and can occur even in the absence of glial infiltration (Awasaki et al., 2006), suggesting that axons are already committed to pruning before visible fragmentation. Together, these findings suggest that initiation of axon degeneration (as defined by microtubule breakdown) may be cell-autonomous to neurons, but does not rule out a role for glia in driving fragmentation in addition to engulfment of debris. Live imaging experiments taking advantage of new techniques to independently label and genetically manipulate engulfing glia and mushroom body γ-neurons with single-cell resolution (Lai and Lee, 2006; Potter et al., 2010) will likely be necessary to fully resolve the role of glia in this process. If glia are indeed promoting axonal degeneration, the next exciting question would be—how do they choose which axons to destroy?

Glia promote synapse formation

A key step in the formation of a functional nervous system is the establishment of appropriate synaptic connections between neurons. Unraveling how neurons make appropriate synaptic connections has been a major focus of neurodevelopment research for decades, and we now understand that synaptogenesis is a multistep process including recognition and selection of appropriate targets, assembly and localization of synaptic components, and stabilization of appropriate connections. Research in the last decade has identified important roles for glia in many of these steps.An important breakthrough was the development of methods to isolate and maintain purified neurons in culture in the absence of glia. Pfrieger and Barres (1997) developed a technique to purify retinal ganglion cell (RGC) neurons and keep them healthy in culture without glia. Intriguingly, they found that purified RGCs cultured alone formed sevenfold fewer functional excitatory synapses than RGCs cultured with astrocyte feeder layers as assayed by electrophysiological recordings, immunostaining for synaptic protein localization, and EM analysis (Pfrieger and Barres, 1997; Nägler et al., 2001; Ullian et al., 2001). Subsequent in vitro studies with other neuronal subtypes demonstrated that these findings were not just specific to rat RGCs (Peng et al., 2003; Ullian et al., 2004; Cao and Ko, 2007). Hints that astrocytes might play similar synaptogenic roles in vivo first came from observations that timing of synaptogenesis between RGCs and their in vivo targets correlated with glial infiltration of the area, even though RGC axons had reached their target region several days prior (Ullian et al., 2001).In culture, the synapse-promoting effects of glia did not require physical contact between the glia and neurons, implicating secreted factors as the primary mediators of glia-induced synaptogenesis (Fig. 3). Fractionation of astrocyte-conditioned media (ACM) was used to identify thrombospondins (TSPs)—extracellular matrix proteins with known roles in cell adhesion—as key glial-derived synaptogenic factors (Fig. 3 A; Christopherson et al., 2005). Purified TSP-1 or TSP-2 mimicked ACM to promote structural synapse formation, whereas depleting TSP-2 from ACM abrogated its synaptogenic effects on cultured neurons. With a specific molecule in hand, researchers could now ask whether a glial-derived molecule was required for synaptogenesis in vivo. Christopherson et al. (2005) showed that TSPs are expressed in astrocytes in vivo when synapses are forming and colocalize with synapse-associated astrocyte processes. Importantly, mice mutant for both TSP-1 and TSP-2 had significantly fewer synapses than wild-type animals (∼40% reduction at P8), demonstrating a requirement for an astrocyte-derived factor for normal synapse development in vivo (Christopherson et al., 2005).Open in a separate windowFigure 3.Model of astrocyte–neuron interactions during synapse formation and maturation. (A) Astrocytes secrete several factors to promote structural synapse formation. Astrocytes secrete thrombospondins (TSPs), which act through α2–δ1 calcium channel subunit/gabapentin receptors to drive formation of structurally intact glutamatergic synapses. The exact mechanism by which TSP binding to α2–δ1 promotes adhesion and structural synapse formation is unknown. Astrocytes also secrete hevin and SPARC. Hevin also promotes structural synapse formation, whereas SPARC antagonizes this function. Competition between hevin and SPARC for a common unknown binding partner on neurons may explain the antagonism. The synapses formed in response to TSPs or hevin are structurally intact with docked vesicles and PSD-95 correctly localized; however, they lack surface expression of AMPARs at the postsynapse and are therefore not fully functional. (B) Other astrocyte-secreted factors influence surface expression and clustering of glutamatergic AMPARs. Glypicans secreted from astrocytes act through an unidentified receptor to promote surface expression and clustering of AMPARs during development. Activity can also induce astrocytes to secrete SPARC, which can perturb integrin interactions with surface AMPARs, leading to decreased surface expression of AMPARs in response to excess activity.In an elegant follow-up study, Eroglu et al. (2009) identified the α2–δ1 calcium channel subunit/gabapentin receptor as the neuronal receptor for glial TSPs, which bind the receptor through their EGF-like domains. α2–δ1 colocalizes with both pre- and postsynaptic puncta, and in vitro overexpression experiments suggest it functions postsynaptically. The intracellular mechanisms by which TSP/α2–δ1 signaling promotes synapse assembly remains an open question, though α2–δ1 modulation of calcium channel function is not likely to be involved because perturbation of calcium channel expression or function is unable to interfere with TSP- or astrocyte-induced synaptogenesis (Eroglu et al., 2009).The extracellular matrix protein hevin is a second astrocyte-derived factor capable of inducing structural synapse formation in vitro (Fig. 3 A). Although purified hevin works just as well as TSPs at inducing glutamatergic synapses in culture, depletion of TSPs from ACM completely abolishes the synaptogenic activity of ACM despite the presence of hevin (Christopherson et al., 2005; Kucukdereli et al., 2011), implying that there might be ACM components that can antagonize hevin function. This inhibitory factor was identified as SPARC (secreted protein acidic and rich in cysteine), a hevin homologue that is also highly expressed in ACM. SPARC selectively interferes with hevin’s synapse-promoting ability, likely by competing with hevin for an as-yet-unidentified common binding partner on neurons. Hevin and SPARC are coexpressed in vivo, with peak expression in the superior colliculus at a time when RGCs are actively forming synapses. hevin mutant mice have reduced, and SPARC mutant mice have increased, RGC-collicular synapses, supporting antagonistic roles for these proteins in vivo. The overall positive effect of synaptogenesis in this circuit during development is likely due to a high hevin/SPARC ratio. (This ratio is low in ACM.) Expression of hevin and SPARC is maintained into adulthood, unlike TSPs (Christopherson et al., 2005), indicating that hevin and SPARC might also play changing roles in synapse maintenance and plasticity throughout life, potentially through dynamic regulation of their relative levels.Although TSPs and hevin play critical roles in promoting synapse formation, these molecules are only capable of promoting structural assembly but not functional maturation of synapses (Christopherson et al., 2005; Eroglu et al., 2009; Kucukdereli et al., 2011). Synapses formed in response to TSPs and hevin are “silent” synapses: synapses that look ultrastructurally normal, but lack postsynaptic AMPA glutamate receptors (AMPARs), rendering them incapable of neurotransmission. Astrocytes or ACM are sufficient to induce the formation of functional synapses, as well as strengthen newly formed synapses by increasing the amplitude and frequency of miniature excitatory postsynaptic potentials (mEPSPs; Ullian et al., 2001). Thus, there must be additional glia-derived factors that work in conjunction with TSPs and hevin to specifically instruct the addition of AMPARs at the postsynapse. This was an outstanding question in the field until the glypicans Gpc-4 and Gpc-6, which are part of the heparan sulfate proteoglycan family, were identified as ACM factors that fulfill this role (Fig. 3 B; Allen et al., 2012). When supplied to isolated neurons in conjunction with TSPs, Gpc-4 and Gpc-6 were each sufficient to increase synaptic activity in cultured RGCs, primarily by increasing surface expression and clustering of postsynaptic AMPAR GluA1 subunits. Synapse function is impaired in gpc-4 knockout mice, as evidenced by reductions in the amplitude of mEPSPs in hippocampal slices, supporting an in vivo role for glypicans in promoting functional synapse formation (Allen et al., 2012). The mechanism by which glypicans induce surface expression and clustering of GluA1 AMPARs is still unknown. In other contexts, glypicans have roles in concentrating morphogens to enhance signaling pathways, but examination of several glypican interactors (including FGFs, Sonic hedgehog, and Wnts) failed to show roles for these molecules in AMPAR regulation. This suggests that Gpc-4 and Gpc-6 might act directly on a postsynaptic receptor to mediate their effects. Identification of the synaptic glypican receptor or binding partner, and of additional molecules that regulate the surface expression and clustering of AMPAR subunits, will be important for determining how astrocytes contribute to synaptic maturation and the plasticity of neural circuits in vivo.It is unlikely the above-described molecules represent the sum of pro-synaptogenic factors supplied by glia. For example, recent work on inhibitory GABAergic synapse formation indicates the effect of TSPs is specific to glutamatergic synapses, having virtually no effect on inhibitory synapse development. Nonetheless, ACM is required for normal inhibitory synapse development (Hughes et al., 2010); thus, ACM must contain additional synaptogenic factor(s) that can affect inhibitory synapses that have yet to be identified. In addition, astrocytes are not the only glial cell type to support synapse formation—peripheral glia also promote NMJ synapse formation and growth in vertebrates and Drosophila through secretion of TGF-β ligands. Schwann cell–derived TGF-β1 can drive synapse formation in Xenopus motor neuron/muscle co-cultures by acting on presynaptic neurons to increase expression of a protein called agrin, which is required to cluster acetylcholine receptors at neuron/muscle contact sites (Feng and Ko, 2008). In Drosophila, the role of glial TGF-β signaling on synapse growth is distinct. In this system, glia express the TGF-β ligand Maverick (Mav), which acts on postsynaptic muscle to modulate muscle release of a distinct TGF-β ligand (Glass bottom boat) that acts directly on neurons to drive synaptic growth (Fuentes-Medel et al., 2012). Thus, although a synaptogenic role for glia cells is highly conserved, diverse mechanisms exist to drive this function that may be specific to species, glial cell type, or synapse type.Several important questions emerge from these studies. First, are the signals discussed above nonspecific permissive signals that allow synapses to form with specificity being driven by entirely distinct factors, or are they capable of selectively driving the formation of specific, appropriate synapses in vivo? Implication of mainly secreted factors as mediators of synapse promotion from in vitro studies suggests the former. There are some studies, however, that have demonstrated close association and dynamic interactions of glial processes and developing synaptic spines that suggest that glia can and do act in synapse-specific ways as well. Whether similar or different molecular mechanisms are involved in these interactions remains to be determined.A second question is how glial synapse promotion is regulated and modulated during and after development. The effects of TSPs and hevin are robust in vitro, and aside from controlling the general timing of expression or the coexpression SPARC and hevin (mechanisms that seem relatively imprecise and slow), it remains unclear how synapse formation is regulated, particularly in vivo both during development and throughout life. Most glial cells are capable of sensing and responding to neural activity, but how activity affects the production or secretion of synaptogenic factors (particularly before synapse formation) has not been fully resolved. It has been recently reported that SPARC is up-regulated in astrocytes in response to neural activity. In this context though, SPARC seems to have a distinct primary function from inhibiting hevin—negative regulation of AMPAR levels at synapses via inhibition of neuronal β-integrin function (Fig. 3 B; Jones et al., 2011). Thus, SPARC appears to play at least two distinct roles in synapse development and plasticity at different times. At early stages, SPARC limits the number of synapses by abrogating hevin function before synapse assembly. After synapses have formed, SPARC can refine synaptic strength by interfering with integrin localization and function that normally serves to stabilize AMPARs at the membrane. Thus, production of SPARC in response to activity can indirectly limit AMPAR clustering at highly active synapses (Jones et al., 2011). Glial regulation of synapse formation, function, and plasticity throughout life is likely to be a rich and fruitful area of future study.Finally, we need to understand how these astrocyte-derived pro-synaptogenic factors function in vivo to coordinate synapse formation and plasticity in different brain regions. This is clearly a complex issue—for example, TSP knockout mice have an ∼40% reduction in cortical synapses and hevin knockout mice show an ∼35% reduction in collicular synapses. These observations indicate that there are likely additional partially redundant glial-derived factors yet to be discovered. Determining how these glial signals interact or synergize with the molecular cues required between pre- and postsynaptic neurons in vivo will be an important future goal.

Glia prune exuberant synaptic connections

Activity-dependent mechanisms ensure that appropriate connections are strengthened while inappropriate or weak connections are eliminated, but the cellular and molecular mechanisms that underlie experience-dependent synapse elimination are largely unknown. A flurry of research over the last several years has identified microglia as key mediators of developmental synaptic pruning by phagocytosing excessive presynaptic inputs.Microglia were once assumed to be relatively quiescent in healthy brains, but advances in live imaging techniques have revealed “resting” microglia to be highly active cells that constantly extend and retract their processes to make contact with neighboring neurons, glia, and synapses (Davalos et al., 2005; Nimmerjahn et al., 2005; Wake et al., 2009). Moreover, microglia activity changes in response to perturbations in neurotransmission, indicating that microglia might be actively monitoring local neural activity, perhaps playing a role in activity-dependent plasticity. High-resolution imaging studies combining live imaging with electron microscopy soon revealed that microglia made frequent contact with dendritic spines during periods of plasticity and remodeling. This contact has consequences on spine morphology and stability with microglia contact frequently predicting the loss of individual spines (Tremblay et al., 2010).Evidence of synaptic elements contained within microglia cytoplasm combined with the known role of microglia as phagocytes, and a report implicating the classical complement cascade with synaptic pruning provided highly suggestive evidence that microglia were involved in activity-dependent synapse elimination via phagocytosis (Stevens et al., 2007; Tremblay et al., 2010; Paolicelli et al., 2011; Schafer et al., 2012). In an elegant study, Schafer et al. (2012) used the mammalian retinogeniculate system to clearly demonstrate the in vivo role for microglia in mediating activity-dependent synaptic pruning that relies on signaling from the classical complement system. This circuit undergoes a well-characterized period of robust activity-dependent synaptic elimination during postnatal development with a clear readout. RGC axons from both eyes form synapses in the lateral geniculate nucleus (LGN) of the thalamus. Due to exuberant connectivity, distinct ipsilateral and contralateral eye domains are poorly defined until approximately postnatal day 5, at which point activity-dependent synaptic pruning results in clearly segregated eye-specific domains (Fig. 4 A). Injecting unique fluorescent anterograde tracers in each eye labels RGC axons and synapses independently, allowing a clear readout of the success of activity-dependent synaptic pruning (Huberman et al., 2008). GFP-expressing microglia in the LGN were closely associated with synapses, and were often found with tracer-labeled presynaptic membrane within their processes and soma during the period (∼P5) of robust synaptic pruning, similar to previous findings in the developing cortex and hippocampus (Tremblay et al., 2010; Paolicelli et al., 2011).Open in a separate windowFigure 4.Microglia refine circuits by complement-mediated phagocytosis of weak synapses. Schematic of retinogeniculate connectivity in wild-type (A) and complement receptor CR3 knockout (B) animals. Inputs from each eye are completely segregated in the LGNs of wild-type animals (A, separate blue and green regions). Synaptic debris from inappropriate connections is observed within local microglia (brown cells) during the pruning process. In CR3 knockout animals (B), segregation of inputs in the LGN is incomplete, resulting in regions with overlapping inputs from both eyes (blue/green regions). Significantly less synaptic debris is observed in microglia in these animals. (C) Proposed model for complement-dependent synaptic pruning by microglia. Weak synapses are tagged for elimination by recruitment of the classical complement molecule C1q. C1q can recruit and activate C3, which then serves as a ligand for the CR3 receptor (expressed exclusively by microglia) to recruit microglia to clear the synapse. How C1q and C3 are localized to and activated at weak synapses is still unknown.To test whether microglia were indeed ingesting synapses destined for elimination, the authors injected one eye with tetrodotoxin (TTX) to create a strong imbalance of activity between the RGCs from each eye. As expected, the authors found significantly more engulfed synapses that originated from the TTX-injected eye as compared with the control-injected eye, indicating that microglia were pruning synapses in an activity-dependent manner. A wealth of anatomical data supported the idea that microglia engulf weak synapses for elimination, but what were the molecular mechanisms that controlled this engulfment? A good candidate came from an earlier study that identified the classical complement components C1q and C3 as synaptically localized molecules required for activity-dependent pruning of retinogeniculate synapses (Stevens et al., 2007). As microglia are the only cell type in the brain that express the complement receptor CR3, it was hypothesized that binding between synaptic C3 and microglial CR3 might mediate synapse–microglia interactions required for engulfment (Fig. 4 C). To test this hypothesis, Schafer et al. (2012) examined CR3 knockout mice and found that CR3-deficient microglia showed an ∼50% reduction in phagocytosed presynaptic material. This in turn resulted in incomplete pruning, as evidenced by incomplete segregation of eye-specific domains in the LGN, and demonstrated a clear role for complement signaling in microglia pruning of synapses (Fig. 4 B).Among the key questions this work raises is how weak synapses are tagged for removal by the complement system. How the diffusible molecules C1q and activated C3 are localized to or activated at specific synapses is unclear, so the identification of complement system binding partners, inhibitors, and activators at synapses and determining how they are regulated by neuronal activity will be an important focus of future studies (Stevens et al., 2007). In addition, although complement signaling plays a major role in synaptic pruning, loss of complement signaling does not completely prevent pruning, causing only an ∼50% reduction. Thus, it is likely that other molecules and signaling pathways contribute to microglial synaptic pruning. One candidate pathway suggested by studies in fly is that of Draper (Megf10 in mouse), which has been shown to mediate developmental axon pruning (Awasaki et al., 2006) and can function in peripheral glia and muscle to mediate engulfment of shed synapses at the NMJ (Fuentes-Medel et al., 2009). This study from Drosophila also elegantly demonstrates the importance of synaptic clearance by glia. Failure to clear synaptic debris leads to severe defects in synapse growth at the NMJ, indicating that the elimination of synaptic debris by glia is an important event required for synaptic plasticity (Fuentes-Medel et al., 2009).In addition to synaptic pruning by microglia, other glial subtypes play important roles in synaptic plasticity. For example, as discussed above, astrocyte-secreted factors including SPARC can influence synaptic plasticity. SPARC expression in the postnatal hippocampus is regulated by activity. SPARC affects clustering and stabilization of glutamate receptor subunits at hippocampal synapses during postnatal development, and is required for normal synaptic plasticity during this time (Jones et al., 2011). Visual cortex plasticity is also strongly affected by glia, for example implantation of immature astrocytes into the brains of older animals can restore ocular dominance plasticity (Müller and Best, 1989). Maturation of oligodendrocytes might also affect the duration of ocular dominance plasticity via releasing factors that bind to neuronal Nogo receptors, as knockout of these receptors can lead to an extension of the plastic period in mice (McGee et al., 2005). These are just a few of the contributions of different glial subtypes to activity-dependent plasticity throughout development (and into adulthood) that have been described thus far. Continued work in this field should reveal many more unexpected roles for glia in modulating plasticity and activity in neural circuits.

Conclusions and future directions

There has been remarkable progress in our understanding of how glia function in the developing nervous system to shape neuronal connectivity. In particular, a number of cellular and molecular mechanisms by which glia regulate neural stem cell biology, and neural fate, survival, and connectivity have begun to be elucidated, but there is undoubtedly much more for us to learn about these remarkable cells. Exciting and fundamental questions about neuron–glia interactions await exploration and molecular or functional insight. For example, determining whether synaptogenic signals from astrocytes are perceived as permissive or instructive will be important, and if they are instructive for specific connections, how is specificity embedded in the message? In the context of neural circuit refinement we need to understand cell–cell communication events and how they are integrated with circuit function. How are “loser” synapses that need to be eliminated molecularly tagged? How does neuronal activity regulate glial pruning function or synaptic tagging during synapse elimination? Are microglia the only glia pruning synapses in mammals? What are the functional consequences of failed pruning in different brain regions? More broadly, we must explore how neural activity shapes glial developmental events, or neuron–glia interactions. It is well known that many types of glia express neurotransmitter receptors that could allow them to “listen in” on neuronal activity if they are in close contact with synapses. What are these used for? Is this a primary way that neurons signal to glia? If so, how does neurotransmitter signaling to glia shape their biology?In contrast to the growing list of astrocyte and microglial functions during embryonic and postnatal development, relatively little is known about specific roles for oligodendrocytes or their precursors (NG2⁺ cells) in the developing nervous system. Oligodendrocytes mediate myelination during the latest stages of development, though the cellular interactions between axons and glia that mediate myelination are still largely unknown. In vivo ablation studies indicate that loss of oligodendrocytes from the developing cerebellum near the onset of myelination disrupts overall cellular architecture and connectivity and alters the gene expression patterns of neurons, suggesting there may be a number of as-yet-identified functions for these cells during development (Mathis et al., 2003; Doretto et al., 2011). Intriguingly, NG2⁺ OPCs receive synapses from neurons during development (Bergles et al., 2000). Why these cells receive direct synaptic input is unclear, as is the exact function of NG2⁺ OPCs, but they are seemingly poised to respond rapidly to neural activity via their direct synaptic connections and generate new oligodendrocytes (De Biase et al., 2010; Mangin et al., 2012). Determining what these cells do in the developing and mature brain is a major question for the field in the future.Finally, although most communication between neurons and glia is likely bi-directional, many of our current models have elucidated only one half of the conversation. In several examples discussed above we have some knowledge of a glial-derived molecule, but many receptors and downstream signaling pathways remain to be identified. It is also notable that the expression of many glial-derived signals often changes over developmental time (e.g., TSPs and Gpcs). Is this the result of intrinsic changes in glial pro-synaptic potential, or is it the result of feedback signaling from neurons? Misregulation of several of the processes and/or molecules discussed above that modulate neural circuit construction are associated with disease. Understanding these functions more deeply and how they are altered in disease may be crucial for generating new therapeutic strategies to program appropriate levels of neural circuit plasticity.     

Two-photon uncaging: New prospects in neuroscience and cellular biology

An uncaging process refers to a fast and efficient release of a biomolecule after photochemical excitation from a photoactivatable precursor. Two-photon excitation produces excited states identical to standard UV excitation while overcoming major limitations when dealing with biological materials, like spatial resolution, tissue penetration and toxicity and has therefore been applied to the uncaging of different biological effectors. A literature survey of two-photon uncaging of biomolecules is described in this article, including applications in cellular- and neurobiology.     

Cell biology in neuroscience: Cellular and molecular mechanisms underlying axon formation,growth, and branching

Proper brain wiring during development is pivotal for adult brain function. Neurons display a high degree of polarization both morphologically and functionally, and this polarization requires the segregation of mRNA, proteins, and lipids into the axonal or somatodendritic domains. Recent discoveries have provided insight into many aspects of the cell biology of axonal development including axon specification during neuronal polarization, axon growth, and terminal axon branching during synaptogenesis.

Introduction

Axon development can be divided into three main steps: (1) axon specification during neuronal polarization, (2) axon growth and guidance, and (3) axon branching and presynaptic differentiation (Fig. 1; Barnes and Polleux, 2009; Donahoo and Richards, 2009). These three steps are exemplified during neocortical development in the mouse: upon neurogenesis, newly born neurons engage long-range migration and polarize (Fig. 1, A and B) by adopting a bipolar morphology with a leading and a trailing process (Fig. 1 C). During migration (approximately from embryonic day [E]11 to E18 in the mouse cortex), the trailing process becomes the axon and extends rapidly while being guided to its final destination (lasts until around postnatal day [P]7 in mouse corticofugal axons with distant targets like the spinal cord; Fig. 1, D–F). Finally, upon reaching its target area, extensive axonal branching occurs during the formation of presynaptic contacts with specific postsynaptic partners (during the second and third postnatal week in the mouse cortex; Fig. 1, G–I). Disruption of any of these steps is thought to lead to various neurodevelopmental disorders ranging from mental retardation and infantile epilepsy to autism spectrum disorders (Zoghbi and Bear, 2012). This review will provide an overview of some of the cellular and molecular mechanisms underlying axon specification, growth, and branching.Open in a separate windowFigure 1.Axon specification, growth, and branching during mouse cortical development. Three stages of the development of callosal axons of cortical pyramidal neurons from the superficial layers 2/3 of the somatosensory cortex in the mouse visualized using long-term in utero cortical electroporation. For this class of model axons, development can be divided in three main stages: (1) neurogenesis and axon specification, occurring mostly at embryonic ages (A–C); (2) axon growth/guidance during the first postnatal week (D–F); and (3) axon branching and synapse formation until approximately the end of the third postnatal week (G–I). A, D, and G show coronal sections of mouse cortex at the indicated ages after in utero cortical electroporation of a GFP-coding plasmid at E15.5 in superficial neuron precursors in one brain hemisphere only (GFP signal in inverted color, dotted line indicates the limits of the brain). B, E, and H are a schematic representation of the main morphological changes observed in callosally projecting axons (red) at the corresponding ages. C shows the typical bipolar morphology of a migrating neuron emitting a trailing process (TP) and a leading process (LP) that will ultimately become the axon and dendrite, respectively. F and I show typical axon projections of layer 2/3 neurons located in the primary somatosensory area at P8 and P21, respectively. Neurons and axons in C, F, and I are visualized by GFP expression (inverted color). Image in C is modified from Barnes et al. (2007) with permission from Elsevier. Images in D, F, G, and I are reprinted from Courchet et al. (2013) with permission from Elsevier.

Neuronal polarization and axon specification

Neuronal polarization is the process of breaking symmetry in the newly born cell to create the asymmetry inherent to the formation of the axonal and somatodendritic compartments (Dotti and Banker, 1987). The mechanisms underlying this process have been studied extensively in vitro and more recently in vivo, but the exact sequence of events has remained elusive (Neukirchen and Bradke, 2011) partly because it is studied in various neuronal cell types that might not use the same extrinsic/intrinsic mechanisms to polarize. It is highly likely that at least three factors underlie neuronal polarization: extracellular cues, intracellular signaling cascades, and subcellular organelle localization. The partition-defective proteins (PARs) are a highly conserved family of proteins including two dyads (Par3/Par6 adaptor proteins and the Par4/Par1 serine/threonine kinases) that are required for polarization and axon formation (Shi et al., 2003, 2004; Barnes et al., 2007; Shelly et al., 2007; Chen et al., 2013), while many other intracellular signaling molecules also support axon formation (Oliva et al., 2006; Rašin et al., 2007; Barnes and Polleux, 2009; Shelly et al., 2010; Cheng et al., 2011; Hand and Polleux, 2011; Cheng and Poo, 2012; Gärtner et al., 2012). These intracellular signaling pathways are influenced by localized extracellular cues that instruct which neurite becomes the axon by either directly promoting axon extension or repressing axon growth in favor of dendritic growth (Adler et al., 2006; Yi et al., 2010; Randlett et al., 2011b; Shelly et al., 2011).The role of organelle subcellular localization during neuronal polarization is a more controversial issue. Initially, the orientation of organelles, including the Golgi complex, centrosomes, mitochondria, and endosomes, was shown to correlate with the neurite that becomes the axon in vitro (Bradke and Dotti, 1997; de Anda et al., 2005, 2010) and in vivo (de Anda et al., 2010). However, more recent studies suggest that the positioning of the centrosome is not necessary for neuronal polarization (Distel et al., 2010; Nguyen et al., 2011). Centrosome localization is likely constrained by microtubule organization within the cell, and therefore the centrosome position within the cell changes dynamically during different stages of polarization (Sakakibara et al., 2013). The question of how the interplay between extracellular cues, intracellular signaling, and organelle localization lead to polarization has pushed the field to perform more extensive in vivo imaging studies as in vitro systems/models have a difficult time recapitulating the complex environment and rely on neurons that were previously polarized in vivo.Like other epithelial cells, neural progenitors present a high degree of polarization along the apico-basal axis (Götz and Huttner, 2005). One of the major questions still needing to be addressed is how, or if, newly born mammalian neurons inherit some level of asymmetry from their parent progenitors (Barnes and Polleux, 2009). Recent studies have attempted to answer this question in vivo but have found that the answer might vary in each neuronal subtype. Retinal ganglion cells (RGCs), retinal bipolar neurons, and tegmental hindbrain nuclei neurons seem to inherit the apical/basolateral polarity from their progenitors (Morgan et al., 2006; Zolessi et al., 2006; Distel et al., 2010; Randlett et al., 2011a). In cortical neurons, hippocampal neurons, and cerebellar granule neurons, this relationship is unclear, in part because newly born cortical neurons first exhibit a multipolar morphology with dynamic neurites emerging from the cell body before adopting a bipolar morphology, suggesting they may not retain a predisposed parental polarity (Hand et al., 2005; Barnes et al., 2007). Other factors also suggest that different neuronal subtypes use different mechanisms during polarization. One such factor is the position where neurons specify their axon relative to the original apical/basolateral axis of their progenitors. As an example, cortical neurons in the mouse brain protrude an axon from the membrane facing the original apical surface toward the ventricular zone (Hand et al., 2005; Barnes et al., 2007; Shelly et al., 2007), whereas zebrafish RGCs form their axon from the membrane on the basolateral side (Zolessi et al., 2006; Randlett et al., 2011b). Another significant difference between cortical neurons and RGCs is related to the timing of axogenesis and dendrogenesis. RGCs tend to form their axons and dendrites at the same time during migration (Zolessi et al., 2006; Randlett et al., 2011b). However, cortical neurons form a long axon during migration before significant dendrite arborization takes place. These differences in the regulation of polarization and sequence of axon versus dendrite outgrowth may be linked to the localization of extracellular cues relative to the immature neuron during polarization (Yi et al., 2010).

Neuronal polarization, cytoskeletal dynamics, and polarized transport

What exactly makes the axonal compartment distinct from the somatodendritic domain? This can most easily be illustrated by focusing on the cytoskeleton that forms the framework of the developing axon. The cytoskeleton is composed of microtubules, actin filaments, and intermediate filaments (also called neurofilaments) along with their associated binding partners. Microtubules are composed of α- and β-tubulin subunits that polymerize to form a long filament intrinsically polarized by the addition of tubulin subunits to only one side of the growing filament called the plus end, while on the opposite side depolymerization occurs. It was discovered more than thirty years ago that the axon of a neuron contains a very uniform distribution of microtubules with the plus end facing away from the cell body (Heidemann et al., 1981). Through the years this observation was confirmed in many neuron cell types, and it was determined that dendrites do not have this uniform plus-end out network of microtubules (Fig. 2; Baas et al., 1988). Dendrites appear to have a complex array of microtubule orientations that may vary between species and/or neuronal subtypes. Current research shows that proximal dendrites are composed of mainly minus-end out microtubules, whereas more distal dendrites transition from an equal distribution of minus-end out and plus-end out microtubules to mainly plus-end out microtubules (Stone et al., 2008; Yin et al., 2011; Ori-McKenney et al., 2012). The orientation of microtubules matters greatly because it determines the relative contribution of microtubule-dependent motor proteins (kinesins and dyneins), which are the main motor proteins carrying various cargoes within cells and in particular are responsible for long-range transport in very large cells such as neurons. Dynein (a minus end–directed microtubule motor) is known to be responsible for both the transport of microtubules away from the cell body and for the uniform polarity of microtubules in the axon (Ahmad et al., 1998; Zheng et al., 2008). Recently, it was discovered that kinesin-1 (a plus end–directed microtubule motor) is required for the minus-end out orientation of microtubules in the dendrites of Caenorhabditis elegans DA9 neurons through selective transport of plus-end out microtubule fragments out of the dendrite (Yan et al., 2013). Another hallmark that differentiates the axonal and somatodendritic compartments is the microtubule-associated proteins (MAPs) that decorate microtubules to regulate their bundling and stability (Hirokawa et al., 2010). Microtubules in the axon are mainly decorated by Tau and MAP1B, whereas microtubules in the dendrites are labeled by proteins of the MAP2a-c family. The role of Tau in axon elongation remains controversial because early reports (Harada et al., 1994; Tint et al., 1998; Dawson et al., 2001) of Tau knockout alone suggested that axons were unaffected, but this apparent lack of phenotype might originate from the functional redundancy between MAPs as Tau/MAP1b double knockout mice show clear axon growth defects (Takei et al., 2000).Open in a separate windowFigure 2.Polarity maintenance and trafficking of somatodendritic and axonal proteins. Neurons are polarized into two main compartments: the somatodendritic domain and the axon. These domains are characterized by the underlying cytoskeleton and their unique protein signatures. The axonal cytoskeleton is defined by its uniform microtubule orientation where each microtubule is oriented with its plus end away from the cell body, while the dendrites contain a mixture of microtubules oriented in both directions. The proximal axon is characterized by a structure known as the axon initial segment (AIS, see inset). This highly ordered structure creates a diffusion barrier between the axonal compartment and the rest of the cell. F-actin is responsible for the cytoplasmic barrier, while sodium channels anchored by Ankyrin G form the basis of the plasma membrane barrier. Tau is retained in the axon by a microtubule-based filter at the AIS. Molecular motors (including kinesin, dynein, and myosin) then use the underlying cytoskeleton to restrict cargo transport to either the axon (such as Cntn1 and L1) or the dendrites (such as PSD95, AMPARs, and NMDARs).The dynamics of actin polymerization into actin filaments (F-actin) also play an important role in defining the axonal compartment, and contain an intrinsic polarity based on the polymerization of the free G-actin subunits (Hirokawa et al., 2010). Beyond the well-documented early role of F-actin dynamics in neurite outgrowth, multiple groups have shown that the disruption of actin polymerization allows dendritically localized proteins to incorrectly enter the axonal compartment (Winckler et al., 1999; Lewis et al., 2009; Song et al., 2009). The existence of a “diffusion barrier” in the proximal part of newly formed axons (Song et al., 2009) was long suspected. One of the current hypotheses is that a dense F-actin meshwork creates a cytoplasmic diffusion barrier shortly after polarization, which in part separates the axonal compartment from the neuronal cell body (Fig. 2, inset). Based on functional analysis and electron microscopy analysis, this “F-actin–based filter” is oriented so that the plus ends point toward the cell body while the minus ends point into the axon (Lewis et al., 2009, 2011; Watanabe et al., 2012). Two recent papers show via high resolution imaging techniques that indeed the axon has a unique F-actin network that is not found in dendrites (Watanabe et al., 2012; Xu et al., 2013). The development of this F-actin meshwork appears to directly precede the formation of the axon initial segment (AIS; Song et al., 2009; Galiano et al., 2012). An intra-axonal diffusion barrier, composed of Spectrins and Ankyrin B, defines the eventual position of the AIS. This boundary excludes Ankyrin G, which instead clusters in the most proximal part of the axon close to the cell body, where the AIS will form (Galiano et al., 2012). Ankyrin G is required for AIS formation and maintenance, and its loss causes the axon to start forming protrusions resembling dendritic spines (Hedstrom et al., 2008). Microtubules also play an important role at the AIS, as recent evidence suggests that Tau is retained in the axon through a microtubule-based diffusion barrier independently of the F-actin based filter (X. Li et al., 2011). The AIS is important in the formation of a plasma membrane barrier between the axonal and somatodendritic domains and its disruption affects both neuronal polarity and function because it is critical for clustering of voltage-dependent sodium channels and action potential initiation (Rasband, 2010).One of the critical cellular mechanisms underlying neuronal polarization is the polarized transport of various cargoes in axons and dendrites. Transport of proteins and various organelles is performed by the microtubule-dependent motor proteins kinesin and dynein (Hirokawa et al., 2010). Studies from many laboratories have demonstrated that kinesin motors can carry cargo to both the axonal and dendritic compartments (Burack et al., 2000; Nakata and Hirokawa, 2003). The mechanism for how selection occurs is not completely understood, but it probably incorporates both the affinity of the kinesin head for microtubules and the cargo bound to the motor protein (Nakata and Hirokawa, 2003; Song et al., 2009; Jenkins et al., 2012). In the axon, dynein works to bring cargo and retrograde signals back to the cell body, whereas in the dendrites it is responsible for much of the transport from the soma into the dendrites (Zheng et al., 2008; Kapitein et al., 2010; Harrington and Ginty, 2013). Additionally, the F-actin–dependent myosin motors can affect the polarized transport of cargos by using the F-actin–based cytoplasmic filter at the AIS to deny or facilitate entry of vesicles into the axon. Loss of the actin filter or myosin Va activity (a plus end–directed motor) allows dendritic cargos into the axon, whereas myosin VI (a minus end–directed motor) both removes axonal proteins from the dendritic surface and funnels vesicles containing axonal proteins through the actin filter at the AIS (Lewis et al., 2009, 2011; Al-Bassam et al., 2012). A current working hypothesis is that vesicles composed of multiple cargoes contain binding sites for each of these motors, and that through unknown mechanisms the activity of the motors can be differentially regulated to control the directionality of transport. An interesting example of how the interplay between different motors and cargo adaptors could lead to polarized transport was recently described for mitochondria (van Spronsen et al., 2013).

Axon growth

Microtubule dynamics regulate axon growth.

After axon specification, axon growth constitutes the second step of axonal development and is tightly linked to axon guidance toward the proper postsynaptic targets. Axon elongation by the growth cone is the product of two opposite forces (Fig. 3): slow axonal transport and the polymerization of microtubules providing a pushing force from the axon shaft, and the retrograde flow of actin providing a pulling force at the front of the growth cone (Letourneau et al., 1987; Suter and Miller, 2011). Although coordinated actin and microtubule dynamics are required for the proper function of the growth cone, it was reported that agents disrupting the actin cytoskeleton have limited consequences on axon elongation and are rather involved in axon guidance in vitro (Marsh and Letourneau, 1984; Ruthel and Hollenbeck, 2000) and in vivo (Bentley and Toroian-Raymond, 1986). Furthermore, local disruption of actin organization in the growth cone of minor neurites allows them to turn into axons (Bradke and Dotti, 1999; Kunda et al., 2001), indicating that the dense actin network present at the periphery of an immature neuronal cell body and in immature neurites may prevent microtubule protrusion and elongation necessary for axon specification.Open in a separate windowFigure 3.Cytoskeletal changes during axon elongation and branching. Representation of axon elongation and collateral branch formation in a cultured neuron. Axon growth is a discontinuous process, and collateral branches often originate from sites where the growth cone paused (gray dotted line), after it has resumed its progression. Other modalities of branch formation can occur through the formation of filopodia and lamellipodia. Red box shows a magnification of the main growth cone. Microtubules from the axon shaft spread into the central (C) zone. Some microtubules pass through the transition (T) zone, containing F-actin arcs, to explore filopodia from the peripheral (P) zone. Upon the proper stimulation by extracellular guidance cues or growth-promoting cues, microtubules are stabilized and invade the P-zone where they provide a pushing force, which, combined with the traction force from the actin treadmilling, provides the force required for growth cone extension. Green box shows the cytoskeletal changes occurring during collateral branch formation in the axon. Filopodia and lamellipodia are primarily F-actin–based protrusions that get invaded by microtubules, then elongate upon microtubule bundling. At later developmental stages, axon branches are stabilized or retracted (blue box) by mechanisms relying on the access to extracellular neurotrophins and/or neuronal activity and synapse formation.Contrary to actin, microtubule polymerization is required to sustain axon elongation and branching (Letourneau et al., 1987; Baas and Ahmad, 1993). Axonal proteins and cytoskeletal elements are transported along the axon through slow axonal transport by molecular motors (Yabe et al., 1999; Xia et al., 2003). It is still controversial whether tubulin and other cytoskeletal elements are transported in the axon as monomers and/or as polymers (Roy et al., 2000; Terada et al., 2000; Wang et al., 2000; Brown, 2003; Terada, 2003). Nonetheless, disruption of the slow transport of tubulin impairs the pushing force resulting from microtubule polymerization and impairs axon elongation (Suter and Miller, 2011). Therefore, it is not surprising that axon growth is affected in vitro and in vivo by disruption of plus-end microtubule-binding proteins such as APC (Shi et al., 2004; Zhou et al., 2004; Yokota et al., 2009; Chen et al., 2011) or EB1 and EB3 (Zhou et al., 2004; Jiménez-Mateos et al., 2005; Geraldo et al., 2008), microtubule-associated proteins such as MAP1B (Black et al., 1994; Takei et al., 2000; Dajas-Bailador et al., 2012; Tortosa et al., 2013), or proteins regulating microtubule severing and reorganization such as KIF2A (Homma et al., 2003), katanin, and spastin (Karabay et al., 2004; Yu et al., 2005; Wood et al., 2006; Butler et al., 2010).The contribution of microtubule dynamics to axon growth is not limited to growth cone dynamics but also involves axonal transport and polymerization along the axon shaft. Moreover, changing the balance between microtubule stabilization and depolymerization by local application of microtubule stabilizing agents is sufficient to instruct one neurite to grow and adopt an axon fate (Witte et al., 2008). Many kinase pathways converge on Tau and other axonal MAPs to regulate their function by phosphorylation (Morris et al., 2011). Among them, the MARK kinases regulate microtubule stability and axonal transport through phosphorylation of Tau (Drewes et al., 1997; Mandelkow et al., 2004). Interestingly, MARK-related kinases SAD-A/B control axon specification in part through phosphorylation of Tau (Barnes et al., 2007) and have very recently been linked to the growth and branching of the axons of sensory neurons (Lilley et al., 2013). Our work recently demonstrated that another family member related to MARKs and SAD kinases, called NUAK1, controls axon branching of mouse cortical neurons through the regulation of presynaptic mitochondria capture (Courchet et al., 2013). To what extent the regulation of Tau and other MAPs by the MARKs, SADs, and NUAK1 kinases contributes to axon elongation remains to be explored.

Where does axon elongation take place?

Growth cone progression and guidance constitute the main driver of axonal growth during development, but this process is unlikely to account for the totality of axon elongation. This is especially true after the axon has reached its final target but the axon shaft keeps growing in proportion to the rest of the body. One mechanism that may contribute to this “interstitial” form of axon elongation during brain/body size increase (see Fig. 1 for an example during postnatal cortex growth) is axon stretching, a process that can induce axon elongation in vitro (Smith et al., 2001; Pfister et al., 2004; Loverde et al., 2011) and in vivo (Abe et al., 2004). Aside from extreme stretching performed through the application of external forces, stretching could also contribute to the natural elongation of the axon in response to the tension resulting from growth cone progression (Suter and Miller, 2011).Axon elongation requires the addition of new lipids, proteins, cytoskeleton elements, and organelles along the axon. Where does the synthesis and incorporation of new components take place? Polysaccharides and cholesterol synthesis mostly occur in the cell body; however, lipid synthesis and/or incorporation can occur along the axon as well (Posse De Chaves et al., 2000; Hayashi et al., 2004). The growth cone is also a site of endocytosis, membrane recycling, and exocytosis (Kamiguchi and Yoshihara, 2001; Winckler and Yap, 2011; Nakazawa et al., 2012). One of the best studied examples of endocytosis and its role in axon growth and neuronal survival is the retrograde trafficking of TrkA receptor by target-derived nerve growth factor (NGF) in the peripheral nervous system (Harrington and Ginty, 2013).

Axon branching and presynaptic differentiation

Where do axon branches form?

The last step of axon development is terminal branching, which allows a single axon to connect to a broad set of postsynaptic targets. Collateral branches are formed along the axon through two distinct mechanisms: the first modality of branching is through splitting or bifurcation of the growth cone, which is linked to axon guidance and to the capacity of one single neuron to reach two targets that are far apart with one single axon. Growth cone splitting is observed in vivo in various neuron types including cortical neurons (Sato et al., 1994; Bastmeyer and O’Leary, 1996; Dent et al., 1999; Tang and Kalil, 2005), sympathetic neurons (Letourneau et al., 1986), motorneurons (Matheson and Levine, 1999), sensory neurons (Ma and Tessier-Lavigne, 2007), or mushroom body neurons in Drosophila (Wang et al., 2002). The second modality, known as interstitial branching, occurs through the formation of collateral branches directly along the axon shaft. Contrary to growth cone splitting, interstitial branching serves the purpose of raising axon coverage locally in order to define their “presynaptic territory”, and may contribute to increased network connectivity (Portera-Cailliau et al., 2005). Although both mechanisms can occur simultaneously in the same neuron, the relative importance of splitting versus interstitial branching is highly divergent from one neuron type to the other (Bastmeyer and O’Leary, 1996; Matheson and Levine, 1999; Portera-Cailliau et al., 2005).In culture, the axon grows in a non-continuous fashion with frequent growth cone pausing. Time-lapse imaging of sensorimotor neurons revealed that interstitial branching often occurs at the site where the growth cone paused, shortly after it has continued its course (Szebenyi et al., 1998). Accordingly, treatments with neurotrophins that slow the growth cone correlate with increased axon branching (Szebenyi et al., 1998). This suggests that growth cone pausing leaves a “mark” along the axon shaft that might predetermine future sites of branching (Kalil et al., 2000). However, local applications of neurotrophins shows that aside from growth cone pause sites the axon shaft remains competent to form collateral branches upon stimulation by extracellular factors (Gallo and Letourneau, 1998; Szebenyi et al., 2001), through the formation of filopodia or lamellipodia. Similar observations in vivo revealed that cortical axons are highly dynamic during development and form multiple filopodia that are the origin of collateral branches (Bastmeyer and O’Leary, 1996). Lamellipodia can be observed as motile, F-actin–dependent “waves” along the axon in vitro (Ruthel and Banker, 1998) and in vivo (Flynn et al., 2009). Moreover, disruption of microtubule organization impairs lamellipodia formation along the axon and is correlated with decreased axon branching (Dent and Kalil, 2001; Tint et al., 2009).

Cytoskeleton dynamics and axon branch formation.

Regardless of what type of protrusion gives rise to a branch, cytoskeletal reorganization in the nascent branch generally follows a similar sequence (Fig. 3): initially F-actin filament reorganization gives rise to a protrusion (filopodia, lamellipodia), followed by microtubule invasion of this otherwise transient structure to consolidate it, before the mature branch starts elongating through microtubule bundling (Gallo, 2011). Actin filaments accumulate along the axon and form “patches” that serve as nucleators for axon protrusions such as filopodia and lamellipodia (Korobova and Svitkina, 2008; Mingorance-Le Meur and O’Connor, 2009; Ketschek and Gallo, 2010). The mRNA for β-actin and regulators of actin polymerization such as Wave1 or Cortactin accumulate along the axons of sensory neurons and form hot-spots of local translation that are associated to NGF-dependent branching (Spillane et al., 2012; Donnelly et al., 2013). Subsequently, microtubules in the axon shaft undergo fragmentation at branch points as a prelude to branch invasion by microtubules (Yu et al., 1994, 2008; Gallo and Letourneau, 1998; Dent et al., 1999; Hu et al., 2012), a process that may disrupt transport locally to help trap molecules and organelles at branch points. Moreover, severed microtubules are transported into branches, a process required for branch stabilization (Gallo and Letourneau, 1999; Ahmad et al., 2006; Qiang et al., 2010; Hu et al., 2012). Interestingly, it is clear that, just like growth cone–mediated axon elongation, F-actin and microtubule reorganization events are interconnected to sustain axon branching (Dent and Kalil, 2001). As an example, microtubule-severing enzymes can also contribute to actin nucleation and filopodia formation (Hu et al., 2012).

Is axon branching linked to axon elongation?

Like in the growth cone, cytoskeleton reorganization constitutes the backbone of branch formation. It is therefore not surprising that many manipulations of the cytoskeleton affect both axon elongation and branch formation (Homma et al., 2003; Chen et al., 2011). Moreover, conditions that primarily disrupt axon elongation could secondarily disrupt branching by impairing the ability of the nascent branch to grow. However, axon elongation and axon branching can be considered as two separate phenomena and can be operationally separated because conditions disrupting one do not systematically affect the other. As an example, the microtubule-severing proteins katanin and spastin have differential consequences on axon elongation (primarily dependent upon katanin function) and branching (mostly spastin mediated; Qiang et al., 2010), taxol treatment (which stabilizes microtubules) affects axon elongation but not branching (Gallo and Letourneau, 1999), and disruption of TrkA endocytosis by knock-down of Unc51-like kinase (ULK1/2) proteins has opposite effects on axon elongation and branching (Zhou et al., 2007). In vivo, superficial layer cortical neurons initially go through a phase of elongation through the corpus callosum without branching (see Fig. 1), then stop elongating and form collateral branches in the contralateral cortex (Mizuno et al., 2007; Wang et al., 2007). It is conceivable that even before myelination, axons are actively prevented from branching at places and stages when they elongate (for example in the white matter of the neocortex) where they tend to be highly fasciculated. The identities of the molecules that inhibit interstitial branching along the axon shaft are currently unknown.

Regulation of axon branching by activity.

Immature neurons display spontaneous activity in the form of calcium waves (Gu et al., 1994; Gomez and Spitzer, 1999; Gomez et al., 2001) and spontaneous vesicular release long before they have completed axon development, which suggested a role for early neuronal activity in axon development and guidance (Catalano and Shatz, 1998). Cell-autonomous silencing of neurons in vivo by transfection of the hyperpolarizing inward-rectifying potassium channel Kir2.1 in olfactory neurons (Yu et al., 2004), in RGCs (Hua et al., 2005) or in cortical pyramidal neurons (Mizuno et al., 2007; Wang et al., 2007), or in vitro through infusion of tetrodotoxin (which blocks action potentials generation) in co-cultures of thalamo-cortical projecting neurons (Uesaka et al., 2007) results in a decrease in terminal axon branching, indicating that synaptic activity is required for axons to fully develop their branching pattern. Moreover, inhibition of synaptic release by expression of tetanus toxin light chain (TeTN-LC; Wang et al., 2007) also abolished terminal axon branching, suggesting that the formation of functional presynaptic release sites is required cell autonomously to control terminal axon branching. However, one potential limitation of the experiments involving TeTN-LC is that it blocks most VAMP-mediated vesicular release (with the exception of VAMP7, also called tetanus toxin–independent VAMP, or TI-VAMP). Therefore TeTN-LC action may not be limited to blocking synaptic vesicle release, but could also inhibit peptide release through vesicles containing neurotrophins for example, or other important trophic factors required for axon branching. More recent experiments through silencing of postsynaptic targets revealed that branching of callosal or thalamocortical axons is also dependent upon the activity of the postsynaptic targets (Mizuno et al., 2010; Yamada et al., 2010), albeit activity of the presynaptic neuron is required earlier during the branching process than activity of the postsynaptic targets (Mizuno et al., 2010). Activity is also required in some neurons at the phase of axon elongation through a feedback loop involving the activity-dependent up-regulation of guidance molecules (Mire et al., 2012).How much does spontaneous or evoked neuronal activity contribute to branching? Reduction of neuronal activity through hyperpolarization induced by overexpression of Kir2.1 significantly reduces axon branching without completely eliminating it (Hua et al., 2005; Mizuno et al., 2007; Wang et al., 2007). Activity seems to serve as a competitive regulator of axon branching with regard to its neighbors because silencing of neighboring axons restores normal branching (Hua et al., 2005). Interestingly, neuronal activity induces neurotrophin expression locally, suggesting that activity can contribute to branching partly through activation of activity-independent branching mechanisms (Calinescu et al., 2011).Neuronal activity can regulate branching through modification of the actin cytoskeleton via RhoA activation (Ohnami et al., 2008), and mRNA accumulates at presynaptic sites, indicating a correlation between local translation and synaptic activity (Lyles et al., 2006; Taylor et al., 2013). Neuronal activity is associated with changes in intracellular Ca²⁺ signaling, which has been shown to play a deterministic function in axon growth (Gomez and Spitzer, 1999). Calcium signaling activates the Ca²⁺/calmodulin-dependent kinases (CAMKs) that are known to regulate axon branching in vitro (Wayman et al., 2004; Ageta-Ishihara et al., 2009) and in vivo (Ageta-Ishihara et al., 2009).

Stabilization and refinement of the axonal arborization.

Axon branches are often formed in excess during development, then later refined to select for specific neural circuits (Luo and O’Leary, 2005). Long-range axon branch retraction has long been observed in layer V cortical neurons that initially project to the midbrain, hindbrain, and spinal cord (O’Leary and Terashima, 1988; Bastmeyer and O’Leary, 1996). At later stages, pyramidal neurons from the primary visual cortex will retract their spinal projection through axon pruning, whereas pyramidal neurons from the primary motor cortex will stabilize this projection but retract their axonal branches growing toward visual targets such as the superior colliculus. The molecular mechanisms controlling this area-specific pattern of axon branch pruning are still poorly understood, but seem to involve extracellular cues such as semaphorins and Rac1-dependent signaling (Bagri et al., 2003; Low et al., 2008; Riccomagno et al., 2012). Another example is the well-characterized refinement of retino-geniculate axons during the selective elimination of binocular input of RGC axon synapses onto relay neurons in the dorsal lateral geniculate nucleus (Muir-Robinson et al., 2002). Interestingly, some axons use caspase-dependent pathways locally to induce the selective retraction of axon branches during the process of pruning (Nikolaev et al., 2009; Simon et al., 2012).Circuit refinement and selective branch retraction can be observed in vivo at the level of the neuromuscular junction where individual branches of motor axons are eliminated asynchronously (Keller-Peck et al., 2001). In the developing CNS, neurotrophin-induced branch retraction can also be observed in a context of competition between neighboring axons (Singh et al., 2008). One other way of stabilizing axon branches is through the formation of synapses with postsynaptic targets. In the visual system, the initial axon arbor is refined to establish ocular dominance through activity-dependent retraction of less active branches (Ruthazer et al., 2003). Time-lapse imaging of RGC axons in zebrafish or in Xenopus tadpole revealed that the formation of presynaptic sites occurs concomitantly to axon branching, and branches that form presynaptic structures are less likely to retract (Meyer and Smith, 2006; Ruthazer et al., 2006). The stabilization of axon branches through formation of synaptic contacts parallels with the stabilization of dendritic branches through synapse formation and stabilization (Niell et al., 2004; J. Li et al., 2011). The role of presynaptic bouton formation goes beyond the stabilization of axonal branches because in vivo, new axon branches can emerge from existing presynaptic terminals (Alsina et al., 2001; Javaherian and Cline, 2005; Panzer et al., 2006).In conclusion, axon growth and branching can be regulated by both activity-dependent and activity-independent mechanisms during development. However, for mammalian CNS axons, much more work is needed to define (1) the precise molecular mechanisms underlying axon branching; (2) the cellular and molecular mechanisms regulating the key transition between axon growth and branching when axons start forming presynaptic contacts with their postsynaptic partners; and (3) the mechanisms regulating axon pruning during synapse elimination.     

New insights into the biology of preeclampsia

Despite recent research progress, the biology of preeclampsia is still poorly understood and neither effective prediction nor causal therapy have yet emerged. Nevertheless, recent studies have documented new and exciting pathophysiological mechanisms for the origin and development of preeclampsia. These studies provide a more differentiated view on alterations of particular peptide systems with strong impact on angiogenesis and cardiovascular regulation in this pregnancy disorder. With the identification of the antiangiogenic factor soluble fms-like tyrosine kinase 1 and the agonistic autoantibody to the angiotensin II type 1 receptor, two factors have been described with a clear linkage to the development of the disease. This review focuses on the most recent and relevant insights into the biology of preeclampsia and develops hypotheses regarding possible links between the reported aspects of preeclampsia.     

铁死亡：一种新的细胞死亡方式

死亡不仅是所有细胞的最终命运,而且它与细胞分裂、增殖一样,在整个机体的生长、发育中具有不可替代的作用.目前认为,除了坏死外,细胞死亡形式分为程序性细胞死亡(programmedcell death,PCD),包括凋亡(apoptosis)和自噬(autophagy),及非程序性细胞死亡(non-programmedcell death,NPCD),包括副凋亡(paraptosis)、细胞有丝分裂灾难(mitotic catastrophe)和胀亡(oncosis)     

Autophagy and Caspases: A New Cell Death Program

Autophagy is used to degrade components of the cytoplasm and functions as a cell survival mechanism during nutrient deprivation. Autophagic structures have also been observed in many types of dying cells, but experimental evidence for autophagy playing a role in the regulation of programmed cell death is limited. We have recently shown that the autophagy genes Atg7 and Beclin1 are required for the death of certain cells, thus demonstrating that this mechanism of proteolysis is involved in both survival and death. The factors that enable autophagy to regulate distinct cell survival and death responses are not clear, and future work is needed to determine the mechanism(s) that regulate autophagic cell death.     

New insights into developing antibiofouling surfaces for industrial photobioreactors

The biofouling formation of the marine microalga Nannochloropsis gaditana on nontoxic surfaces was quantified on rigid materials, both coated (with fouling release coatings and nanoparticle coatings) and noncoated, to cover a wide range of surface properties from strongly hydrophobic to markedly hydrophilic under conditions similar to those prevailing in outdoor massive cultures of marine microalgae. The effect of seawater on surfaces that presented the best antibiofouling properties was also evaluated. The adhesion intensity on the different surfaces was compared with the predictions of the biocompatibility theories developed by Baier and Vogler using water adhesion tension (τ⁰) as the quantitative parameter of surface wettability. For the most hydrophobic surfaces, τ⁰ ≤ 0, the microalgae adhesion density increased linearly with τ⁰, following the Baier's theory trend. However, for the rest of the surfaces, τ⁰ ≥ 0, a tendency toward minimum adhesion was observed for amphiphilic surfaces with a τ⁰ = 36 mJ/m², a value close to that which minimizes cell adhesion according to Vogler's theory. The understanding and combination of the two biocompatibility theories could help to design universal antibiofouling surfaces that minimize the van der Waals forces and prevent foulant adsorption by using a thin layer of hydration.     

Food behaviour and obesity: insights from decision neuroscience

Neuroimaging allows to estimate brain activity when individuals are doing something. The location and intensity of this estimated activity provides information on the dynamics and processes that guide choice behaviour and associated actions that should be considered a complement to behavioural studies. Decision neuroscience therefore sheds new light on whether the brain evaluates and compares alternatives when decisions are made, or if other processes are at stake. This work helped to demonstrate that the situations faced by individuals (risky, uncertain, delayed in time) do not all have the same (behavioural) complexity, and are not underlined by activity in the cerebral networks. Taking into account brain dynamics of people (suffering from obesity or not) when making food consumption decisions might allow for improved strategies in public health prevention, far from the rational choice theory promoted by neoclassical economics.     

New insights regarding the biology of Giardia lamblia

Giardia lamblia is one of the most common causes of intestinal disease in humans. To adapt to environments both inside and outside of the host's small intestine, this protozoan parasite undergoes significant developmental changes during its life cycle. In this review, we analyze and discuss the most recent findings regarding the process of Giardia trophozoites differentiation into infective cysts as well as the mechanism of antigenic variation, which allows the parasite to cause chronic and recurrent infections in susceptible individuals.     

New insights into the biology of cytokinin degradation

A survey of recent results is presented concerning the role of cytokinin degradation in plants, which is catalyzed by cytokinin oxidase/dehydrogenase (CKX) enzymes. An overview of Arabidopsis CKX gene expression suggests that their differential regulation by biotic and abiotic factors contributes significantly to functional specification. Here, we show using reporter gene and semiquantitative RT-PCR analyses regulation of individual CKX genes by cytokinin, auxin, ABA, and phosphate starvation. Partially overlapping expression domains of CKX genes and cytokinin-synthesizing IPT genes in meristematic tissues and endo-reduplicating cells lend support for a locally restricted function of cytokinin. On the other hand, their expression in vascular tissue suggests a function in controlling transported cytokinin. Recent studies led to a model for the biochemical reaction mechanism of CKX-mediated catalysis, which was refined on the basis of the three-dimensional enzyme structure. Last but not least, the developmental functions of CKX enzymes are addressed. The recent identification of the rice OSCKX2 gene as an important novel breeding tool is highlighted. Together the results corroborate the relevance of metabolic control in determining cytokinin activity.     

Systems biology in neuroscience: bridging genes to cognition

Systems biology is a new branch of biology aimed at understanding biological complexity. Genomic and proteomic methods integrated with cellular and organismal analyses allow modelling of physiological processes. Progress in understanding synapse composition and new experimental and bioinformatics methods indicate the synapse is an excellent starting point for global systems biology of the brain. A neuroscience systems biology programme, organized as a consortium, is proposed.     

Social neuroscience: mirror neurons recorded in humans

New single-cell recordings show that humans do have mirror neurons, and in more brain regions than previously suspected. Some action-execution neurons were seen to be inhibited during observation, possibly preventing imitation and helping self/other discrimination.     

肿瘤细胞死亡的一种新形式——铁死亡

铁死亡是近年来发现的一种程序性细胞死亡新形式,其主要特征是在发生于线粒体内的铁依赖性脂质过氧化物损伤诱导的细胞死亡。铁死亡细胞在形态、蛋白质及基因水平的变化均不同于细胞凋亡、坏死和自噬。2012年,铁死亡概念首次被提出后,铁死亡逐渐成为科学研究的热点。Erastin以及RSL3是铁死亡的诱导剂,谷胱甘肽过氧化物酶4（glutathione peroxidase 4,GPX4）是铁死亡的关键调节点,GPX4的表达量减少或活性降低均可诱导铁死亡的发生。胱氨酸-谷氨酸逆向转运蛋白（system Xc^-）可将细胞内的谷氨酸排出,同时将细胞外胱氨酸转运入细胞内,促进细胞内谷胱甘肽的合成,维持GPX4酶的活性。新近的研究表明,p62-keap1-Nrf2、P53-SAT1-ALOX15是铁死亡的关键调控通路,p53、BECN1以及BAP1是铁死亡的关键调节因子。Erastin以及RSL3可以选择性杀死RAS突变的肿瘤细胞,且越来越多的研究也证明,诱导肿瘤细胞铁死亡在免疫治疗以及逆转耐药方面均有着重要作用。因此,调控肿瘤细胞铁死亡很可能成为治疗肿瘤的新手段。本文就诱导肿瘤细胞铁死亡的机制及其进展作一综述。     

肿瘤细胞死亡的一种新形式——铁死亡

铁死亡是近年来发现的一种程序性细胞死亡新形式,其主要特征是在发生于线粒体内的铁依赖性脂质过氧化物损伤诱导的细胞死亡。铁死亡细胞在形态、蛋白质及基因水平的变化均不同于细胞凋亡、坏死和自噬。2012年,铁死亡概念首次被提出后,铁死亡逐渐成为科学研究的热点。Erastin以及RSL3是铁死亡的诱导剂,谷胱甘肽过氧化物酶4（glutathione peroxidase 4,GPX4）是铁死亡的关键调节点,GPX4的表达量减少或活性降低均可诱导铁死亡的发生。胱氨酸-谷氨酸逆向转运蛋白（system Xc^-）可将细胞内的谷氨酸排出,同时将细胞外胱氨酸转运入细胞内,促进细胞内谷胱甘肽的合成,维持GPX4酶的活性。新近的研究表明,p62-keap1-Nrf2、P53-SAT1-ALOX15是铁死亡的关键调控通路,p53、BECN1以及BAP1是铁死亡的关键调节因子。Erastin以及RSL3可以选择性杀死RAS突变的肿瘤细胞,且越来越多的研究也证明,诱导肿瘤细胞铁死亡在免疫治疗以及逆转耐药方面均有着重要作用。因此,调控肿瘤细胞铁死亡很可能成为治疗肿瘤的新手段。本文就诱导肿瘤细胞铁死亡的机制及其进展作一综述。     

Epigenetics and Cell Death: DNA Hypermethylation in Programmed Retinal Cell Death

Background

Vertebrate genomes undergo epigenetic reprogramming during development and disease. Emerging evidence suggests that DNA methylation plays a key role in cell fate determination in the retina. Despite extensive studies of the programmed cell death that occurs during retinal development and degeneration, little is known about how DNA methylation might regulate neuronal cell death in the retina.

Methods

The developing chicken retina and the rd1 and rhodopsin-GFP mouse models of retinal degeneration were used to investigate programmed cell death during retinal development and degeneration. Changes in DNA methylation were determined by immunohistochemistry using antibodies against 5-methylcytosine (5mC) and 5-hydroxymethylcytosine (5hmC).

Results

Punctate patterns of hypermethylation paralleled patterns of caspase3-dependent apoptotic cell death previously reported to occur during development in the chicken retina. Degenerating rd1 mouse retinas, at time points corresponding to the peak of rod cell death, showed elevated signals for 5mC and 5hmC in photoreceptors throughout the retina, with the most intense staining observed in the peripheral retina. Hypermethylation of photoreceptors in rd1 mice was associated with TUNEL and PAR staining and appeared to be cCaspase3-independent. After peak rod degeneration, during the period of cone death, occasional hypermethylation was observed in the outer nuclear layer.

Conclusion

The finding that cell-specific increases of 5mC and 5hmC immunostaining are associated with the death of retinal neurons during both development and degeneration suggests that changes in DNA methylation may play a role in modulating gene expression during the process of retinal degeneration. During retinal development, hypermethylation of retinal neurons associates with classical caspase-dependent apoptosis as well as caspase-3 independent cell death, while hypermethylation in the rd1 mouse photoreceptors is primarily associated with caspase-3 independent programmed cell death. These findings suggest a previously unrecognized role for epigenetic mechanisms in the onset and/or progression of programed cell death in the retina.     

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.