Cell biology in neuroscience: Death of developing neurons: New insights and implications for connectivity

The concept that target tissues determine the survival of neurons has inspired much of the thinking on neuronal development in vertebrates, not least because it is supported by decades of research on nerve growth factor (NGF) in the peripheral nervous system (PNS). Recent discoveries now help to understand why only some developing neurons selectively depend on NGF. They also indicate that the survival of most neurons in the central nervous system (CNS) is not simply regulated by single growth factors like in the PNS. Additionally, components of the cell death machinery have begun to be recognized as regulators of selective axonal degeneration and synaptic function, thus playing a critical role in wiring up the nervous system.

Why do so many neurons die during development?

Programmed cell death occurs throughout life, as cell turnover is part of homeostasis and maintenance in most organs and tissues. The situation in the nervous system is principally different, as the vast majority of neurons undergo their last round of cell division early in development. Soon after exiting the cell cycle, neurons start elongating axons to innervate their targets. It is during this period that they are highly susceptible to undergo programmed cell death: a large percentage, as much as 50% in several ganglia in the peripheral nervous system (PNS) as well as in various central nervous system (CNS) areas, is eliminated around the time that connections are being made with other cells. Later in development, the propensity of neurons to initiate apoptosis progressively decreases. The likelihood for a neuron to undergo apoptosis seems to be determined by a tightly regulated apoptotic machinery (summarized in Fig. 1). Therefore, modulation of the expression levels or the activity of components of this apoptotic balance changes the sensitivity to death-promoting cues, allowing temporal restriction of cell death.Open in a separate windowFigure 1.Core components of the apoptotic machinery. The likelihood that a neuron undergoes apoptosis is determined by the interplay of the tightly interlinked apoptotic machinery, many components of which are highly conserved between species. The critical, and often terminal, step in programmed cell death is the proteolytic activation of the executor caspases (such as caspase 3, 6, 7) by the initiator caspases (i.e., caspase 8, 9, and 10; Riedl and Salvesen, 2007). In mammalian cells, initiation of the executor caspases is regulated by two distinct protein cascades: the intrinsic pathway, also known as the mitochondrial pathway, and the extrinsic pathway. The intrinsic pathway integrates a number of intra- and extracellular signal modalities, such as redox state (for example, the reactive oxygen species; Franklin, 2011), DNA damage (Sperka et al., 2012), ER stress (Puthalakath et al., 2007) and growth factor deprivation (Deckwerth et al., 1998; Putcha et al., 2003; Bredesen et al., 2005), or activation of the p75NTR neurotrophin receptor by pro-neurotrophins (Nykjaer et al., 2005). The stressors converge onto pro- and anti-apoptotic members of the Bcl-2 protein family (for example: BCL-2, BCL-X_l, BAX, and tBID; Youle and Strasser, 2008). These proteins regulate the release of cytochrome c from mitochondria, which activates the initiator caspase 9 through Apaf1 (Riedl and Salvesen, 2007). The extrinsic pathway links activation of ligand-bound death receptors (such as Fas/CD95 and TNFR) to the initiator caspase 8 and 10, through formation of the death-inducing signaling complex (DISC; LeBlanc and Ashkenazi, 2003; Peter and Krammer, 2003). Together with additional regulatory elements (including the Inhibitors of apoptosis proteins [IAP]; Vaux and Silke, 2005) and cFLIP (Scaffidi et al., 1999; Wang et al., 2005), the apoptotic machinery forms a balance that determines the propensity of the neuron to undergo apoptosis.Programmed cell death eliminates many neurons during development, even in organisms comprised of only few cells, such as Caenorhabditis elegans. As neurons and their targets are initially separated, it is possible that the initial generation of an overabundance of neurons is simply part of a mechanism to ensure that distal targets are adequately innervated (Oppenheim, 1991; Conradt, 2009; Chen et al., 2013). In various tissues other than the nervous system, programmed cell death is used to eliminate cells that are no longer needed, defective, or harmful to the function of the organism. However, there is strong evidence that the elimination of superfluous neurons in the developing nervous system is not essential. For example, early work in C. elegans revealed that preventing programmed cell death does not result in significant behavioral alterations (Ellis and Horvitz, 1986). In the C57BL/6 mouse strain, deletion of the executor caspases 3 and 7 (Fig. 1) has a remarkably limited neuropathological and morphological impact in the CNS (Leonard et al., 2002; Lakhani et al., 2006) compared with the 129X1/SvJ strain, in which deletion of these caspases causes neurodevelopmental defects (Leonard et al., 2002). Similar conclusions were reached by blocking the Bcl-2–associated X protein (BAX)–dependent pathway in many neuronal populations, including motoneurons (Buss et al., 2006a). A recent study in the developing retina showed that in mice lacking the central apoptotic regulator BAX, the normal mosaic distribution of intrinsically photosensitive retinal ganglion cells (ipRGCs) was perturbed (Chen et al., 2013). Although this abnormal distribution is dispensable for the intrinsic photosensitivity of the ipRGCs, it is required for establishing proper connections to other neurons in the retina, which is necessary for rod/cone photo-entrainment (Chen et al., 2013). Even though this finding highlights a physiological role for programmed cell death in the CNS, the functional consequences remain rather underwhelming in the face of a process that eliminates such large numbers of neurons (Purves and Lichtman, 1984; Oppenheim, 1991; Miller, 1995; Gohlke et al., 2004). It thus appears that apoptotic removal of the surplus neurons generated during development mainly serves the purpose to optimize the size of the nervous system to be minimal, but sufficient.

A molecular substrate for the neurotrophic theory

Quantitatively, programmed cell death of neurons in the PNS and CNS is most dramatic when neurons start contacting the cells they innervate. Because experimental manipulations such as target excision typically lead to the death of essentially all innervating neurons (Oppenheim, 1991), the concept emerged that the fate of developing neurons is regulated by their targets. This notion is often referred to as the “neurotrophic theory” (Hamburger et al., 1981; Purves and Lichtman, 1984; Oppenheim, 1991), but it is important to realize that it evolved in the absence of direct mechanistic or molecular support (Purves, 1988). Originally described as a diffusible agent promoting nerve growth, the eponymous NGF later provided a strong and very appealing molecular foundation for this theory (Korsching and Thoenen, 1983; Edwards et al., 1989; Hamburger, 1992). The tyrosine kinase receptor tropomyosin receptor kinase A (TrkA), which was initially identified as an oncogene (Martin-Zanca et al., 1986), was fortuitously discovered to be the critical receptor necessary for the prevention of neuronal cell death by NGF (Klein et al., 1991). Both the remarkable expression pattern of TrkA in NGF-dependent neurons and the onset of its expression during development (Martin-Zanca et al., 1990) provided further additional support for the neurotrophic theory. However, for a surprisingly long time, the question was not asked as to why only specific populations of neurons strictly depend on NGF for survival, while others do not. Indeed, it was only recently shown that TrkA causes cell death of neurons by virtue of its mere expression, and that this death-inducing activity is prevented by addition of NGF (Nikoletopoulou et al., 2010). These findings thus indicate that TrkA acts as a “dependence receptor,” a concept introduced after observations that various cell types die when receptors are expressed in the absence of their cognate ligands (Bredesen et al., 2005; Tauszig-Delamasure et al., 2007). Accordingly, embryonic mouse sympathetic or sensory neurons survive in the absence of NGF when TrkA is deleted (Nikoletopoulou et al., 2010). The closely related neurotrophin receptor TrkC also acts as a dependence receptor (Tauszig-Delamasure et al., 2007; Nikoletopoulou et al., 2010). Here, it is interesting to note a series of older, convergent results indicating that deletion of neurotrophin-3 (NT3), the TrkC ligand, leads to a significantly larger loss of sensory and sympathetic neurons in the PNS than the deletion of TrkC (Tessarollo et al., 1997). This phenotypic discrepancy fits well with the idea that inactivation of the ligand of a dependence receptor is expected to yield a more profound phenotype than inactivation of the receptor itself (Tauszig-Delamasure et al., 2007). How TrkA and TrkC induce apoptosis remains to be fully elucidated. It seems that proteolysis is involved, either of TrkC itself (Tauszig-Delamasure et al., 2007), as was suggested for other dependence receptors (Bredesen et al., 2005), or by the proteolysis of the neurotrophin receptor p75NTR, which associates with both TrkA and TrkC (Fig. 2; Nikoletopoulou et al., 2010). Surprisingly, although TrkA and TrkC cause cell death, the structurally related TrkB receptor does not (Nikoletopoulou et al., 2010), a difference that appears to be accounted for by their differential localization in the cell membrane. TrkA and TrkC colocalize with p75NTR in lipid rafts, whereas TrkB, which also associates with p75NTR (Bibel et al., 1999), is excluded from lipid rafts (Fig. 2; unpublished data). Interestingly, the transmembrane domains of TrkA and TrkC are closely related, and differ clearly from that of TrkB. It turns out that a chimeric protein of TrkB with the transmembrane domain of TrkA causes cell death, which can be prevented by the addition of the TrkB ligand brain-derived neurotrophic factor (BDNF; unpublished data). The suggestion that the lipid raft localization of TrkA and TrkC is important for their death-inducing function is in line with a number of reports indicating that certain apoptotic proteins preferentially localize in lipid rafts in the plasma membrane. After activation of the extrinsic apoptosis pathway, translocation of the activated receptors to lipid rafts in the membrane is required for assembling the death-inducing signaling complex (DISC; Davis et al., 2007; Song et al., 2007). Indeed, regulators of the extrinsic pathway (e.g., cFLIP; Fig. 1) prevent this translocation, explaining how they attenuate cell death induction (Song et al., 2007). Similarly, the localization of the dependence receptor DCC (deleted in colorectal cancer) in lipid rafts is a prerequisite for its pro-apoptotic activity in absence of its ligand, Netrin-1 (Furne et al., 2006).Open in a separate windowFigure 2.TrkA and TrkC as dependence receptors: mode of action and contrast with TrkB. All Trk receptors associate with the pan-neurotrophin receptor p75NTR (Bibel et al., 1999). A critical step in the induction of apoptosis by TrkA is the release of the intracellular death domain of p75NTR by the protease γ-secretase (Nikoletopoulou et al., 2010), which is localized in lipid rafts (Urano et al., 2005). Our membrane fractionation studies indicate that while TrkA and TrkC associate with p75NTR in lipid rafts, TrkB associated with p75NTR is excluded from this membrane domain (unpublished data). The 24–amino acid transmembrane domain of the Trk receptors may be responsible for this differential localization (see text).Despite the fact that TrkB does not act as a dependence receptor, its activation by BDNF is required for the survival of several populations of cranial sensory neurons (Ernfors et al., 1995; Liu et al., 1995). It appears that other death-inducing receptors predispose these neurons to be eliminated, such as p75NTR, which is expressed at high levels in some of these ganglia, or TrkC in vestibular neurons (Stenqvist et al., 2005). This latter case is of special interest, as NT3 is known not to be required for the survival of these neurons (Stenqvist et al., 2005). In addition to inducing apoptosis in the absence of their ligand, TrkA and TrkC have long been recognized to have a pro-survival function similar to TrkB, as can be inferred from the loss of specific populations of peripheral sensory neurons in mutants lacking these receptors (Klein et al., 1994; Smeyne et al., 1994).

Cell death in the CNS

Although TrkA is primarily expressed in peripheral sympathetic and sensory neurons, it is also found in a small population of cholinergic neurons in the basal forebrain (Sobreviela et al., 1994), a proportion of which requires NGF for survival (Hartikka and Hefti, 1988; Crowley et al., 1994; Müller et al., 2012). Selective deletion of TrkA was recently shown not to cause the death of these neurons (Sanchez-Ortiz et al., 2012). This supports the notion that TrkA acts as a dependence receptor for this small population of CNS neurons, like for peripheral sensory and sympathetic neurons. TrkA activation by NGF is essential for the maturation, projections, and function of these neurons (Sanchez-Ortiz et al., 2012), as was previously described for sensory neurons in the PNS as well (Patel et al., 2000).Whether or not receptors other than TrkA act as dependence receptors in the CNS is an important open question, particularly because TrkB, which is expressed highly by most CNS neurons, does not act as a dependence receptor (Nikoletopoulou et al., 2010). In retrospect, the structural similarities between TrkA and TrkB, just like those between NGF and BDNF (Barde, 1989), have substantially misled the field by suggesting that BDNF would act in the CNS like NGF in the PNS. Adding to the confusion were early findings showing that BDNF supports the growth of spinal cord motoneurons in vitro or in vivo after axotomy (Oppenheim et al., 1992; Sendtner et al., 1992; Yan et al., 1992). However, in the absence of lesion, deletion of BDNF does not lead to significant losses of neurons in the developing or adult CNS (Ernfors et al., 1994a; Jones et al., 1994; Rauskolb et al., 2010), unlike the case in some populations of PNS neurons. The poor correlation of the role of BDNF in CNS development and in axotomy and in vitro experiments is surprising, especially because the role of NGF in vivo could in essence be recapitulated by in vitro experiments. Although the reasons for this discrepancy are not fully understood, the strong up-regulation of death-inducing molecules such as p75NTR after axotomy (Ernfors et al., 1989) may be a part of the explanation. At present, most of the growth factors promoting the survival of PNS neurons fail to show significant survival properties for developing neurons in the CNS, as for example was shown for NT3 (Ernfors et al., 1994b; Fariñas et al., 1994), glial cell line–derived neurotrophic factor (GDNF; Henderson et al., 1994), ciliary neurotrophic factor (CNTF; DeChiara et al., 1995), and several others.In the developing CNS, neuronal activity and neurotransmitter input seem to play a more significant role than single growth factors in regulating neuronal survival. In particular, it has been known for a long time that blocking synaptic transmission at the neuromuscular junction has a pro-survival effect on spinal cord motoneurons (Pittman and Oppenheim, 1978; Oppenheim et al., 2008). By contrast, surgical denervation of afferent connections leads to increased apoptosis of postsynaptic neurons (Okado and Oppenheim, 1984), whereas inhibiting glycinergic and GABAergic synaptic transmission has both pro- and anti-apoptotic effects on motoneurons (Banks et al., 2005). Throughout the developing brain, blocking glutamate-mediated synaptic transmission involving NMDA receptors markedly increases normally occurring neuronal death (Ikonomidou et al., 1999; Heck et al., 2008). The mechanism involves a reduction of neuronal expression of anti-apoptotic proteins, such as B-cell lymphoma 2 (BCL-2; Hansen et al., 2004). Conversely, a limited increase in neuronal activity leads to down-regulation of the pro-apoptotic genes BAX and caspase 9 (Léveillé et al., 2010), thereby reducing the propensity of these cells to initiate programmed cell death (Hardingham et al., 2002). In addition to directly modulating the expression of apoptotic proteins, neuronal activity affects the expression of several secreted growth factors, such as BDNF (Hardingham et al., 2002; Hansen et al., 2004) and GDNF (Léveillé et al., 2010). So, even though BDNF is not a major survival factor in the developing CNS, it appears to be critical for activity-dependent neuroprotection (Tremblay et al., 1999). A recent publication revealed that certain populations of neurons in the CNS do not follow the predictions of the neurotrophic theory and showed that apoptosis of cortical inhibitory neurons is independent of cues present in the developing cerebral cortex (Southwell et al., 2012). This study indicates that programmed cell death of a large proportion of interneurons in the CNS is regulated by intrinsic mechanisms that are largely resistant to the presence or absence of extrinsic cues (Dekkers and Barde, 2013).Taken together, even though the extent of naturally occurring cell death in the different regions of the CNS is not nearly as well characterized as in the PNS, let alone quantified, it appears that its regulation may significantly differ. Although single secreted neurotrophic factors seem to be largely dispensable for survival, neuronal activity and other intrinsic mechanisms drive the propensity of the neurons in the CNS to undergo apoptosis. An important open question in this context is a possible involvement of non-neuronal cells, such as glial cells (see Corty and Freeman, in this issue).

The apoptotic machinery as a regulator of connectivity

Activation of the executor caspases has been most studied in cell bodies and typically results in the demise of the entire cell (Williams et al., 2006). However, recent evidence shows that caspases are also activated locally in neuronal processes and branches destined to be eliminated, for example in axons overshooting their targets that are subsequently pruned back to establish the precise adult connectivity (Finn et al., 2000; Raff et al., 2002; Luo and O’Leary, 2005; Buss et al., 2006b). Initially, axonal degeneration and axon pruning were thought to be independent of caspases (Finn et al., 2000; Raff et al., 2002). Later work in Drosophila melanogaster (Kuo et al., 2006; Williams et al., 2006) and in mammalian neurons (Plachta et al., 2007; Nikolaev et al., 2009; Vohra et al., 2010) demonstrated that interfering with the apoptotic balance or the executor caspases can prevent or at least delay axonal degeneration. Simon et al. (2012) have found that a caspase 9 to caspase 3 cascade is crucial for axonal degeneration induced by NGF withdrawal, with caspase 6 activation playing a significant but subsidiary role. Upstream of the caspases, BCL-2 family members such as BAX and BCL-X_l are required (Nikolaev et al., 2009; Vohra et al., 2010). It is conceivable that the failure of ipRGCs in BAX-deficient mice to form appropriate connections to other cells in the retina (Chen et al., 2013) may be in part attributable to defective axonal degeneration. Surprisingly, Apaf1 appears not to be involved in this process (Cusack et al., 2013), suggesting that axon degeneration depends on the concerted activation of the intrinsic initiator complex in a different way from apoptosis.Strikingly, a series of recent studies showed that several caspases and components of the intrinsic pathway also affect normal synaptic physiology in adulthood (Fig. 3, A–D). Here, pro-apoptotic proteins are predominantly involved in weakening the synapses, whereas the anti-apoptotic proteins have been mainly associated with synaptic strengthening (Fig. 3 B). In particular, caspase 3 promotes long-term depression (LTD), a stimulation paradigm that results in a period of decreased synaptic transmission (Li et al., 2010), and also prevents long-term potentiation (LTP), the converse situation leading to strengthened synaptic transmission (Jo et al., 2011). Likewise, the proapoptotic BCL-2 family members BAX and BAD stimulate LTD (Jiao and Li, 2011). By contrast, the anti-apoptotic protein BCL-X_l increases synapse numbers and strength (H. Li et al., 2008), and the inhibitor of apoptosis protein (IAP) family member survivin was reported to be involved in LTP in the hippocampus (Iscru et al., 2013) and in activity-dependent gene regulation (O’Riordan et al., 2008).Open in a separate windowFigure 3.Canonical and noncanonical functions of the apoptotic machinery. (A) The apoptotic machinery is not only involved in eliminating cells destined to die, but is also a central player in refining neuronal connectivity, by regulating synaptic transmission and by generating the adult connectivity through axon pruning (Luo and O’Leary, 2005; Hyman and Yuan, 2012). But how the canonical and noncanonical roles of the apoptotic machinery are interlinked and spatially restricted is not well understood. (B) In the adult nervous system, the pro-apoptotic proteins BAX, caspase 9, and caspase 3 promote weakening of synapses (long-term depression [LTD]; Li et al., 2010; Jiao and Li, 2011; Jo et al., 2011), while the anti-apoptotic proteins Bcl-X_l and the IAP survivin promote synaptic strengthening (long-term potentiation [LTP]; Li et al., 2008a; Iscru et al., 2013). It is unclear how the activation of these pathways is restricted to a single synapse, but a recent review suggested that the proteasomal degradation of activated caspases may prevent their diffusion (Hyman and Yuan, 2012). (C) Caspase activation is now known to be required for axon pruning during development to generate the adult refined connectivity (Luo and O’Leary, 2005; Simon et al., 2012). Different pathways are activated depending on the stimulus leading to degeneration. Growth factor deprivation during development leads to activation the executor caspases 3 and 6 (Simon et al., 2012) through the intrinsic apoptotic pathway, although its core protein Apaf1 does not seem to be required for this process (Cusack et al., 2013). On the other hand, a traumatic injury leads to reduced influx of NMNAT2 into the axon, which negatively affects the stability and function of mitochondria and leads to an increased calcium concentration (Wang et al., 2012). The effector caspase, caspase 6, is dispensable for this form of axonal degeneration (Vohra et al., 2010; Simon et al., 2012). Regulatory proteins such as the IAPs and also the proteasome seem to play a role in limiting the extent of activation to the degenerating part of the axon (Wang et al., 2012; Cusack et al., 2013; Unsain et al., 2013). (D) Simplified schematic of the main pro- and anti-apoptotic components. DISC, death-induced signaling complex. IAP, inhibitor of apoptosis protein. See Fig. 1 for details.These findings indicate that the apoptotic machinery acts at different levels in the cell, ranging from driving sub-lethal degradation of a compartment (Fig. 3 C) and attenuating synaptic transmission at the neuronal network level (Fig. 3 B) to destroying the entire cell during development or in disease (Fig. 3 D). How the cell spatially restricts the extent of activation of the apoptotic machinery is yet unclear. For example, elimination of the somata of developing neurons after neurotrophin deprivation is preceded by axonal degeneration, but not all instances of axonal degeneration lead to the death of the neuron (Campenot, 1977; Raff et al., 2002). Local regulation of caspase activation by IAPs is well established as a means for ensuring the elimination of neuronal processes in D. melanogaster (Kuo et al., 2006; Williams et al., 2006). Recent findings suggest a similar role for IAP in mammalian neurons, where it limits caspase activation to the degenerating axon (Fig. 3 C; Cusack et al., 2013; Unsain et al., 2013). The spontaneous mutation Wallerian degeneration slow (WldS; Lunn et al., 1989) has been instrumental to understand that trauma-induced axon degeneration is a regulated process different from, and independent of, cell body degeneration (Wang et al., 2012), but also distinct from axon pruning (Hoopfer et al., 2006). Work on the chimeric protein encoded by the WldS mutation also led to the identification of the protein NMNAT2 (nicotinamide mononucleotide adenylyltransferase 2) as a labile axon survival factor (Gilley and Coleman, 2010). How the WldS chimeric protein and NMNAT2 result in axon protection is unclear, but several lines of evidence seem to converge on local regulation of mitochondrial function and motility (Avery et al., 2012; Fang et al., 2012).Related to the spatial limiting of apoptotic activity is the question of how a local source of neurotrophins leads to the rescue of a developing peripheral neuron. When neurons encounter a source of neurotrophins, only the receptors close to the target will be activated, whereas the others, located further away, are not. The cell, therefore, needs to integrate a pro-survival signal from the activated receptors, and death-inducing signals from the nonactivated dependence receptors. The continued signaling of activated neurotrophin receptors that are retrogradely transported to the soma (Grimes et al., 1996; Howe et al., 2001; Wu et al., 2001; Harrington et al., 2011) likely play a role in counteracting the pro-apoptotic signaling proximal to the source of neurotrophins. It will be interesting to investigate whether similar mechanisms play a role in axon pruning and traumatic axon degeneration as well.

Programmed cell death in the adult brain

Most of the nervous system becomes post-mitotic early in development. In rodents, two brain areas retain the capacity to generate new neurons in the adult: the sub-ventricular zone, which generates neurons that migrate toward the olfactory bulb, and the sub-granular zone of the dentate gyrus of the hippocampus, where neurons are generated that integrate locally. Similar to what is observed during embryonic development, these adult-generated neurons are produced in excess, and a large fraction undergoes apoptosis when contacting its designated targets (Petreanu and Alvarez-Buylla, 2002; Kempermann et al., 2003; Ninkovic et al., 2007). Preventing apoptosis of adult-generated neurons in the olfactory bulb only has limited functional consequences (Kim et al., 2007), whereas a similar maneuver in the dentate gyrus does lead to impaired performance in memory tasks (Kim et al., 2009). Why superfluous hippocampal neurons would need to be eliminated for proper function is a matter of speculation, but may be linked with the fact that these are excitatory projection neurons, whereas in the olfactory bulb only axon-less inhibitory granule cells are integrated. The extent of survival in both these areas critically depends on the activity of the neuronal network in which these newly born neurons have to integrate (Petreanu and Alvarez-Buylla, 2002; Kempermann et al., 2006; Ninkovic et al., 2007). In this context, BDNF, the expression level of which is well known to be regulated by network activity, supports the survival of young adult–generated neurons and possibly even stimulates the proliferation of neural progenitors (Y. Li et al., 2008; Waterhouse et al., 2012). Interestingly, in young adult mouse mutants that exhibit spontaneous epileptic seizures, significantly higher levels of BDNF have been measured (Lavebratt et al., 2006; Heyden et al., 2011). Concomitantly, the entire hippocampal formation is considerably enlarged by as much as 40% (Lavebratt et al., 2006; Angenstein et al., 2007), which in turn is dependent on the epileptic seizures (Lavebratt et al., 2006). Whether or not there is a causal relationship between increased BDNF levels and hippocampal volume remains to be established.

Conclusion

Now that is has become clear that action of the apoptotic machinery can be limited spatially and temporally, several questions need to be addressed: how do neurons integrate intrinsic and extrinsic pro- and anti-apoptotic signals; and how they are spatially restricted to allow degradation of a dendrite or axon, or modulation of synaptic transmission? Another important issue is the regulation of cell death by intrinsic mechanisms in the central nervous system of vertebrates, not least because programmed cell death is observed in the CNS in a number of neurodegenerative diseases (Vila and Przedborski, 2003). Indeed, several of the central apoptotic components discussed here are also involved in these disorders (Hyman and Yuan, 2012). New insights in the regulation of programmed cell death in the developing nervous system may therefore continue to help to better understand the pathophysiological mechanisms of neurodegenerative disorders.     

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.     

The Apical Actin Fringe Contributes to Localized Cell Wall Deposition and Polarized Growth in the Lily Pollen Tube

In lily (Lilium formosanum) pollen tubes, pectin, a major component of the cell wall, is delivered through regulated exocytosis. The targeted transport and secretion of the pectin-containing vesicles may be controlled by the cortical actin fringe at the pollen tube apex. Here, we address the role of the actin fringe using three different inhibitors of growth: brefeldin A, latrunculin B, and potassium cyanide. Brefeldin A blocks membrane trafficking and inhibits exocytosis in pollen tubes; it also leads to the degradation of the actin fringe and the formation of an aggregate of filamentous actin at the base of the clear zone. Latrunculin B, which depolymerizes filamentous actin, markedly slows growth but allows focused pectin deposition to continue. Of note, the locus of deposition shifts frequently and correlates with changes in the direction of growth. Finally, potassium cyanide, an electron transport chain inhibitor, briefly stops growth while causing the actin fringe to completely disappear. Pectin deposition continues but lacks focus, instead being delivered in a wide arc across the pollen tube tip. These data support a model in which the actin fringe contributes to the focused secretion of pectin to the apical cell wall and, thus, to the polarized growth of the pollen tube.Pollen tubes provide an excellent model for studying the molecular and physiological processes that lead to polarized cell growth. Because all plant cell growth results from the regulated yielding of the cell wall in response to uniform turgor pressure (Winship et al., 2010; Rojas et al., 2011), the cell wall of the pollen tube must yield only at a particular spot: the cell apex, or tip. To accomplish the extraordinary growth rates seen in many species, and to balance the thinning of the apical wall due to rapid expansion, the pollen tube delivers prodigious amounts of wall material, largely methoxylated pectins, to the tip in a coordinated manner. Recent studies suggest that the targeted exocytosis increases the extensibility of the cell wall matrix at the tip, which then yields to the existing turgor pressure, permitting the tip to extend or grow (McKenna et al., 2009; Hepler et al., 2013). There are many factors that influence exocytosis in growing pollen tubes; in this study, we investigate the role of the apical actin fringe.For many years, it has been known that an actin structure exists near the pollen tube tip, yet its exact form has been a matter of some contention (Kost et al., 1998; Lovy-Wheeler et al., 2005; Wilsen et al., 2006; Cheung et al., 2008; Vidali et al., 2009; Qu et al., 2013). The apical actin structure has been variously described as a fringe, a basket, a collar, or a mesh. Using rapid freeze fixation of lily (Lilium formosanum) pollen tubes followed by staining with anti-actin antibodies, the structure appears as a dense fringe of longitudinally oriented microfilaments, beginning 1 to 5 µm behind the apex and extending 5 to 10 µm basally. The actin filaments are positioned in the cortical cytoplasm close to the plasma membrane (Lovy-Wheeler et al., 2005). More recently, we used Lifeact-mEGFP, a probe that consistently labels this palisade of longitudinally oriented microfilaments in living cells (Vidali et al., 2009; Fig. 1A, left column). For the purposes of this study, we will refer to this apical organization of actin as a fringe.Open in a separate windowFigure 1.The actin fringe and the thickened pollen tube tip wall are stable, although dynamic, structures during pollen tube growth. A, The left column shows a pollen tube transformed with Lifeact-mEGFP imaged with a spinning-disc confocal microscope. Maximal projections from every 15 s are shown. The right column shows epifluorescence images of a pollen tube stained with PI. Again, images captured every 15 s are shown. Bars = 10 μm. B, The data from the pollen tube in A expressing Lifeact-mEGFP were subjected to kymograph analysis using an 11-pixel strip along the image’s midline. C, The first three frames from the pollen tube in A and B were assigned the colors red, blue, and green, respectively, and then overlaid. Areas with white show the overlap of all three. The fringe is stable, but most of its constituent actin is not shared between frames.Many lines of evidence demonstrate that actin is required for pollen tube growth. Latrunculin B (LatB), which blocks actin polymerization, inhibits pollen tube growth and disrupts the cortical fringe at concentrations as low as 2 nm. Higher concentrations are needed to block pollen grain germination and cytoplasmic streaming (Gibbon et al., 1999; Vidali et al., 2001). Actin-binding proteins, including actin depolymerizing factor-cofilin, formin, profilin, and villin, and signaling proteins, such as Rho-of-Plants (ROP) GTPases and their effectors (ROP interacting crib-containing proteins [RICs]), also have been shown to play critical roles in growth and actin dynamics (Fu et al., 2001; Vidali et al., 2001; Allwood et al., 2002; Chen et al., 2002; Cheung and Wu, 2004; McKenna et al., 2004; Gu et al., 2005; Ye et al., 2009; Cheung et al., 2010; Staiger et al., 2010; Zhang et al., 2010a; Qu et al., 2013; van Gisbergen and Bezanilla, 2013).Our understanding of the process of exocytosis and pollen tube elongation has been influenced by ultrastructural images of pollen tube tips, which reveal an apical zone dense with vesicles (Cresti et al., 1987; Heslop-Harrison, 1987; Lancelle et al., 1987; Steer and Steer, 1989; Lancelle and Hepler, 1992; Derksen et al., 1995). It has long been assumed that these represent exocytotic vesicles destined to deliver new cell wall material. This model of polarized secretion has been challenged in recent years in studies using FM dyes. Two groups have suggested that exocytosis occurs in a circumpolar annular zone (Bove et al., 2008; Zonia and Munnik, 2008). However, other studies, using fluorescent beads attached to the cell surface, indicate that the maximal rate of expansion, and of necessity the greatest deposition of cell wall material, occurs at the apex along the polar axis of the tube (Dumais et al., 2006; Rojas et al., 2011). Similarly, our experiments with propidium iodide (PI; McKenna et al., 2009; Rounds et al., 2011a) and pectin methyl esterase fused to GFP (McKenna et al., 2009) show that the wall is thickest at the very tip and suggest that wall materials are deposited at the polar axis, consistent with the initial model of exocytosis (Lancelle and Hepler, 1992). Experiments using tobacco (Nicotiana tabacum) pollen and a receptor-like kinase fused to GFP also indicate that exocytosis occurs largely at the apical polar axis (Lee et al., 2008).Many researchers argue that apical actin is critical for exocytosis (Lee et al., 2008; Cheung et al., 2010; Qin and Yang, 2011; Yan and Yang, 2012). More specifically, recent work suggests that the fringe participates in targeting vesicles and thereby contributes to changes in growth direction (Kroeger et al., 2009; Bou Daher and Geitmann, 2011; Dong et al., 2012). In this article, using three different inhibitors, namely brefeldin A (BFA), LatB, and potassium cyanide (KCN), we test the hypothesis that polarized pectin deposition in pollen tubes requires the actin fringe. Our data show that during normal growth, pectin deposition is focused to the apex along the polar axis of the tube. However, when growth is modulated, different end points arise, depending on the inhibitor. With BFA, exocytosis stops completely, and the fringe disappears, with the appearance of an actin aggregate at the base of the clear zone. LatB, as shown previously (Vidali et al., 2009), incompletely degrades the actin fringe and leaves a rim of F-actin around the apical dome. Here, we show that, in the presence of LatB, pectin deposition continues, with the focus of this activity shifting in position frequently as the slowly elongating pollen tube changes direction. With KCN, the actin fringe degrades completely, but exocytosis continues and becomes depolarized, with pectin deposits now occurring across a wide arc of the apical dome. This dome often swells as deposition continues, only stopping once normal growth resumes. Taken together, these results support a role for the actin fringe in controlling the polarity of growth in the lily pollen tube.     

Calcineurin B-Like Protein-Interacting Protein Kinase CIPK21 Regulates Osmotic and Salt Stress Responses in Arabidopsis

The role of calcium-mediated signaling has been extensively studied in plant responses to abiotic stress signals. Calcineurin B-like proteins (CBLs) and CBL-interacting protein kinases (CIPKs) constitute a complex signaling network acting in diverse plant stress responses. Osmotic stress imposed by soil salinity and drought is a major abiotic stress that impedes plant growth and development and involves calcium-signaling processes. In this study, we report the functional analysis of CIPK21, an Arabidopsis (Arabidopsis thaliana) CBL-interacting protein kinase, ubiquitously expressed in plant tissues and up-regulated under multiple abiotic stress conditions. The growth of a loss-of-function mutant of CIPK21, cipk21, was hypersensitive to high salt and osmotic stress conditions. The calcium sensors CBL2 and CBL3 were found to physically interact with CIPK21 and target this kinase to the tonoplast. Moreover, preferential localization of CIPK21 to the tonoplast was detected under salt stress condition when coexpressed with CBL2 or CBL3. These findings suggest that CIPK21 mediates responses to salt stress condition in Arabidopsis, at least in part, by regulating ion and water homeostasis across the vacuolar membranes.Drought and salinity cause osmotic stress in plants and severely affect crop productivity throughout the world. Plants respond to osmotic stress by changing a number of cellular processes (Xiong et al., 1999; Xiong and Zhu, 2002; Bartels and Sunkar, 2005; Boudsocq and Lauriére, 2005). Some of these changes include activation of stress-responsive genes, regulation of membrane transport at both plasma membrane (PM) and vacuolar membrane (tonoplast) to maintain water and ionic homeostasis, and metabolic changes to produce compatible osmolytes such as Pro (Stewart and Lee, 1974; Krasensky and Jonak, 2012). It has been well established that a specific calcium (Ca²⁺) signature is generated in response to a particular environmental stimulus (Trewavas and Malhó, 1998; Scrase-Field and Knight, 2003; Luan, 2009; Kudla et al., 2010). The Ca²⁺ changes are primarily perceived by several Ca²⁺ sensors such as calmodulin (Reddy, 2001; Luan et al., 2002), Ca²⁺-dependent protein kinases (Harper and Harmon, 2005), calcineurin B-like proteins (CBLs; Luan et al., 2002; Batistič and Kudla, 2004; Pandey, 2008; Luan, 2009; Sanyal et al., 2015), and other Ca²⁺-binding proteins (Reddy, 2001; Shao et al., 2008) to initiate various cellular responses.Plant CBL-type Ca²⁺ sensors interact with and activate CBL-interacting protein kinases (CIPKs) that phosphorylate downstream components to transduce Ca²⁺ signals (Liu et al., 2000; Luan et al., 2002; Batistič and Kudla, 2004; Luan, 2009). In several plant species, multiple members have been identified in the CBL and CIPK family (Luan et al., 2002; Kolukisaoglu et al., 2004; Pandey, 2008; Batistič and Kudla, 2009; Weinl and Kudla, 2009; Pandey et al., 2014). Involvement of specific CBL-CIPK pair to decode a particular type of signal entails the alternative and selective complex formation leading to stimulus-response coupling (D’Angelo et al., 2006; Batistič et al., 2010).Several CBL and CIPK family members have been implicated in plant responses to drought, salinity, and osmotic stress based on genetic analysis of Arabidopsis (Arabidopsis thaliana) mutants (Zhu, 2002; Cheong et al., 2003, 2007; Kim et al., 2003; Pandey et al., 2004, 2008; D’Angelo et al., 2006; Qin et al., 2008; Tripathi et al., 2009; Held et al., 2011; Tang et al., 2012; Drerup et al., 2013; Eckert et al., 2014). A few CIPKs have also been functionally characterized by gain-of-function approach in crop plants such as rice (Oryza sativa), pea (Pisum sativum), and maize (Zea mays) and were found to be involved in osmotic stress responses (Mahajan et al., 2006; Xiang et al., 2007; Yang et al., 2008; Tripathi et al., 2009; Zhao et al., 2009; Cuéllar et al., 2010).In this report, we examined the role of the Arabidopsis CIPK21 gene in osmotic stress response by reverse genetic analysis. The loss-of-function mutant plants became hypersensitive to salt and mannitol stress conditions, suggesting that CIPK21 is involved in the regulation of osmotic stress response in Arabidopsis. These findings are further supported by an enhanced tonoplast targeting of the cytoplasmic CIPK21 through interaction with the vacuolar Ca²⁺ sensors CBL2 and CBL3 under salt stress condition.     

Characterization of the Possible Roles for B Class MADS Box Genes in Regulation of Perianth Formation in Orchid

To investigate sepal/petal/lip formation in Oncidium Gower Ramsey, three paleoAPETALA3 genes, O. Gower Ramsey MADS box gene5 (OMADS5; clade 1), OMADS3 (clade 2), and OMADS9 (clade 3), and one PISTILLATA gene, OMADS8, were characterized. The OMADS8 and OMADS3 mRNAs were expressed in all four floral organs as well as in vegetative leaves. The OMADS9 mRNA was only strongly detected in petals and lips. The mRNA for OMADS5 was only strongly detected in sepals and petals and was significantly down-regulated in lip-like petals and lip-like sepals of peloric mutant flowers. This result revealed a possible negative role for OMADS5 in regulating lip formation. Yeast two-hybrid analysis indicated that OMADS5 formed homodimers and heterodimers with OMADS3 and OMADS9. OMADS8 only formed heterodimers with OMADS3, whereas OMADS3 and OMADS9 formed homodimers and heterodimers with each other. We proposed that sepal/petal/lip formation needs the presence of OMADS3/8 and/or OMADS9. The determination of the final organ identity for the sepal/petal/lip likely depended on the presence or absence of OMADS5. The presence of OMADS5 caused short sepal/petal formation. When OMADS5 was absent, cells could proliferate, resulting in the possible formation of large lips and the conversion of the sepal/petal into lips in peloric mutants. Further analysis indicated that only ectopic expression of OMADS8 but not OMADS5/9 caused the conversion of the sepal into an expanded petal-like structure in transgenic Arabidopsis (Arabidopsis thaliana) plants.The ABCDE model predicts the formation of any flower organ by the interaction of five classes of homeotic genes in plants (Yanofsky et al., 1990; Jack et al., 1992; Mandel et al., 1992; Goto and Meyerowitz, 1994; Jofuku et al., 1994; Pelaz et al., 2000, 2001; Theißen and Saedler, 2001; Pinyopich et al., 2003; Ditta et al., 2004; Jack, 2004). The A class genes control sepal formation. The A, B, and E class genes work together to regulate petal formation. The B, C, and E class genes control stamen formation. The C and E class genes work to regulate carpel formation, whereas the D class gene is involved in ovule development. MADS box genes seem to have a central role in flower development, because most ABCDE genes encode MADS box proteins (Coen and Meyerowitz, 1991; Weigel and Meyerowitz, 1994; Purugganan et al., 1995; Rounsley et al., 1995; Theißen and Saedler, 1995; Theißen et al., 2000; Theißen, 2001).The function of B group genes, such as APETALA3 (AP3) and PISTILLATA (PI), has been thought to have a major role in specifying petal and stamen development (Jack et al., 1992; Goto and Meyerowitz, 1994; Krizek and Meyerowitz, 1996; Kramer et al., 1998; Hernandez-Hernandez et al., 2007; Kanno et al., 2007; Whipple et al., 2007; Irish, 2009). In Arabidopsis (Arabidopsis thaliana), mutation in AP3 or PI caused identical phenotypes of second whorl petal conversion into a sepal structure and third flower whorl stamen into a carpel structure (Bowman et al., 1989; Jack et al., 1992; Goto and Meyerowitz, 1994). Similar homeotic conversions for petal and stamen were observed in the mutants of the AP3 and PI orthologs from a number of core eudicots such as Antirrhinum majus, Petunia hybrida, Gerbera hybrida, Solanum lycopersicum, and Nicotiana benthamiana (Sommer et al., 1990; Tröbner et al., 1992; Angenent et al., 1993; van der Krol et al., 1993; Yu et al., 1999; Liu et al., 2004; Vandenbussche et al., 2004; de Martino et al., 2006), from basal eudicot species such as Papaver somniferum and Aquilegia vulgaris (Drea et al., 2007; Kramer et al., 2007), as well as from monocot species such as Zea mays and Oryza sativa (Ambrose et al., 2000; Nagasawa et al., 2003; Prasad and Vijayraghavan, 2003; Yadav et al., 2007; Yao et al., 2008). This indicated that the function of the B class genes AP3 and PI is highly conserved during evolution.It has been thought that B group genes may have arisen from an ancestral gene through multiple gene duplication events (Doyle, 1994; Theißen et al., 1996, 2000; Purugganan, 1997; Kramer et al., 1998; Kramer and Irish, 1999; Lamb and Irish, 2003; Kim et al., 2004; Stellari et al., 2004; Zahn et al., 2005; Hernandez-Hernandez et al., 2007). In the gymnosperms, there was a single putative B class lineage that duplicated to generate the paleoAP3 and PI lineages in angiosperms (Kramer et al., 1998; Theißen et al., 2000; Irish, 2009). The paleoAP3 lineage is composed of AP3 orthologs identified in lower eudicots, magnolid dicots, and monocots (Kramer et al., 1998). Genes in this lineage contain the conserved paleoAP3- and PI-derived motifs in the C-terminal end of the proteins, which have been thought to be characteristics of the B class ancestral gene (Kramer et al., 1998; Tzeng and Yang, 2001; Hsu and Yang, 2002). The PI lineage is composed of PI orthologs that contain a highly conserved PI motif identified in most plant species (Kramer et al., 1998). Subsequently, there was a second duplication at the base of the core eudicots that produced the euAP3 and TM6 lineages, which have been subject to substantial sequence changes in eudicots during evolution (Kramer et al., 1998; Kramer and Irish, 1999). The paleoAP3 motif in the C-terminal end of the proteins was retained in the TM6 lineage and replaced by a conserved euAP3 motif in the euAP3 lineage of most eudicot species (Kramer et al., 1998). In addition, many lineage-specific duplications for paleoAP3 lineage have occurred in plants such as orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009), Ranunculaceae, and Ranunculales (Kramer et al., 2003; Di Stilio et al., 2005; Shan et al., 2006; Kramer, 2009).Unlike the A or C class MADS box proteins, which form homodimers that regulate flower development, the ability of B class proteins to form homodimers has only been reported in gymnosperms and in the paleoAP3 and PI lineages of some monocots. For example, LMADS1 of the lily Lilium longiflorum (Tzeng and Yang, 2001), OMADS3 of the orchid Oncidium Gower Ramsey (Hsu and Yang, 2002), and PeMADS4 of the orchid Phalaenopsis equestris (Tsai et al., 2004) in the paleoAP3 lineage, LRGLOA and LRGLOB of the lily Lilium regale (Winter et al., 2002), TGGLO of the tulip Tulipa gesneriana (Kanno et al., 2003), and PeMADS6 of the orchid P. equestris (Tsai et al., 2005) in the PI lineage, and GGM2 of the gymnosperm Gnetum gnemon (Winter et al., 1999) were able to form homodimers that regulate flower development. Proteins in the euAP3 lineage and in most paleoAP3 lineages were not able to form homodimers and had to interact with PI to form heterodimers in order to regulate petal and stamen development in various plant species (Schwarz-Sommer et al., 1992; Tröbner et al., 1992; Riechmann et al., 1996; Moon et al., 1999; Winter et al., 2002; Kanno et al., 2003; Vandenbussche et al., 2004; Yao et al., 2008). In addition to forming dimers, AP3 and PI were able to interact with other MADS box proteins, such as SEPALLATA1 (SEP1), SEP2, and SEP3, to regulate petal and stamen development (Pelaz et al., 2000; Honma and Goto, 2001; Theißen and Saedler, 2001; Castillejo et al., 2005).Orchids are among the most important plants in the flower market around the world, and research on MADS box genes has been reported for several species of orchids during the past few years (Lu et al., 1993, 2007; Yu and Goh, 2000; Hsu and Yang, 2002; Yu et al., 2002; Hsu et al., 2003; Tsai et al., 2004, 2008; Xu et al., 2006; Guo et al., 2007; Kim et al., 2007; Chang et al., 2009). Unlike the flowers in eudicots, the nearly identical shape of the sepals and petals as well as the production of a unique lip in orchid flowers make them a very special plant species for the study of flower development. Four clades (1–4) of genes in the paleoAP3 lineage have been identified in several orchids (Hsu and Yang, 2002; Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009; Mondragón-Palomino et al., 2009). Several works have described the possible interactions among these four clades of paleoAP3 genes and one PI gene that are involved in regulating the differentiation and formation of the sepal/petal/lip of orchids (Tsai et al., 2004; Kim et al., 2007; Mondragón-Palomino and Theißen, 2008, 2009). However, the exact mechanism that involves the orchid B class genes remains unclear and needs to be clarified by more experimental investigations.O. Gower Ramsey is a popular orchid with important economic value in cut flower markets. Only a few studies have been reported on the role of MADS box genes in regulating flower formation in this plant species (Hsu and Yang, 2002; Hsu et al., 2003; Chang et al., 2009). An AP3-like MADS gene that regulates both floral formation and initiation in transgenic Arabidopsis has been reported (Hsu and Yang, 2002). In addition, four AP1/AGAMOUS-LIKE9 (AGL9)-like MADS box genes have been characterized that show novel expression patterns and cause different effects on floral transition and formation in Arabidopsis (Hsu et al., 2003; Chang et al., 2009). Compared with other orchids, the production of a large and well-expanded lip and five small identical sepals/petals makes O. Gower Ramsey a special case for the study of the diverse functions of B class MADS box genes during evolution. Therefore, the isolation of more B class MADS box genes and further study of their roles in the regulation of perianth (sepal/petal/lip) formation during O. Gower Ramsey flower development are necessary. In addition to the clade 2 paleoAP3 gene OMADS3, which was previously characterized in our laboratory (Hsu and Yang, 2002), three more B class MADS box genes, OMADS5, OMADS8, and OMADS9, were characterized from O. Gower Ramsey in this study. Based on the different expression patterns and the protein interactions among these four orchid B class genes, we propose that the presence of OMADS3/8 and/or OMADS9 is required for sepal/petal/lip formation. Further sepal and petal formation at least requires the additional presence of OMADS5, whereas large lip formation was seen when OMADS5 expression was absent. Our results provide a new finding and information pertaining to the roles for orchid B class MADS box genes in the regulation of sepal/petal/lip formation.     

Focus Issue on Plant Cell Walls: A Comprehensive Toolkit of Plant Cell Wall Glycan-Directed Monoclonal Antibodies

A collection of 130 new plant cell wall glycan-directed monoclonal antibodies (mAbs) was generated with the aim of facilitating in-depth analysis of cell wall glycans. An enzyme-linked immunosorbent assay-based screen against a diverse panel of 54 plant polysaccharides was used to characterize the binding patterns of these new mAbs, together with 50 other previously generated mAbs, against plant cell wall glycans. Hierarchical clustering analysis was used to group these mAbs based on the polysaccharide recognition patterns observed. The mAb groupings in the resulting cladogram were further verified by immunolocalization studies in Arabidopsis (Arabidopsis thaliana) stems. The mAbs could be resolved into 19 clades of antibodies that recognize distinct epitopes present on all major classes of plant cell wall glycans, including arabinogalactans (both protein- and polysaccharide-linked), pectins (homogalacturonan, rhamnogalacturonan I), xyloglucans, xylans, mannans, and glucans. In most cases, multiple subclades of antibodies were observed to bind to each glycan class, suggesting that the mAbs in these subgroups recognize distinct epitopes present on the cell wall glycans. The epitopes recognized by many of the mAbs in the toolkit, particularly those recognizing arabinose- and/or galactose-containing structures, are present on more than one glycan class, consistent with the known structural diversity and complexity of plant cell wall glycans. Thus, these cell wall glycan-directed mAbs should be viewed and utilized as epitope-specific, rather than polymer-specific, probes. The current world-wide toolkit of approximately 180 glycan-directed antibodies from various laboratories provides a large and diverse set of probes for studies of plant cell wall structure, function, dynamics, and biosynthesis.Cell walls play important roles in the structure, physiology, growth, and development of plants (Carpita and Gibeaut, 1993). Plant cell wall materials are also important sources of human and animal nutrition, natural textile fibers, paper and wood products, and raw materials for biofuel production (Somerville, 2007). Many genes thought to be responsible for plant wall biosynthesis and modification have been identified (Burton et al., 2005; Lerouxel et al., 2006; Mohnen et al., 2008), and 15% of the Arabidopsis (Arabidopsis thaliana) genome is likely devoted to these functions (Carpita et al., 2001). However, phenotypic analysis in plants carrying cell wall-related mutations has proven particularly difficult. First, cell wall-related genes are often expressed differentially and at low levels between cells of different tissues (Sarria et al., 2001). Also, plants have compensatory mechanisms to maintain wall function in the absence of a particular gene (Somerville et al., 2004). Thus, novel tools and approaches are needed to characterize wall structures and the genes responsible for their synthesis and modification.Monoclonal antibodies (mAbs) developed against cell wall polymers have emerged as an important tool for the study of plant cell wall structure and function (Knox, 2008). Previous studies have utilized mAbs that bind epitopes present on rhamnogalacturonan I (RG-I; Freshour et al., 1996; Jones et al., 1997; Willats et al., 1998; McCartney et al., 2000; Clausen et al., 2004; Altaner et al., 2007), homogalacturonan (Willats et al., 2001; Clausen et al., 2003), xylogalacturonan (Willats et al., 2004), xylans and arabinoxylans (McCartney et al., 2005), xyloglucan (Freshour et al., 1996, 2003; Marcus et al., 2008), arabinogalactan(protein) (Pennell et al., 1991; Puhlmann et al., 1994; Dolan et al., 1995; Smallwood et al., 1996), and extensins (Smallwood et al., 1995) to localize these epitopes in plant cells and tissues. In addition, mAbs have been used to characterize plants carrying mutations in genes thought to be associated with cell wall biosynthesis and metabolism (Orfila et al., 2001; Seifert, 2004; Persson et al., 2007; Cavalier et al., 2008; Zabotina et al., 2008). Despite their utility, the available set of mAbs against carbohydrate structures is relatively small given the structural complexity of wall polymers (Ridley et al., 2001; O''Neill and York, 2003), and knowledge of their epitope specificity is limited. Thus, additional mAbs specific to diverse epitope structures and methods for rapid epitope characterization are needed (Somerville et al., 2004).Here, we report the generation of 130 new mAbs that bind to diverse epitopes present on a broad spectrum of plant cell wall glycans. In addition, approximately 50 previously reported or generated mAbs were included in the ELISA-based screens used to group the antibodies according to their binding patterns against a diverse panel of 54 polysaccharides. The resulting ELISA data were analyzed by hierarchical clustering to illustrate the relationships between the available mAbs. Nineteen groups of mAbs were identified from the clustering analysis. Some initial information regarding possible epitopes recognized by some of these antibodies could be inferred from the clustering analysis.     

Cell biology in development: The cell biology of planar cell polarity

Planar cell polarity (PCP) refers to the coordinated alignment of cell polarity across the tissue plane. Key to the establishment of PCP is asymmetric partitioning of cortical PCP components and intercellular communication to coordinate polarity between neighboring cells. Recent progress has been made toward understanding how protein transport, endocytosis, and intercellular interactions contribute to asymmetric PCP protein localization. Additionally, the functions of gradients and mechanical forces as global cues that bias PCP orientation are beginning to be elucidated. Together, these findings are shedding light on how global cues integrate with local cell interactions to organize cellular polarity at the tissue level.The collective alignment of cell polarity across the tissue plane is a phenomenon known as planar cell polarity (PCP). Exemplified by the uniform orientation of bristles covering the insect epidermis or of the hairs covering the mammalian body surface (Fig. 1 A), PCP patterns can align over thousands, even billions of cells. This phenomenon is controlled by the so-called PCP pathway, which integrates global directional cues to produce locally polarized cell behaviors. There has been a recent surge in interest in PCP after discoveries that various processes such as vertebrate gastrulation, mammalian ear patterning and hearing, and neural tube closure all require a conserved set of PCP genes (Heisenberg et al., 2000; Tada and Smith, 2000; Wallingford et al., 2000; Kibar et al., 2001; Murdoch et al., 2001; Curtin et al., 2003; Montcouquiol et al., 2003; Copley et al., 2013). Since that time, the PCP pathway has been found to coordinate cell behaviors in numerous diverse settings including polarized ciliary beating in the trachea and brain ventricles (Tissir et al., 2010; Vladar et al., 2012), oriented cell divisions (Gong et al., 2004; Baena-López et al., 2005; Ségalen et al., 2010; Mao et al., 2011), lung branching (Yates et al., 2010), and hair follicle alignment (Guo et al., 2004; Devenport and Fuchs, 2008), to name a few (Fig. 1). Genetic disruptions to PCP cause severe developmental abnormalities in vertebrates, notably neural tube defects, left/right patterning defects, and ciliopathies, which highlights the essential requirement for PCP in development (Kibar et al., 2001; Murdoch et al., 2001; Curtin et al., 2003; Wang et al., 2006a,b; Kim et al., 2010; Song et al., 2010).Open in a separate windowFigure 1.Planar cell polarity and the core PCP components. (A and B) The Drosophila wing blade and mammalian epidermis illustrate the phenomenon of PCP. In both cases, hairs point in a single direction along the tissue axis, where they align locally with neighboring hairs and globally across the tissue. Whereas Drosophila wing hairs are produced by single cells, mammalian hairs emerge from multicellular hair follicles, which orient as a unit. A conserved PCP pathway controls the collective alignment of both types of structures. (C) Core PCP components localize to the plasma membrane and asymmetrically segregate along the epithelial plane as indicated.Like many types of cell polarity, the establishment of PCP involves (1) a global orienting cue, (2) asymmetric segregation of dedicated polarity proteins, and (3) translation of polarity information into polarized outputs. But unlike other types of cell polarity, the PCP mechanisms we currently understand involve coupling between adjacent cells, allowing for the alignment of polarity over many cell distances.First described in insects and then genetically dissected in Drosophila melanogaster, PCP was long confined to the realm of experimental embryology and genetics until the discovery that the protein products of several PCP genes were localized asymmetrically within the cell, thrusting PCP into the domain of cell biology (for review see Strutt and Strutt, 2009). The challenge to understanding PCP on a molecular level is that long-range PCP is, in essence, an in vivo phenomenon that is difficult to recapitulate in a tissue culture dish. However, recent advances in imaging technology combined with increasingly sophisticated genetic tools are helping us to decipher the in vivo cell biology of PCP. In this review, I highlight some of the recent advances made toward understanding the cell biology underlying the establishment of coordinated polarized cell behaviors. For clarity, I limit discussions of PCP phenomena that meet the definition of PCP proposed by Goodrich and Strutt (2011): namely, that “cell–cell communication causes two or more cells to adopt coordinated polarity” in a process that is mechanistically “dependent upon planar polarity proteins.” Other aligned cellular patterns or examples of noncanonical Wnt signaling, sometimes described as “Wnt/PCP” signaling, will not be discussed.

PCP components and molecular asymmetries

Two molecular systems control PCP behavior, the “core” and the “Fat–Dachsous (Ft–Ds)” PCP pathways. A key feature of both is the asymmetric distribution of their constituents (Fig. 2). The core PCP pathway is composed of the multipass transmembrane proteins Frizzled (Fz), Van Gogh (Vang; also known as Strabismus/Stbm), and Flamingo (Fmi; also known as Starry night/Stan), and the cytosolic components Dishevelled (Dsh), Prickle (Pk), and Diego (Dgo). On one edge of the cell reside Fz, Dsh, and Dgo, and on the opposite side lie Vang and Pk (Figs. 1 C and 2 B; Axelrod, 2001; Strutt, 2001; Feiguin et al., 2001; Tree et al., 2002; Bastock et al., 2003). The atypical cadherin, Fmi, resides on both sides, where it forms homodimers between neighboring cells (Usui et al., 1999; Shimada et al., 2001). These molecular asymmetries are observed in sensory hair cells of the vertebrate inner ear (Wang et al., 2005, 2006a,b; Montcouquiol et al., 2006; Deans et al., 2007; Song et al., 2010), the mammalian epidermis (Devenport and Fuchs, 2008; Devenport et al., 2011), brain ventricles (Tissir et al., 2010), and trachea (Vladar et al., 2012). Mutations in core PCP components lead to a loss or randomization of polarity and misalignment of cellular structures along the tissues axis.Open in a separate windowFigure 2.Asymmetric localization of PCP components. (A) PCP asymmetry develops progressively from an initially uniform distribution of core PCP proteins. Fz, Dsh, and Dgo (red) localize to the distal/posterior edge, whereas Vang and Pk (turquoise) localize to the proximal/anterior side. Fmi (dark blue) localizes to both sides, where it forms homodimers between neighboring cells. (B) Feedback interactions between core PCP components. A Fz–Fmi complex interacts preferentially with a Vang–Fmi complex between cells, whereas proximal and distal complexes antagonize one another within the cell. (C) Ds and Fj are expressed in opposing gradients in the Drosophila wing blade. Fj positively modulates Ft activity, leading to a gradient of Ft activity across the wing (not depicted). (D) Graded expression of Ds and Fj leads to asymmetric cellular localization of Ds and Ft, which form heterodimers between adjacent cells. Dachs, a downstream component of the Ft–Ds pathway, also localizes asymmetrically in association with Ds.The Ft–Ds pathway includes the large protocadherins Ft and Ds and the Golgi resident transmembrane kinase, Four-jointed (Fj; for review see Matis and Axelrod, 2013; Thomas and Strutt, 2012). Similar to the core system, Ft–Ds also displays molecular asymmetries in flies. Ds and its ligand Ft accumulate on opposite cell edges, where they form intercellular heterophilic interactions (Fig. 2 D; Matakatsu and Blair 2004; Ambegaonkar et al., 2012; Brittle et al., 2012). Unlike the core components, Ds and Fj are expressed in complementary gradients in the Drosophila eye and developing wing, which contribute to the cellular asymmetries of Ds and Ft (Fig. 2, C and D). Whether Ft–Ds–Fj gradients and asymmetries are conserved in vertebrate systems has yet to be determined.

Segregation of cortical polarity proteins: Shaking hands with the enemy

The asymmetric segregation of Fz–Dsh–Fmi and Vang–Pk–Fmi complexes to opposite sides of the cell relies on their mutual exclusion intracellularly and their preferential binding between neighboring cells (Fig. 2 B; for review see Strutt and Strutt, 2009). There is mutual interdependence among core PCP components for their asymmetric localization. Depletion of any one core PCP component results in a loss of asymmetry of all the others. In addition, PCP asymmetry develops progressively from initially uniform distributions (Fig. 2 A). Thus, PCP asymmetry can be thought of not as a simple hierarchy of interactions, but the result of feedback amplification of an initial directional bias.

Intercellular interactions.

PCP requires cell–cell communication, mediated by the transmembrane components of the core system, where it is thought that Fz–Fmi on one cell interacts with Vang–Fmi on its neighbor. These interactions are best understood in the Drosophila wing blade, where PCP controls the alignment of wing hairs along the proximal–distal axis (Figs. 1 A and 2, A and B). In the wing, Vang–Pk localize to the proximal face of each cell, whereas Fz–Dsh–Dgo localize distally adjacent to the wing hair (Figs. 1 C and 2, A and B; Axelrod, 2001; Strutt, 2001; Tree et al., 2002; Bastock et al., 2003; Das et al., 2004). By generating mutant clones and examining PCP localization at the clone border, the intercellular interactions between neighboring cells can be assessed in vivo. For example, when Fz is lacking within a clone, leaving only Vang–Fmi available at cell junctions, then Fz–Fmi in adjacent wild-type cells is recruited to the clone border (Chen et al., 2008). Vang mutant clones produce a similar effect, but in this case the excess Fz recruits Vang to clone borders (Bastock et al., 2003). What mediates these intercellular asymmetric interactions? One possibility is that Vang and Fz interact directly, and in vitro binding assays between the Fz extracellular domain and Vang suggest that this mechanism is possible (Wu and Mlodzik, 2008). However, mutants of Fz or Vang lacking their extracellular domains can still recruit one another between cells, which suggests that something else must bridge the two proteins (Chen et al., 2008). The seven-pass transmembrane cadherin, Fmi, likely performs this function. Fmi is essential for the junctional recruitment of Fz and Vang, and Fmi homodimers appear to be functionally asymmetric (Chen et al., 2008; Strutt and Strutt, 2008; Struhl et al., 2012). Clonal overexpression of Fmi preferentially recruits Fz to the clone border, even in the absence of Vang, which suggests that excess or unpaired Fmi is in a configuration that has higher affinity for Fmi–Fz than Fmi–Vang (Chen et al., 2008; Strutt and Strutt, 2008). Thus, Fmi may exist in two forms depending on whether it is paired with Fz or Vang, but the molecular basis for this difference is not known (Chen et al., 2008; Strutt and Strutt, 2008; Struhl et al., 2012).

Amplification of asymmetry.

Intercellular Fz–Fmi and Vang–Fmi complexes can form between cells in any orientation, so how do they resolve into discrete and opposed asymmetric domains? One way is through clustering of Fz–Fmi and Vang–Fmi complexes of the same orientation, and the cytoplasmic PCP components are particularly important for this function. As PCP complexes grow increasingly asymmetric, they cluster into discrete puncta that are stably associated with the plasma membrane and are resistant to endocytosis (Strutt et al., 2011). FRAP analysis of Fz-containing puncta demonstrated that they are highly stable compared with diffuse Fz-GFP, and have limited lateral mobility within the membrane. In the absence of Dsh, Pk, or Dgo, the size, intensity, and stability of Fz-containing puncta are diminished (Strutt et al., 2011), whereas overexpression causes Fz accumulation and coalescence into larger puncta (Feiguin et al., 2001; Tree et al., 2002; Bastock et al., 2003). Although the precise mechanisms driving PCP puncta formation are not known, the cytoplasmic components do not affect endocytosis, which suggests that they contribute to puncta formation by clustering intercellular complexes (Strutt et al., 2011). Pk can interact homophilically (Jenny et al., 2003; Ayukawa et al., 2014), which might promote clustering of proximal Vang–Pk–Fmi complexes. It will also be interesting to determine whether the cytoskeleton is directly connected to PCP complexes to minimize lateral mobility within the membrane.A second mechanism contributing to PCP asymmetry is directed transport. Live imaging of fluorescently tagged PCP proteins in pupal wings showed that Fz- and Dsh-containing particles travel across the cell in a proximal-to-distal direction (Shimada et al., 2006; Matis et al., 2014; Olofsson et al., 2014). These particles most likely represent endosomes undergoing transcytosis, as they arise from the proximal cortex and are labeled by the endocytic tracer FM4-64. This mechanism could serve to amplify asymmetry or even provide the initial polarity bias by removing proximal Fz–Dsh–Fmi complexes and transporting them to the distal side. Directed PCP transport is mediated by an array of subapical, noncentrosomal microtubules (MTs) that align along the proximal–distal axis, with the plus ends oriented with a slight distal bias (Hannus et al., 2002; Shimada et al., 2006; Harumoto et al., 2010; Matis et al., 2014; Olofsson et al., 2014). Ft and Ds are required for proximal–distal MT alignment (Harumoto et al., 2010), which suggests that the Ft-Ds system may feed into the core PCP system by orienting cytoskeletal architecture to deliver Fz–Dsh–Fmi complexes to the distal edge of the cell.Directed transport of Vang-containing endosomes has not been reported in flies, but selective trafficking could target Vang to specific membrane domains. In mammalian cells, exit of the Vang homologue Vangl2 from the trans-Golgi network (TGN) requires Arfrp1 (an Arf-like GTPase) and the clathrin adaptor complex AP-1, neither of which are required for the transport of a mammalian Fz homologue Fz6 or Fmi/Celsr1, which suggests that the differential sorting of PCP complexes to opposite sides of the cell could initiate at TGN export (Guo et al., 2013). Whether newly synthesized Vang and Fz proteins are transported to opposing cell surfaces from the TGN has not yet been explored.Microtubule orientation also correlates with PCP asymmetry in mouse trachea epithelial cells, where PCP coordinates the alignment of motile cilia (Vladar et al., 2012). MTs are planar polarized with their plus ends oriented toward the Fz–Dvl domain, and disruption of MTs with nocodazole impairs core PCP localization. Similarly, MTs are needed to establish Pk asymmetry in gastrulating zebrafish embryos (Sepich et al., 2011). However, in the skin epithelium, MTs align perpendicular to the axis of PCP asymmetry (unpublished data). Thus, directed transport along MTs may not be required in all tissue types for the establishment of PCP asymmetry.

Negative regulation.

Repulsive interactions between Vang- and Fz-containing complexes may also contribute to the amplification of asymmetry, and cytoplasmic proteins have been proposed to perform this function. Pk and Dgo both bind to Dsh in vitro, interacting with the same domain on Dsh in a mutually exclusive manner (Jenny et al., 2005). In addition, overexpression of Pk can prevent Dsh translocation to the membrane (Tree et al., 2002; Carreira-Barbosa et al., 2003), which suggests that Pk binding to Dsh could displace it from the proximal side of the cell. On the distal side, Dgo binding to Dsh would prevent association with Pk, thus enhancing Dsh distal localization. This increase in Dsh and Pk asymmetry would then positively feed back by clustering the transmembrane components into stable membrane domains.Modulation of PCP protein levels by ubiquitin-mediated degradation also leads to feedback by restricting the amount of one PCP protein to antagonize another. In flies, regulation of Dsh by a Cullin-3-BTB E3 ubiquitin ligase complex limits its levels at cell junctions (Strutt et al., 2013a). Reduction of Cullin-3 leads to an increase in overall core PCP protein levels, a reduction of asymmetry, and defects in wing hair polarity, which is consistent with Dsh overexpression phenotypes (Strutt et al., 2013a). SkpA, a subunit of the SCF E3 ligase, regulates Pk levels by promoting its degradation in a Vang-dependent manner (Strutt et al., 2013b). In mice, Smurf E3 ligases ubiquitinate Pk and promote its local degradation by binding to phosphorylated Dvl2 (a mammalian homologue of Dsh; Narimatsu et al., 2009). Smurfs are required for Pk localization in the inner ear and floor plate, and their removal leads to defects in convergent extension (CE) and stereocilia alignment (Narimatsu et al., 2009). Thus, targeting Pk for degradation either balances total Pk protein levels or targets a specific pool of Pk for ubiquitination and proteasome degradation.

Tissue-level polarity cues: This way or that?

What provides the tissue-level polarity cue that biases core PCP asymmetry in one direction over another? This is perhaps the most fundamental, yet poorly understood, element of PCP. Current models propose that an upstream, graded cue provides an initial bias in PCP asymmetry by regulating the levels, localization, or activity of one or more of the core proteins. Gradients are attractive candidates for providing global polarity cues, as they can act across many cells and define the tissue boundaries over which polarity must be oriented.

Ft–Ds–Fj.

Unlike the core proteins, Ds and Fj are nonuniformly expressed in the Drosophila eye, wing, and abdominal segments, and as such, the Ft–Ds module has been proposed to provide a global polarity cue (Fig. 2, C and D; for review see Ma et al., 2003; Yang et al., 2002; Thomas and Strutt, 2012; Matis and Axelrod, 2013). Ft and Ds are heterodimeric cadherins, regulated by the Golgi kinase Fj (Ishikawa et al., 2008; Brittle et al., 2010; Simon et al., 2010). The complementary expression patterns of Ds and Fj are thought to give rise to asymmetric Ft and Ds protein localization, with Ft and Ds localizing to opposite sides of each cell (Fig. 2 D; Ambegaonkar et al., 2012; Bosveld et al., 2012; Brittle et al., 2012). Because Fj positively regulates the activity of Ft, a gradient of Ft activity is expressed across the wing complementary to that of its ligand, Ds (Simon et al., 2010). Mutations in the Ft–Ds system give rise to swirling wing hair patterns, and disrupt the global alignment of core PCP proteins, but not their asymmetric distributions.An appealing model for symmetry breaking in the early Drosophila wing is that cellular asymmetries of Ft–Ds polarize MT organization and promote the distal transport of Fz–Dsh–Fmi vesicles (Shimada et al., 2001, Harumoto et al., 2010; Matis and Axelrod, 2013; Matis et al., 2014). This would produce an initial bias in Fz–Dsh localization, which would then be amplified by feedback interactions. However, several pieces of evidence have prevented the model from gaining universal acceptance. First, Ds and Fj gradients are oriented in opposite directions with respect to the core PCP proteins in the wing compared with the eye and abdomen. This discrepancy has been rectified with the finding by two independent groups that cells interpret Ft–Ds–Fj gradients differently depending on which of two Pk isoforms is expressed (Ayukawa et al., 2014; Olofsson et al., 2014). Second, the Ft–Ds system can orient PCP independently of the core pathway, and thus the two systems orient polarity in parallel, as opposed to in a single, common pathway (Casal et al., 2006). Third, Ft–Ds mutations affect core PCP orientation only regionally in the wing, which suggests that, if Ft–Ds provides a global bias, other, redundant cues must also exist (Matakatsu and Blair, 2006; Matis et. al., 2014). Finally, the direction of Ft–Ds and core PCP asymmetry diverges late in wing development, where the two systems become completely uncoupled. Intriguingly, the extent of coupling depends on which isoform of Pk is expressed (Merkel et al., 2014). Perhaps the simplest explanation for Ft–Ds function is that it can both transmit polarity information independent of the core system and organize the cytoskeleton to provide an initial bias of core PCP asymmetry, but which mechanism predominates depends on the tissue and developmental stage.

Wnts.

Wnt proteins have long been considered attractive candidates to provide tissue-level polarity cues because Fz and Dsh are primary components of the Wnt–β-catenin signaling pathway. Wnts are secreted glycoproteins that bind to Fz and other receptors, and often display graded expression. In vertebrates, Wnts are clearly important regulators of PCP, but whether they act instructively or permissively is unclear. In zebrafish, Wnt5a and Wnt11 are required for CE movements during gastrulation, but uniform expression of Wnt11 rescues the mutant phenotype, which suggests that it is permissive rather than instructive (Heisenberg et al., 2000; Kilian et al., 2003). Wnt5a is expressed in a gradient along the axis of polarity in the mouse inner ear, where it interacts genetically with Vangl2 in cochlear hair cell orientation (Qian et al., 2007). In the mouse limb, Wnt5a and its atypical receptor Ror2 are required for limb elongation and the asymmetric localization of Vangl2 at the proximal face of converging and extending chondrocytes (Gao et al., 2011). Wnt5a is expressed in a distal-to-proximal gradient, which induces a gradient of Vangl2 phosphorylation. The functional consequences of Vangl2 phosphorylation are unknown but Vangl2 cellular asymmetry appears to be strongest distally, where Wnt5a and Vangl2 phosphorylation levels are highest (Gao et al., 2011).While several studies had argued against the involvement of Wnt proteins in Drosophila PCP (Lawrence et al., 2002; Chen et al., 2008), it was recently discovered that Wingless (Wg) and Wnt4a act redundantly to orient PCP in the wing, particularly near the wing margin (Wu et al., 2013). Misexpression of Wg or Wnt4a reorients wing hair polarity in a pattern reminiscent of Fz loss of function, which suggests that Wnt gradients may orient polarity by antagonizing Fz. Consistently, the ability of Fz and Vang to recruit one another between adjacent cells in culture was inhibited by the addition of Wg or Wnt4a, which suggests that Wnts could provide a polarizing cue by diminishing Fz–Vang interactions at the margin of the wing, where Wnt expression is highest (Wu et al., 2013). However, Wnt4a overexpression also reorients MT alignment, suggesting that Wnts may act as polarity cues thorough an effect on the cytoskeleton (Matis et. al., 2014). Alternatively, Sagner et al. (2012) suggest that Wg orients core PCP indirectly through its effects on wing patterning and growth. Although the evidence for Wnt gradients as global PCP cues is accumulating, the mechanisms by which they regulate core protein levels or activity remain to be elucidated.

Mechanical forces.

Anisotropic mechanical forces that accompany growth and morphogenesis can also provide global polarizing cues. During wing development, PCP reorients in response to extensive morphogenetic changes that elongate the wing along the proximal–distal axis. In early pupal wings, PCP aligns toward the wing margin and then reorients during wing elongation and contraction of the wing hinge (Aigouy et al., 2010). These morphogenetic changes have broad effects on cell behavior, inducing cell elongation, oriented divisions, and cell rearrangements with a concomitant reorientation of PCP. Severing the wing pouch from the hinge blocks cell flows and PCP reorientation, which suggests that the anisotropic tension from hinge contraction drives tissue flow and the reorientation of polarity (Aigouy et al., 2010). Although this model doesn’t explain what initially biases PCP, it does demonstrate how the morphogenetic processes that shape tissues can completely remodel global PCP alignment. This is an attractive model to explain how PCP aligns over very large tissues, like the mammalian skin, where hairs consistently reorient along regions of extensive tissue elongation such as the face, limbs, and ears.

Downstream effectors of PCP: Steering the wheel

If PCP is the cell’s compass, it is also the steering wheel, directing downstream, polarized cell behaviors in response to global directional cues. PCP can polarize a wide range of cell behaviors, which suggests that it can intersect with numerous downstream effectors. We focus here on three examples where the molecular mechanisms linking core PCP to their polarized outputs have recently been elucidated.

Distal positioning of wing hairs.

Each cell of the Drosophila wing blade emits a single actin-rich protrusion from its distal edge. The placement of the wing hair strongly correlates with the position of Fz–Dsh–Fmi, which suggests that core proteins may localize cytoskeletal regulators to distinct positions within the cell (Strutt and Warrington, 2008). On the proximal side, Vang recruits a group of proteins that negatively regulate actin prehair formation: Inturned, Fuzzy, and Fritz (Adler et al., 2004; Strutt and Warrington, 2008). These three proteins regulate Multiple Wing Hairs, a GTPase-binding/formin-homology 3 (GBD/FH3) domain protein thought to repress actin polymerization (Strutt and Warrington, 2008; Yan et al., 2008). This restricts actin nucleation to distal positions within the cell, and in the absence of Multiple Wing Hairs, ectopic actin bundles form across the apical surface (Wong and Adler, 1993). On the distal side, casein kinase 1 γ CK1g/gilamesh is required to further refine prehair nucleation to a single site through a parallel mechanism involving Rab11-dependent vesicle traffic to the site of prehair formation (Gault et al., 2012). Rho and Rho kinase (Drok) have also been implicated in wing hair formation, but their roles are difficult to dissect due to the numerous functions of Rho in cell shape and cell division (Winter et al., 2001; Yan et al., 2009).

Actomyosin contraction and convergent extension (CE).

CE was the first vertebrate process to be linked molecularly to PCP (Wallingford et al., 2000). During CE, mesenchymal cells elongate, form mediolateral-directed protrusions, and intercalate mediolaterally, narrowing the mediolateral axis while simultaneously lengthening the anterior–posterior (A-P) axis (Fig. 3 A; Keller, 2002). Mediolateral polarization, elongation, and intercalation are lost when core PCP components are disrupted, leading to a failure in CE (Tada and Smith, 2000; Wallingford et al., 2000; Goto and Keller, 2002; Jessen et al., 2002). While several PCP-dependent mechanisms have been proposed to mediate CE movements, two recent studies provide direct mechanistic links between asymmetrically localized core PCP components and CE behaviors. In neuroepithelial cells, PCP specifies the localization of myosin to the A-P faces of intercalating cells. Fmi/Celsr1 and Dvl recruit the formin DAAM1 to the A-P junction, which in turn binds and activates PDZ-RhoGEF. This likely activates RhoA and myosin contractility specifically at A-P junctions, resulting in medial-directed cell intercalation and neural plate bending (Nishimura et al., 2012). A similar mechanism was found to drive CE movements of mesenchymal cells during Xenopus gastrulation. In this case, Fritz and Dsh help to localize septins to mediolateral vertices, where they spatially restrict cortical actomyosin contractility and junctional shrinking to A-P cell edges, thus driving cell intercalation (Fig. 3 A; Kim et al., 2010; Shindo and Wallingford, 2014). Together these studies show how asymmetric PCP localization produces collectively polarized cell behaviors through spatial modulation of the cytoskeleton.Open in a separate windowFigure 3.Polarized cell behaviors controlled by PCP. (A) PCP drives convergent extension (CE). CE in vertebrates is driven by mediolateral intercalation, which narrows the mediolateral axis while simultaneously lengthening the A-P axis. Mediolateral intercalation is accompanied by cell polarization and elongation and the formation of mediolateral protrusions, all of which require core PCP function. Pk localizes anteriorly (Ciruna et. al., 2006; Yin et al., 2008), whereas Dsh localizes posteriorly (Yin et. al., 2008). In addition, PCP proteins recruit myosin to A-P cell borders, leading to actomyosin contractility and junctional shrinking. (B) Asymmetric cell division. Drosophila sensory organ precursors (SOPs) divide asymmetrically along the epithelial plane, giving rise to distinct anterior and posterior daughters. Spindle alignment along the A-P axis is PCP dependent. Dsh interacts with Mud/NuMA and the dynein complex posteriorly while Vang links Pins/LGN-Mud/NuMA-dynein on the anterior. This links astral MTs to the A-P cortex, bringing the spindle into register with the A-P axis. (C) Positioning of the kinocilium in the inner ear. The placement of kinocilium in sensory hair cells of the inner ear determines the position of V-shaped stereocilia bundles. Gαi and mPins/LGN localize on the abneural side on the hair cell, where they are required for abneural positioning the MT-based kinocilium. The collective alignment of kinocilia and stereocilia bundles across the epithelium requires the core PCP component Vangl2. Vangl2 (light green) localizes to the abneural side of supporting cells. Whether Fz (dark blue) associates on the opposite face is not yet clear (Ezan et. al., 2013).

Positioning of centrosomes and cilia.

PCP regulates the positioning of MT-based structures including the mitotic spindle and cilia. In Drosophila sensory organ precursors and early zebrafish embryos, PCP controls mitotic spindle orientation along the epithelial plane by interacting with the highly conserved spindle orientation complex, which links astral MTs to the cell cortex through Mud/NuMA-mediated recruitment of the dynein complex (Ségalen et al., 2010). To orient the spindle, posteriorly localized Dsh binds to Mud/NuMA, which recruits the dynein complex and astral MTs to the posterior cortex. On the anterior side, Pins/LGN recruits Mud/NuMA, bringing the spindle into A-P alignment (Fig. 3 B). Similarly, PCP was recently shown to interact with the spindle orientation machinery to position the kinocilium in nondividing cells of the inner ear (Ezan et al., 2013; Tarchini et al., 2013). In vestibular hair cells, Gαi and mPins/LGN localize to the abneural cortex, opposite Vangl2, where they are required for kinocilium positioning and subsequent alignment of stereocilia bundles (Fig. 3 C; Ezan et al., 2013). MT plus ends and dynein also show an abneural bias suggesting that Gαi-mPins/LGN induces pulling on MTs by a similar mechanism that orients the centrosome during spindle orientation. Vangl2 is required for the alignment of Gαi-Pins/LGN crescents between cells, coordinating kinocilia positioning and stereocilia polarity across the tissue (Fig. 3 C; Ezan et al., 2013). Thus the PCP pathway co-opts the spindle orientation machinery to specify not only the division plane but also cilia position in nondividing cells. As PCP is required for asymmetric cilia positioning in a wide range of cell types, including the node (Antic et al., 2010; Borovina et. al., 2010; Song et al., 2010; Hashimoto et. al., 2010), it will be interesting to determine whether this mechanism is conserved.

Concluding remarks

PCP is a fundamental and highly conserved process coordinating a vast number of polarized cell behaviors. While the number of functions ascribed to PCP continues to grow, an understanding of the mechanisms establishing PCP is still far from complete. The development of cellular asymmetry from uniform distributions is not well understood, and will benefit from recent advances in high-resolution, time-lapse imaging with photoconvertible fluorescent tags. Other important issues to resolve include deciphering the structural domains and biochemical interactions mediating intercellular communication, identifying the global cues that orient PCP especially in vertebrates, and deciphering the mechanisms by which complex multicellular structures, like lung branches and hair follicles, are oriented by the PCP machinery.