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. Author manuscript; available in PMC: 2015 Aug 10.
Published in final edited form as: Adv Exp Med Biol. 2013;782:23–38. doi: 10.1007/978-1-4614-5465-6_2

The molecular basis of experience-dependent motor system development

Robert G Kalb 1, Lei Zhang 2, Weiguo Zhou 2
PMCID: PMC4530779  NIHMSID: NIHMS712936  PMID: 23296479

Abstract

Neurons in the vertebrate nervous system acquire their mature features over an extended period in pre-natal and early post-natal life. The interaction of the organism with its environment (“experience”) has been shown to profoundly influence sensory neuron development. Over the past ~2 decades, it has become increasingly clear that motor system development is also experience-dependent. Glutamate receptors of the N-methyl-D-aspartate (NMDA) subtype have been implicated in both sensory and motor system experience-dependent development. An additional molecular mechanism involves the GluA1 subunit of the 2-amino-3-(5-methyl-3-oxo-1,2-oxazol-4-yl)propanoic acid (AMPA) subtype glutamate receptors. GluA1-dependent development operates in an NMDA-R independent manner and uses a distinct set of signaling molecules. The synapse associated protein of 97 kDa molecular weight (SAP97) is key. A deeper understanding of how experiences guides motor system development may lead to new ways to improve function after central nervous system insult.

Overview

Nervous system operation depends on the ability of neurons to process a panoply of excitatory and inhibitory signals and generate meaningful output. The size and complexity of the dendritic tree is a critical determinant of the computational work of neurons (Stuart, 1999). In addition, the geometry of the dendritic tree can regulate who communicates with a neuron by controlling the quantitative and qualitative nature of the afferent input (Hume & Purves, 1981). Theorists, taking a wiring optimization approach, view synaptic connectivity and neuronal morphology as inextricably linked because it is the most efficient fit of network wiring within a given volume of neuropil (Chklovskii, 2004). The spectrum of animal behavior, from 0.5 mm round worm wiggling on a petri dish to Glen Gould playing Bach, reflects the precision with which neurons elaborate their dendritic tree and are innervated.

Cell and molecular biology of activity-dependent development

The process of dendrite elaboration is often divided into an initial, synaptic activity-independent phase and a subsequent, synaptic activity-dependent phase (Goodman & Shatz, 1993; Shatz, 1990). The first phase sets up the basic architecture of the tree and is likely to be under strong genetic control (Gao & Bogert, 2003; Jan & Jan, 2003). The synaptic activity-dependent phase of dendrite elaboration is believed to fine tune structure into a precise configuration (H. T. Cline, 2001; Constantine-Paton, Cline, & Debski, 1990). There is much evidence to support the “synaptotrophic hypothesis” as a mechanism for activity-dependent regulation of dendrite arbor development ((J.E. Vaughn, 1989; J. E. Vaughn, Barber, & Sims, 1988), reviewed by (H. Cline & Haas, 2008)). Developing axons and dendrites undergo exploratory growth and make nascent synapses. Adhesion molecules such as neurexins (NRX) and neuroligins (NLG) (Chen, Tari, She, & Haas, 2010) (Thyagarajan & Ting, 2010) are likely to be involved. Repeated use of a synapse leads to stabilization of the synapse if pre-and post-synaptic elements are coincidently active (Bi & Poo, 2001; Engert, Tao, Zhang, & Poo, 2002; Ruthazer, Akerman, & Cline, 2003). Axons and dendrites that bear stable synapses are retained (or perhaps grow) and conversely, portions of axons and dendrites that do not bear stable synapses are withdrawn (Katz & Constantine-Paton, 1988). Spontaneous synaptic activity is the initial driver of these events and subsequently environmentally-evoked synaptic activity does the heavy lifting. Experience-dependent refinement of neuronal architecture and synaptic connectivity sculpts each nervous system to perform best in the environment in which the animal was reared (L. I. Zhang, Tao, & Poo, 2000).

One form of activity-dependent development involves activation of the NMDA subtype of glutamate receptor (H.T. Cline, Debski, & Constantine-Paton, 1987; R.G. Kalb, 1994; Kleinschmidt, Bear, & Singer, 1987). This leads to a substantial rise in dendritic calcium, which is thought to be the key trigger for subsequent events. The precise ordering of what occurs next is not entirely clear, but the literature supports the view is that there are three linked main events. Event #1 is the secretion/elaboration of extracellular factors such as BDNF, Wnts and nitric oxide (Cramer, Angelucci, Hahm, Bogdanov, & Sur, 1996; Inglis, Furia, Zuckerman, Strittmatter, & Kalb, 1998; H. H. Wu, Williams, & McLoon, 1994). There is evidence that secretion/elaboration is activity-dependent and blocking their action can prevent synapse stablization and dendrite growth. ((A. K. McAllister, Katz, & Lo, 1996; A.K. McAllister, Katz, & Lo, 1997)but see (Lu, 2003)). Event #2 is the activation of intracellular signaling molecules such as Ca++/calmodulin-dependent kinase (CamK) type I (Wayman et al., 2006), CamK II (G.-Y. Wu & Cline, 1998; Zou & Cline, 1999) (Gaudilliere, Konishi, de la Iglesia, Yao, & Bonni, 2004) and CamK IV (Redmond, Kashani, & Ghosh, 2002), MAP kinase(Ha & Redmond, 2008; Redmond et al., 2002; G. Y. Wu, Deisseroth, & Tsien, 2001), βcatenin(Yu & Malenka, 2003) (Peng et al., 2009)and RhoA GTP’ases (Z. Li, Aizenman, & Cline, 2002; Z. Li, Van Aelst, & Cline, 2000; Sin, Haas, Ruthazer, & Cline, 2002). The role of these molecules has been studied using pharmacological inhibitors and expression of dominant-negative and constitutively-active forms of these proteins. Event #3 is new gene expression and the list of contributing transcription factors includes CREB (S. Li, Zhang, Takemori, Zhou, & Xiong, 2009; Redmond et al., 2002; Wayman et al., 2006), CREST (Aizawa et al., 2004), NeuroD (Gaudilliere et al., 2004) and MEF2A (Shalizi et al., 2006). As above, the role of these molecules has been studied employing molecular genetic techniques. Modification of cytoskeletal elements, maturation of silent synapses (NMDA-R only → AMPA-R + NMDA-R), and the precise apposition of pre- and postsynaptic membranes incorporating adhesion molecules are all necessary steps in this process. With so many events apparently occurring simultaneously it is difficult to discern the epistatic relationships. How activity-dependent processes dovetail with dendrite growth promoting processes not-shown-to-be-activity-dependent (such as activation of PI3’K and mTOR (Jaworski, Spangler, Seeburg, Hoogenraad, & Sheng, 2005; Kumar, Zhang, Swank, Kunz, & Wu, 2005) and Notch signaling (Redmond, Oh, Hicks, Weinmaster, & Ghosh, 2000)) only complicates matters more.

Experience-dependent motor system development

The normal development of the locomotor system (from behavior, to connectivity within the segmental spinal cord, to motor neuron dendrite architecture) emerges during pre-natal and early postnatal life (Altman & Sudarshan, 1975; Curfs, Gribnau, & Dederen, 1994; Curfs, Gribnau, & Dideren, 1993; Donatelle, 1977; Pellis, Pellis, & Teitelbaum, 1991; Seebach & Ziskind-Conhaim, 1994; Snider, Zhang, Yusoof, Gorukanti, & Tsering, 1992). Below, I outline the evidence that locomotor development is experience-dependent and that the molecular machinery that drives this process can involve activation of NMDA-Rs. In addition, I will provide evidence for a second set of molecules that appear to act in parallel with NMDA-Rs to drive motor system development. We have found that GluA1 subunit of the AMPA-R, in concert with an intracellular binding partner called SAP97, promote motor system development by an NMDA-R-independent mechanism.

Experience-dependent motor system development and the NMDA-R

We begin with studies by Kerry Walton’s group of neonatal rats reared in space. The force of gravity at the surface of the earth is called “1G” and anything less than that is referred to as “microgravity”. Walton’s group studied a cohort of animals that spent about 2 weeks of early postnatal life in the Space Shuttle. She showed that young rats that develop in microgravity have demonstrably different locomotor behavior than those that develop in earth (K. D. Walton, Harding, Anschel, Harris, & Llinas, 2005). These observations echo her previous work using the tail suspension model (K.D. Walton, Lieberman, Llinas, Begin, & Llinas, 1992). Work from my lab using these mice demonstrated that the motor neuron dendritic tree also undergoes experience-dependent development (Inglis, Zuckerman, & Kalb, 2000). The parsimonious construct is that at least some on the alterations in motor neuron dendrite structure subserve the alterations in locomotor function that follow microgravity rearing.

These behavioral and anatomical studies prompted us to ask whether NMDA-Rs were involved in activity-dependent maturation of motor neuron dendritic architecture. We began by asking whether NMDA-R components were expressed by developing motor neurons. In situ hybridization studies show that newborn motor neurons express NR1, NR2A and NR2C at particularly high levels and over the subsequent next few weeks of life, the abundance of these mRNAs falls off considerably (Stegenga & Kalb, 2001). The NR1 subunit undergoes alternative splicing, and an analysis of specific NR1 variants reveals that NR1A, NR1B, NR1-2 and NR1-4 are expressed at particularly high levels in newborn motor neurons. The abundance of these splice variants falls subsequently in early postnatal life (Stegenga & Kalb, 2001). This work demonstrates that motor neurons express a unique repertoire of NMDA-R subunits in early postnatal life.

Coincident with the period when motor neurons express a distinct type of NMDA-R, the dendrites of motor neurons are undergoing substantial growth (Curfs et al., 1993; Lindsay, Greer, & Feldman, 1991; Núñez-Abades, He, Barrionuevo, & Cameron, 1994). Overall tree size and number of branches increase ~2 to 3-fold between postnatal day 7 (P7) and P21. Antagonism of NMDA-Rs with APV or MK-801 inhibits the growth of motor neuron dendrites of NMDA-R (R.G. Kalb, 1994). In contrast to their effects on developing dendrites, antagonism of NMDA-Rs in adult animals has no effect on motor neuron dendrite architecture. These results indicate that during a critical period in early postnatal life, activation of NMDA-Rs promotes the elaboration of motor neurons dendrites. In subsequent work we showed that the dendrite growth promoting actions of NMDA-Rs are mediated by the second messenger, nitric oxide (Inglis et al., 1998). Overall this work highlights the distinct parallel between the experience-dependent development of sensory and motor systems.

A novel form of activity-dependent development utilizes GluA1 containing AMPA-R

In addition to characterizing the expression pattern of NMDA-R subunits, we also examined the expression pattern of AMPA-R subunits. Although all subunits undergo developmentally regulated expression, we were impressed that neonatal motor neurons express particularly high levels of the GluA1 (mRNA and protein) (Jakowec, Fox, Martin, & Kalb, 1995; Jakowec, Yen, & Kalb, 1995). (This is the unedited version of the protein that contains the “flip” alternatively spliced exon and unless otherwise stated, I will use “GluA1” to denote “GluA1(Q)flip”). Electrophysiologic studies show that neonatal motor neurons display Ca++-permeable AMPA receptors (as would be expected from assembled tetramers enriched with GluA1 or even homomeric GluA1 tetramers) (Carriedo, Yin, & Weiss, 1996; Vandenberghe, Robberecht, & Brorson, 2000). This raises the possibility that the special type of AMPA receptors expressed by neonatal motor neurons is part of the molecular mechanism of experience-dependent dendrite development. The first good clue that this could be the case was a study in which we overexpressed (OE) GluA1 in mature motor neurons (after the period of developmental dendrite growth) (Inglis et al., 2002). We found that this led to large-scale remodeling of the dendritic tree with a marked increase in dendrite branching. Expression of a version of GluR1 with an arginine in the critical “Q/R editing site” (GluA1(R)) had no effects on dendrite architecture. AMPA-R assembled from GluA1(R) are calcium impermeable and pass very little current upon activation with glutamate. These in vivo observations suggest that GluA1 can promote dendrite growth and this depends on the ability of GluA1 containing AMPA-R to depolarize cells. Subsequent in vivo and in vitro work provide strong support for the idea that calcium permeability of GluA1 containing AMPA-R is a major determinant of its effect on dendrite growth (Jeong et al., 2006).

One could imagine at least two ways in which increasing the expression of AMPA-R assembled with GluA1 might promote dendrite growth: 1) By enhancing the ability of cells to depolarize upon afferent stimulation, GluA1 might facilitate the activation of NMDA-Rs. In this scenario, GluA1 acts upstream of NMDA-R mediated events; 2) GluA1 may act through an NMDA-R-independent pathway to regulated dendrite growth. In this scenario, GluA1 acts in parallel to NMDA-R mediated events. There are several reasons for favoring the second scenario. First, administration of MK-801 to rats OE GluA1 did not block the pro-dendrite growth effect. We know MK-801 was at an effective concentration in the brain because we could not evoke LTP in the dentate gyrus from animals treated with MK-801 (see figure 6 of (Inglis et al., 2002)). So OE GluA1 led to increased dendrite branching even though NMDA-Rs were effectively antagonized. Second, NMDA-R mRNAs are developmentally regulated and not expressed by mature motor neurons (Stegenga & Kalb, 2001). So GluA1-mediated dendrite remodeling can occur in neurons that do not express NMDA-R subunits. Finally we more formally examined the role of NMDA-Rs in GluA1-dependent dendrite growth in vitro.

We grew dissociated spinal cord neurons in vitro and expressed GluA1 by transfection and treated some cells with the NMDA-R open-channel blocker MK-801 (10 μM). Three groups of neurons were studied morphologically: 1) GFP, 2) GFP + GluA1 + vehicle and 3) GFP + GluA1 + MK-801. MK801 or vehicle was administered daily to the cultures and after 5 days in vitro (DIV), the cultures were fixed, immunostained for GFP to enhance the cell labeling, camera lucida drawings generated and quantitatively analyzed (Figure 1). Compared with GFP alone, GluA1 led to an ~ 30% increase in dendrite branches (F(2,60) = 3.507, p = 0.03), ~ 60% increase in overall arbor size (F(2,60) = 17.005, p < 0.0001), ~60% increase in average dendrite length(F(2,60) = 9.370, p = 0.0003) and a 40% increase in the length of the longest dendrite (F(2,60) = 10.817, p < 0.0001). Treatment with MK801 did not influence the pro-growth effects of GluA1; the dendritic arbor of neurons in the “GluA1 + MK-801” were not statistically different from the dendrite arbor of the neurons in the “GluA1” group. These observations establish that overexpression of GluA1 stimulates dendrite growth in a manner that is independent of NMDA-Rs.

Figure 1. Overexpression of WT GluA1 stimulates dendrite growth in vitro in an NMDA-R-independent manner.

Figure 1

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Top, representative camera lucida images of neurons expression GFP alone, or GFP + GluA1 (treated or not with MK801). The chart below provides quantitative analysis of dendrites as well as statistical analysis using ANOVA. The number of neurons drawn is noted in parentheses next to the column title. There is statistically significant increase in branching and overall tree size in neurons overexpressing GluA1 and these effects are not influenced by NMDA-R antagonism.

In addition to NMDA-R, voltage-gated calcium channels are activated by membrane depolarization, and so these channels might participate in the GluA1 form of dendrite growth. To examine this issue we undertook a second set of experiments we used the L-type calcium channel blocker nifedipine (20 μM). Three groups of neurons were studied morphologically: 1) GFP, 2) GFP + GluA1 + vehicle and 3) GFP + GluA1 + nifedipine. Nifedipine or vehicle was administered daily and cells were prepared for analysis as above (Figure 2). Compared with GFP alone, GluA1 led to an ~ 40% increase in dendrite branches (F(2,63) = 8.396, p = 0.0006), ~ 40% increase in overall arbor size (F(2,63) = 12.99, p < 0.0001), and ~20% increase in average dendrite length(F(2,63) = 9.085, p = 0.0004). Treatment with nifedipine blocked all the pro-dendrite growth effects of GluA1 overexpression on branching and even suppressed overall tree growth and elaboration of the longest dendrite in comparison with neurons treated with vehicle. It is interesting that recent work has shown that L-type calcium channels play a critical role in homeostatic synaptic plasticity (Goold & Nicoll, 2011). Thus the activity-dependent dendrite growth elicited by GluA1 is NMDA-R-independent and requires activation of voltage-gated calcium channels.

Figure 2. Overexpression of WT GluA1 stimulates dendrite growth in vitro in a voltage gated calcium channel-dependent manner.

Figure 2

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Top, representative camera lucida images of neurons expression GFP alone, or GFP + GluA1 (treated or not with nifedipine). The chart below provides quantitative analysis of dendrites as well as statistical analysis using ANOVA. The number of neurons drawn is noted in parentheses next to the column title. There is statistically significant increase in branching and overall tree size in neurons overexpressing GluA1 and these effects are blocked by voltage gated calcium channel antagonism.

The above described work argues that expression of GluA1 is sufficient to promote dendrite growth. Simply overexpressing the protein can trigger the elaboration of dendrites (either in vitro or in vivo). To determine if GluA1 is necessary for dendrite growth under endogenous conditions requires that we reduce or eliminate its expression in neurons and determine the effects on morphogenesis. In vitro studies using RNAi technology reveal that reducing GluA1 in neurons inhibits the normal elaboration of dendrites (L. Zhang et al., 2008). The effects are dose-dependent and cell autonomous. These observations support the view that expression of GluA1 by neurons is necessary for the normal morphogenesis of the dendritic tree.

The in vivo role of GluA1 in motor system development

Activity-dependent growth of dendrites is a component of experience-dependent motor system development. In light of this we wondered if the above in vitro observations were also seen in vivo. If so, we would be well positioned to determine the broader impact of GluA1 on other features of experience-dependent motor development such as behavior and circuit connectivity.

GluA1 null mice (GluA1−/−) are viable and with the exception of selective defects in cognitive function are believed to be normal (Zamanillo et al., 1999). We began our studies of these animals by asking if GluA1−/− neonatal motor neurons elaborate a normal dendritic tree. Both at P10 and P23, the dendritic tree of GluA1−/− motor neurons is smaller than WT control mice (L. Zhang et al., 2008). It is worth noting that in addition to motor neurons, interneurons within the neonatal ventral horn also express GluA1 at very high levels (Jakowec, Fox et al., 1995; Jakowec, Yen et al., 1995). And so it is possible that loss of GluA1 from interneurons that innervate motor neurons influences the capacity of motor neurons to elaborate a normal dendritic tree. To examine this issue directly we generated conditional knock-out mice. Mating Hb9-Cre mice with mice bearing a LoxP flanked allele of GluA1 generates mice with loss of one copy of GluA1. Using an appropriate breeding we generated mice that are homozygous for the LoxP flanked allele and Cre recombinase and these mice have ablation of GluA1 expression restricted to motor neurons. Analysis of dendrite from these animals (GluA1deltaHb9) revealed a reduction of dendrite size and branching, similar but not as severe as what we observed in the GluA1−/− mice (L. Zhang et al., 2008). There was no effect on dendrite structure in various control mice (i.e., Hb9-Cre alone or GluA1LoxP alone). These in vivo observations support the view that the expression of endogenous GluA1 by motor neurons is required for the normal elaboration of the dendritic tree during development.

Does the loss of GluA1 influence other aspect of the motor system, such as circuitry within the segmental spinal cord? To address the first question, we used the pseudorabies virus (PRV) tracing system. A recombinant PRV was generated that expressed GFP in cells. Upon injection into the hamstring leg muscle, PRV-GFP particles are retrogradely transported to motor neurons. PRV-GFP particles are then exported into dendrites where they cross synapses from motor neurons into innervating pre-motor interneurons. Thus, the distribution of GFP-labeled cells in the spinal cord reflect the pattern of pre-motor innervation of motor neurons. When we applied this approach to GluA1−/− and GluA1+/+ mice, we found that much of the GFP labeling was identical between groups. However, fewer labeled interneurons were seen in the ipsilateral Rexed lamina VIII of lumbar segment 4 of GluA1−/− than GluA1+/+ mice. In addition, there were fewer labeled interneurons in the multiple contralateral Rexed lamina of lumbar segment 2 – 5 of GluA1−/− than GluA1+/+ mice (L. Zhang et al., 2008). These results suggest that segmental spinal cord connectivity is different between the GluA1−/− and GluA1+/+ mice.

Do these alterations in dendrite structure and interneuronal connectivity manifest in behavioral differences between the GluA1−/− and GluA1+/+ mice? To examine this, we subject the two strains of mice to a battery of locomotor tasks including treadmill running, rotarod and fore- and hind-limb grip strength. At P23 and in adulthood, the GluA1−/− mice performed poorer in every single test in comparison with the GluA1+/+ mice (L. Zhang et al., 2008). Similar trends were seen when we studied the GluA1deltaHb9 although the degree of locomotor impairment was less than seen in the GluA1−/− mice. These differences in motor function could not be ascribed to a difference in motor neuron number. The weakness phenotype of the GluA1−/− mice was associated with an increase in type I muscle fibers in the gastrocnemius. Thus, elimination of GluA1 from motor neurons (as well as other neurons presumably in the ventral horn) leads to abnormal development of the neuromuscular unit. The dendritic tree of GluA1−/− motor neurons is stunted, the pattern of pre-motor interneuron connectivity is perturbed, muscle fiber type specification is distorted and this leads to poorer locomotor performance in comparison with WT animals. These observations point to the critical role that GluA1 plays in the normal activity-dependent development of the motor system.

SAP97 translates the GluA1-generated signal into dendrite growth

Our working hypothesis is that synaptic activation of AMPA-Rs assembled with the GluA1 subunit initiate an activity-dependent pro-dendrite growth signal. Some data suggest that the electrophysiological properties of GluA1-containing AMPA-R regulates how GluA1 influences dendrite morphology, and this is linked to AMPA-R calcium permeability (Jeong et al., 2006). We wondered if, in addition, intracellular proteins that bind GluA1 are also important determinants.

The extreme C-terminal four amino acids of GluA1 act as a ligand for the synapse-associated protein of 97 kDa molecular weight called SAP97 (Cai, Coleman, Niemi, & Keinanen, 2002). SAP97 is a MAGUK-class scaffolding protein and is the only known binding partner of the extreme C-terminus of GluA1. MAGUK proteins are enriched in the postsynaptic density where they play a variety of roles in synaptic function including chaperoning glutamate receptor subunits into and out of the synapse, receptor clustering and modulation of receptor electrophysiological function (Palmer, Cotton, & Henley, 2005; Sheng & Sala, 2001; Shepherd & Huganir, 2007). SAP97 is a modular protein with multiple protein-protein interaction domains. Detailed below, we have explored the dendrite growth promoting role of SAP97 and its binding to GluA1 in a series of in vitro and vivo experiments.

In co-immunoprecipitation (coIP) experiments using spinal cord or cerebral cortex tissue, we find that GluA1 and SAP97 are part of a physical complex (Zhou et al., 2008). When the two full length proteins are expressed in a heterologous system, we can again demonstrate a physical complex in the coIP assay. Two approaches were taken to establish the portions of each protein required for the physical complex. First we deleted the C-terminal 7 amino acids of GluA1 (GluA1Δ7). While full length SAP97 will coIP full length GluA1, it will not coIP GluA1Δ7. Second the crystal structure of PDZ domains is known and it is possible to introduce mutations such that the PDZ domain becomes incompetent to bind ligands (Morais Cabral et al., 1996). We engineered such mutations into PDZ2 of GluA1 (K323A, K326A) and found that mutant PDZ2 SAP97 did not co-IP full length GluA1 (Zhou et al., 2008). Thus GluA1 and SAP97 are part of a physical complex that is likely to be mediated by the binding of the extreme C-terminus of GluA1 to PDZ2 of SAP97.

What biology, if any, is influenced by the GluA1/SAP97 complex? We began by studying the trafficking of GluA1 through the secretory pathway to populate synapses. To address this issue we used a strain of mice in which the wild type version of GluA1 has been replaced with a version that is lacking the C-terminal 7 amino acids (Kim et al., 2005). In the homozygous state, this “knock-in” mouse only expresses GluA1Δ7. We find that GluA1Δ7 is synthesized in the endoplasmic reticulum at normal levels, is processed normally in the Golgi apparatus, hetero-oligomerizes normally with other AMPA-R subunits and inserts into synapses normally (Zhou et al., 2008). Electrophysiological studies of the hippocampus of GluA1Δ7 mice show normal basal synaptic transmission as well as normal LTP/LTP (Kim et al., 2005). Thus despite the fact that GluA1Δ7 does not physically associate with SAP97, the subunit behaves like wild type GluA1. In marked contrast, SAP97 does not traffic to synapses in the GluA1Δ7 mice. These observations indicate that GluA1 chaperones SAP97 into synapses.

What is SAP97 doing, in association with GluA1, at synapses? We took two approaches to address this issue. First we determined the effect on dendrite growth of eliminating SAP97 from neurons. When we knocked down SAP97 with an shRNA, the neuronal dendritic tree is smaller and less branched than WT neurons (Zhou et al., 2008). This implies that endogenous SAP97 is required for normal elaboration of the dendritic tree. We also found that knockdown of SAP97 blocked the dendrite growth promoting action of GluA1 overexpression. To validate these in vitro observations, we wanted to study mice in which SAP97 is ablated. Unfortunately SAP97 null mice die at birth owing to cranio-facial abnormalities. To overcome this problem we generated mice in which SAP97 is eliminated specifically in motor neurons. This was achieved using the Hb9-Cre mice mated to mice bearing a floxxed allele of SAP97. We found that the dendrites of motor neuron from the SAP97deltaHb9 mice are smaller and less branched than WT mice (Zhou et al., 2008). Thus studies both in vitro and in vivo demonstrate that SAP97 plays a key role in the normal development of the neuronal dendritic tree. In addition, all of the dendrite growth promoting actions of GluA1 are lost in the absence of SAP97. This suggests that SAP97 acts to translate activity from GluA1-containing AMPA-Rs into growth.

Our second approach to understanding what GluA1 and SAP97 are doing at synapses focused on the nature of their physical relationship. Must SAP97 be physically tethered to GluA1 to promote dendrite growth? Or, is co-localization of both proteins to the plasma membrane sufficient for GluA1 to promote dendrite growth? We undertook a series of experiments to explore this issue. First, we found that GluA1, but not GluA1Δ7, over expressed in neurons in vitro is dendrite growth promoting (Zhou et al., 2008). In addition, we found that co-expression of GluA1 with SAP97 has a synergistic dendrite growth promoting action, while co-expression of GluA1Δ7 with SAP97 leads to modest dendrite growth (equivalent to the dendrite growth promoting action of SAP97 itself). So even though GluA1Δ7 traffics to the cell surface and hetero-oligomerizes normally with other AMPA-R subunits, the lack of physical association with SAP97 blocks the dendrite growth promoting action of this subunit.

In the next set of experiments we added a palmitoylation sequence to SAP97 (palSAP97) and we show that this leads to membrane targeting of the protein. Both SAP97 and palSAP97 have equivalent dendrite growth promoting actions when overexpressed in neurons. Armed with this tool we took two approached to look at the necessity of a physical interaction between GluA1 and SAP97 for the promotion of dendrite growth. First, we asked if co-expression of palSAP97 with GluA1Δ7 rescued the dendrite growth promoting activity of this version of GluA1. Remarkably the combination of palSAP97 with GluA1Δ7 promoted dendrite growth to the same degree that the combination of SAP97 + GluA1 did (Zhou et al., 2008).

In our final set of in vitro studies we palmitoylated the version of SAP97 that contained mutations in PDZ2 that disrupted its physical association with GluA1 (mutPDZ2-palSAP97). In co-expression studies in heterologous cells we found that GluA1 will coIP palSAP97, but will not coIP mutPDZ2-palSAP97 (Zhou et al., 2008). So even though mutPDZ2-palSAP97 targets to the plasma membrane this is not sufficient to lead to a physical association with GluA1. We then asked about the dendrite growth promoting action of mutPDZ2-palSAP97 and we found that when co-expressed with GluA1, both palSAP97 and mutPDZ2-palSAP97 were equally effective in promoting dendrite growth (Zhou et al., 2008). Thus using two different strategies to disrupt the physical association of SAP97 with GluA1, we come to the same conclusion: co-expression of SAP97 with GluA1 synergistically promoted dendrite growth as long as both proteins are targeted to the plasma membrane. While the native proteins associate as part of a physical complex, experimental manipulations that de-link the two proteins demonstrate that co-localization, not physical interaction, are required for dendrite growth.

Potential Implications

Why should we care about this pathway of activity-dependent neuronal plasticity? One reason is that knowledge of this pathway may lead to ways of promoting plasticity in adults. One potential beneficiary might be individuals with spinal cord injury (R. G. Kalb, 2003). After a thoracic spinal cord lesion, the circuitry in the lumbar spinal cord can be engaged by repetitive activation of selected neuronal pathways (e.g., standing training, ambulation training) which results in remarkable improvement in motor behavior. This is seen both in experimental animals and humans (Barbeau & Rossignol, 1987; Dietz, Colombo, Jensen, & Baumgartner, 1995; Edgerton et al., 1997; Edgerton, Tillakaratne, Bigbee, de Leon, & Roy, 2004; Fung, Stewart, & Barbeau, 1990; Lovely, Gregor, Roy, & Edgerton, 1990; Rossignol, 2000; Wernig, Muller, Nanassy, & Cagol, 1995; Wernig, Nanassy, & Muller, 1998; Wirz, Colombo, & Dietz, 2001). The mechanism for this effect is use-dependent modification of spinal cord circuitry (Gazula, Roberts, Luzzio, Jawad, & Kalb, 2004) and so we think that enhancement of activity-dependent plasticity within the spinal cord will have a salubrious effect on functional recovery.

Another reason to study this form of activity-dependent neuronal plasticity relates to developmental disorders of brain. Abnormalities in dendrite structure (i.e., size, branching, and spines) are commonly seen in childhood diseases such as mental retardation, Autism and Autism-spectrum disorders (Dierssen & Ramakers, 2006; Kaufmann & Moser, 2000). Several lines of evidence indicate that in some forms of these childhood diseases, the primary defect is in activity-dependent development. Many genes linked to familial forms of impaired cognitive and emotional development are involved in activity-dependent synapse formation or stabilization (i.e., actin related proteins such as cofilin, LIMK and debrin; Rho-GTP’ase regulators such as oligophrenin-1 and Kalirin-7; and trophic factors such as BDNF, NRGN-ErbB4) (Lin & Koleske). In a study using homozygosity mapping to discover recessive disease genes in autistic patients (Morrow et al., 2008), significant genetic heterogeneity was found. One of the more remarkable findings of this study was that many autism-associated genes are regulated by neuronal activity. For example, the expression of the candidate gene DIA1 is regulated by activity and this transcription factor that controls the expression of other activity-regulated transcripts such as MEF2, NPAS4, CREB, EGR, SRF and others. If we start with the proposition that perturbation of experience-dependent cortical development underlies some of the defects in autism, then it is critical to understand the varieties of normal activity-dependent development. In this regard it is perhaps noteworthy that genetic studies link SAP97 to schizophrenia (Sato, Shimazu, Yamamoto, & Nishikawa, 2008; Toyooka et al., 2002) and autism (Willatt et al., 2005). It is possible that exploration of motor system development will provide a window onto previously unknown aspects of brain operation.

Acknowledgments

This work was supported in the past by the US Public Health Service (NS29837). We thank R. Sprengel, P Seeburg and R. Huganir for several of the murine strains used in these studies.

Cited Literature

  1. Aizawa H, Hu SC, Bobb K, Balakrishnan K, Ince G, Gurevich I, et al. Dendrite development regulated by CREST, a calcium-regulated transcriptional activator. Science. 2004;303(5655):197–202. doi: 10.1126/science.1089845. [DOI] [PubMed] [Google Scholar]
  2. Altman J, Sudarshan K. Postnatal development of locomotion in the laboratory rat. Animal Behavior. 1975;23:896–920. doi: 10.1016/0003-3472(75)90114-1. [DOI] [PubMed] [Google Scholar]
  3. Barbeau H, Rossignol S. Recovery of locomotion after chronic spinalization in the adult cat. Brain Res. 1987;412:84–95. doi: 10.1016/0006-8993(87)91442-9. [DOI] [PubMed] [Google Scholar]
  4. Bi G, Poo M. Synaptic modification by correlated activity: Hebb’s postulate revisited. Annu Rev Neurosci. 2001;24:139–166. doi: 10.1146/annurev.neuro.24.1.139. [DOI] [PubMed] [Google Scholar]
  5. Cai C, Coleman SK, Niemi K, Keinanen K. Selective binding of synapse-associated protein 97 to GluR-A alpha-amino-5-hydroxy-3-methyl-4-isoxazole propionate receptor subunit is determined by a novel sequence motif. J Biol Chem. 2002;277(35):31484–31490. doi: 10.1074/jbc.M204354200. [DOI] [PubMed] [Google Scholar]
  6. Carriedo SG, Yin HZ, Weiss JH. Motor neurons are selectively vulnerable to AMPA/KA receptor-mediated injury in vitro. J Neurosci. 1996;16:4069–4079. doi: 10.1523/JNEUROSCI.16-13-04069.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
  7. Chen SX, Tari PK, She K, Haas K. Neurexin-neuroligin cell adhesion complexes contribute to synaptotropic dendritogenesis via growth stabilization mechanisms in vivo. Neuron. 2010;67(6):967–983. doi: 10.1016/j.neuron.2010.08.016. [DOI] [PubMed] [Google Scholar]
  8. Chklovskii DB. Synaptic connectivity and neuronal morphology: two sides of the same coin. Neuron. 2004;43(5):609–617. doi: 10.1016/j.neuron.2004.08.012. [DOI] [PubMed] [Google Scholar]
  9. Cline H, Haas K. The regulation of dendritic arbor development and plasticity by glutamatergic synaptic input: a review of the synaptotrophic hypothesis. J Physiol. 2008;586(6):1509–1517. doi: 10.1113/jphysiol.2007.150029. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Cline HT. Dendritic arbor development and synaptogenesis. Curr Opin Neurobiol. 2001;11(1):118–126. doi: 10.1016/s0959-4388(00)00182-3. [DOI] [PubMed] [Google Scholar]
  11. Cline HT, Debski EA, Constantine-Paton M. N-methyl-D-aspartate receptor antagonist desegregates eye-specific columns. PNAS. 1987;84:4342–4345. doi: 10.1073/pnas.84.12.4342. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Constantine-Paton M, Cline HT, Debski E. Patterned activity, synaptic convergence, and the NMDA receptor in developing visual pathways. Annu Rev Neurosci. 1990;13:129–154. doi: 10.1146/annurev.ne.13.030190.001021. [DOI] [PubMed] [Google Scholar]
  13. Cramer KS, Angelucci A, Hahm JO, Bogdanov MB, Sur M. A role for nitric oxide in the development of the ferret retinogeniculate projection. J Neurosci. 1996;16:7995–8004. doi: 10.1523/JNEUROSCI.16-24-07995.1996. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Curfs MHJM, Gribnau AAM, Dederen PJWC. Selective elimination of transient corticospinal projections in the rat cervical spinal cord gray matter. Dev Brain Res. 1994;78:182–190. doi: 10.1016/0165-3806(94)90025-6. [DOI] [PubMed] [Google Scholar]
  15. Curfs MHJM, Gribnau AAM, Dideren PJWC. Postnatal maturation of the dendritic fields of motoneuron pools supplying flexor and extensor muscles of the distal forelimb in the rat. Development. 1993;117:535–541. doi: 10.1242/dev.117.2.535. [DOI] [PubMed] [Google Scholar]
  16. Dierssen M, Ramakers GJ. Dendritic pathology in mental retardation: from molecular genetics to neurobiology. Genes Brain Behav. 2006;5(Suppl 2):48–60. doi: 10.1111/j.1601-183X.2006.00224.x. [DOI] [PubMed] [Google Scholar]
  17. Dietz V, Colombo G, Jensen L, Baumgartner L. Locomotor capacity of spinal cord in paraplegic patients. Ann Neurol. 1995;37(5):574–582. doi: 10.1002/ana.410370506. [DOI] [PubMed] [Google Scholar]
  18. Donatelle JM. Growth of the Corticospinal tract and the development of placing reactions in the postnatal rat. J Comp Neurol. 1977;175:207–232. doi: 10.1002/cne.901750205. [DOI] [PubMed] [Google Scholar]
  19. Edgerton VR, de Leon RD, Tillakaratne N, Recktenwald MR, Hodgson JA, Roy RR. Use-dependent plasticity in spinal stepping and standing. Adv Neurol. 1997;72:233–247. [PubMed] [Google Scholar]
  20. Edgerton VR, Tillakaratne NJ, Bigbee AJ, de Leon RD, Roy RR. Plasticity of the spinal neural circuitry after injury. Annu Rev Neurosci. 2004;27:145–167. doi: 10.1146/annurev.neuro.27.070203.144308. [DOI] [PubMed] [Google Scholar]
  21. Engert F, Tao HW, Zhang LI, Poo MM. Moving visual stimuli rapidly induce direction sensitivity of developing tectal neurons. Nature. 2002;419(6906):470–475. doi: 10.1038/nature00988. [DOI] [PubMed] [Google Scholar]
  22. Fung J, Stewart JE, Barbeau H. The combined effects of clonidine and cyproheptadine with interactive training on the modulation of locomotion in spinal cord injured subjects. J Neurol Sci. 1990;100(1–2):85–93. doi: 10.1016/0022-510x(90)90017-h. [DOI] [PubMed] [Google Scholar]
  23. Gao FB, Bogert BA. Genetic control of dendritic morphogenesis in Drosophila. Trends Neurosci. 2003;26(5):262–268. doi: 10.1016/S0166-2236(03)00078-X. [DOI] [PubMed] [Google Scholar]
  24. Gaudilliere B, Konishi Y, de la Iglesia N, Yao G, Bonni A. A CaMKII-NeuroD signaling pathway specifies dendritic morphogenesis. Neuron. 2004;41(2):229–241. doi: 10.1016/s0896-6273(03)00841-9. [DOI] [PubMed] [Google Scholar]
  25. Gazula VR, Roberts M, Luzzio C, Jawad AF, Kalb RG. Effects of limb exercise after spinal cord injury on motor neuron dendrite structure. J Comp Neurol. 2004;476(2):130–145. doi: 10.1002/cne.20204. [DOI] [PubMed] [Google Scholar]
  26. Goodman CS, Shatz CJ. Developmental Mechanisms That Generate Precise Patterns of Neuronal Connectivity. Cell. 1993;10:77–98. doi: 10.1016/s0092-8674(05)80030-3. [DOI] [PubMed] [Google Scholar]
  27. Goold CP, Nicoll RA. Single-cell optogenetic excitation drives homeostatic synaptic depression. Neuron. 2011;68(3):512–528. doi: 10.1016/j.neuron.2010.09.020. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Ha S, Redmond L. ERK mediates activity dependent neuronal complexity via sustained activity and CREB-mediated signaling. Dev Neurobiol. 2008;68(14):1565–1579. doi: 10.1002/dneu.20682. [DOI] [PubMed] [Google Scholar]
  29. Hume RI, Purves D. Geometry of neonatal neurons and the regulation of synapse elimination. Nature. 1981;293:469–471. doi: 10.1038/293469a0. [DOI] [PubMed] [Google Scholar]
  30. Inglis FM, Crockett R, Korada S, Abraham WC, Hollmann M, Kalb RG. The AMPA receptor GluR1 regulates dendritic architecture of motor neurons. J Neurosci. 2002;22:8042–8051. doi: 10.1523/JNEUROSCI.22-18-08042.2002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Inglis FM, Furia F, Zuckerman KE, Strittmatter SM, Kalb RG. The role of nitric oxide and NMDA receptors in the development of motor neuron dendrites. J Neurosci. 1998;18:10493–10501. doi: 10.1523/JNEUROSCI.18-24-10493.1998. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Inglis FM, Zuckerman KE, Kalb RG. Experience-dependent development of spinal motor neurons. Neuron. 2000;26:299–305. doi: 10.1016/s0896-6273(00)81164-2. [DOI] [PubMed] [Google Scholar]
  33. Jakowec MW, Fox AJ, Martin LJ, Kalb RG. Quantitative and Qualitative Changes in AMPA Receptor Expression During Spinal Cord Development. Neuroscience. 1995;67:893–907. doi: 10.1016/0306-4522(95)00026-f. [DOI] [PubMed] [Google Scholar]
  34. Jakowec MW, Yen L, Kalb RG. In situ hybridization analysis of AMPA receptor subunit gene expression in the developing rat spinal cord. Neuroscience. 1995;67:909–920. doi: 10.1016/0306-4522(95)00094-y. [DOI] [PubMed] [Google Scholar]
  35. Jan YN, Jan LY. The control of dendrite development. Neuron. 2003;40(2):229–242. doi: 10.1016/s0896-6273(03)00631-7. [DOI] [PubMed] [Google Scholar]
  36. Jaworski J, Spangler S, Seeburg DP, Hoogenraad CC, Sheng M. Control of dendritic arborization by the phosphoinositide-3′-kinase-Akt-mammalian target of rapamycin pathway. J Neurosci. 2005;25(49):11300–11312. doi: 10.1523/JNEUROSCI.2270-05.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Jeong GB, Werner M, Gazula VR, Itoh T, Roberts M, David S, et al. Bi-directional control of motor neuron dendrite remodeling by the calcium permeability of AMPA receptors. Mol Cell Neurosci. 2006;32(3):299–314. doi: 10.1016/j.mcn.2006.04.008. [DOI] [PubMed] [Google Scholar]
  38. Kalb RG. Regulation of motor neuron dendrite growth by NMDA receptor activation. Development. 1994;120:3063–3071. doi: 10.1242/dev.120.11.3063. [DOI] [PubMed] [Google Scholar]
  39. Kalb RG. Getting the spinal cord to think for itself. Arch Neurol. 2003;60(6):805–808. doi: 10.1001/archneur.60.6.805. [DOI] [PubMed] [Google Scholar]
  40. Katz LC, Constantine-Paton M. Relationships between segregated afferents and postsynaptic neurons in the optic tectum of three-eyed frogs. J Neurosci. 1988;8:3160–3180. doi: 10.1523/JNEUROSCI.08-09-03160.1988. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Kaufmann WE, Moser HW. Dendritic anomalies in disorders associated with mental retardation. Cereb Cortex. 2000;10(10):981–991. doi: 10.1093/cercor/10.10.981. [DOI] [PubMed] [Google Scholar]
  42. Kim CH, Takamiya K, Petralia RS, Sattler R, Yu S, Zhou W, et al. Persistent hippocampal CA1 LTP in mice lacking the C-terminal PDZ ligand of GluR1. Nat Neurosci. 2005;8(8):985–987. doi: 10.1038/nn1432. [DOI] [PubMed] [Google Scholar]
  43. Kleinschmidt A, Bear MF, Singer W. Blockade of “NMDA” receptors disrupts experience-dependent plasticity of kitten striate cortex. Science. 1987;238:355–358. doi: 10.1126/science.2443978. [DOI] [PubMed] [Google Scholar]
  44. Kumar V, Zhang MX, Swank MW, Kunz J, Wu GY. Regulation of dendritic morphogenesis by Ras-PI3K-Akt-mTOR and Ras-MAPK signaling pathways. J Neurosci. 2005;25(49):11288–11299. doi: 10.1523/JNEUROSCI.2284-05.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Li S, Zhang C, Takemori H, Zhou Y, Xiong ZQ. TORC1 regulates activity-dependent CREB-target gene transcription and dendritic growth of developing cortical neurons. J Neurosci. 2009;29(8):2334–2343. doi: 10.1523/JNEUROSCI.2296-08.2009. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Li Z, Aizenman CD, Cline HT. Regulation of rho GTPases by crosstalk and neuronal activity in vivo. Neuron. 2002;33(5):741–750. doi: 10.1016/s0896-6273(02)00621-9. [DOI] [PubMed] [Google Scholar]
  47. Li Z, Van Aelst L, Cline HT. Rho GTPases regulate distinct aspects of dendritic arbor growth in Xenopus central neurons in vivo. Nat Neurosci. 2000;3(3):217–225. doi: 10.1038/72920. [DOI] [PubMed] [Google Scholar]
  48. Lin YC, Koleske AJ. Mechanisms of synapse and dendrite maintenance and their disruption in psychiatric and neurodegenerative disorders. Annu Rev Neurosci. 33:349–378. doi: 10.1146/annurev-neuro-060909-153204. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Lindsay AD, Greer JJ, Feldman JL. Phrenic Motoneuron Morphology in the Neonatal Rat. J Comp Neurol. 1991;308:169–179. doi: 10.1002/cne.903080204. [DOI] [PubMed] [Google Scholar]
  50. Lovely RG, Gregor RJ, Roy RR, Edgerton VR. Weight-bearing hindlimb stepping in treadmill-exercised adult spinal cats. Brain Res. 1990;514:206–218. doi: 10.1016/0006-8993(90)91417-f. [DOI] [PubMed] [Google Scholar]
  51. Lu B. Pro-region of neurotrophins: role in synaptic modulation. Neuron. 2003;39(5):735–738. doi: 10.1016/s0896-6273(03)00538-5. [DOI] [PubMed] [Google Scholar]
  52. McAllister AK, Katz LC, Lo DC. Neurotrophin regulation of cortical dendritic growth requires activity. Neuron. 1996;17(6):1057–1064. doi: 10.1016/s0896-6273(00)80239-1. [DOI] [PubMed] [Google Scholar]
  53. McAllister AK, Katz LC, Lo DC. Opposing roles for endogenous BDNF and NT-3 in regulating cortical dendrite growth. Neuron. 1997;18:767–778. doi: 10.1016/s0896-6273(00)80316-5. [DOI] [PubMed] [Google Scholar]
  54. Morais Cabral JH, Petosa C, Sutcliffe MJ, Raza S, Byron O, Poy F, et al. Crystal structure of a PDZ domain. Nature. 1996;382(6592):649–652. doi: 10.1038/382649a0. [DOI] [PubMed] [Google Scholar]
  55. Morrow EM, Yoo SY, Flavell SW, Kim TK, Lin Y, Hill RS, et al. Identifying autism loci and genes by tracing recent shared ancestry. Science. 2008;321(5886):218–223. doi: 10.1126/science.1157657. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Núñez-Abades PA, He F, Barrionuevo G, Cameron WE. Morphology of Developing Rat Genioglossal Motoneurons Studied In Vitro: Changes in Length, Branching Pattern, and Spatial Distribution of Dendrites. J Comp Neurol. 1994;339:401–420. doi: 10.1002/cne.903390308. [DOI] [PubMed] [Google Scholar]
  57. Palmer CL, Cotton L, Henley JM. The molecular pharmacology and cell biology of alpha-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptors. Pharmacol Rev. 2005;57(2):253–277. doi: 10.1124/pr.57.2.7. [DOI] [PMC free article] [PubMed] [Google Scholar]
  58. Pellis V, Pellis S, Teitelbaum A descriptive analysis of the postnatal development of contact-righting in rats (Rattus norvegicus) Devel Psychobiol. 1991;24(4):237–263. [Google Scholar]
  59. Peng YR, He S, Marie H, Zeng SY, Ma J, Tan ZJ, et al. Coordinated changes in dendritic arborization and synaptic strength during neural circuit development. Neuron. 2009;61(1):71–84. doi: 10.1016/j.neuron.2008.11.015. [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Redmond L, Kashani AH, Ghosh A. Calcium regulation of dendritic growth via CaM kinase IV and CREB-mediated transcription. Neuron. 2002;34(6):999–1010. doi: 10.1016/s0896-6273(02)00737-7. [DOI] [PubMed] [Google Scholar]
  61. Redmond L, Oh SR, Hicks C, Weinmaster G, Ghosh A. Nuclear Notch1 signaling and the regulation of dendritic development. Nat Neurosci. 2000;3(1):30–40. doi: 10.1038/71104. [DOI] [PubMed] [Google Scholar]
  62. Rossignol S. Locomotion and its recovery after spinal injury. Curr Opin Neurobiol. 2000;(10):708–716. doi: 10.1016/s0959-4388(00)00151-3. [DOI] [PubMed] [Google Scholar]
  63. Ruthazer ES, Akerman CJ, Cline HT. Control of axon branch dynamics by correlated activity in vivo. Science. 2003;301(5629):66–70. doi: 10.1126/science.1082545. [DOI] [PubMed] [Google Scholar]
  64. Sato J, Shimazu D, Yamamoto N, Nishikawa T. An association analysis of synapse-associated protein 97 (SAP97) gene in schizophrenia. J Neural Transm. 2008;115(9):1355–1365. doi: 10.1007/s00702-008-0085-9. [DOI] [PubMed] [Google Scholar]
  65. Seebach BS, Ziskind-Conhaim L. Formation of transient inappropriate sensorimotor synapses in developing rat spinal cord. J Neurosci. 1994;14:4520–4528. doi: 10.1523/JNEUROSCI.14-07-04520.1994. [DOI] [PMC free article] [PubMed] [Google Scholar]
  66. Shalizi A, Gaudilliere B, Yuan Z, Stegmuller J, Shirogane T, Ge Q, et al. A calcium-regulated MEF2 sumoylation switch controls postsynaptic differentiation. Science. 2006;311(5763):1012–1017. doi: 10.1126/science.1122513. [DOI] [PubMed] [Google Scholar]
  67. Shatz CJ. Impulse activity and the patterning of connections during CNS development. Neuron. 1990;5:745–756. doi: 10.1016/0896-6273(90)90333-b. [DOI] [PubMed] [Google Scholar]
  68. Sheng M, Sala C. PDZ domains and the organization of supramolecular complexes. Annu Rev Neurosci. 2001;24:1–29. doi: 10.1146/annurev.neuro.24.1.1. [DOI] [PubMed] [Google Scholar]
  69. Shepherd JD, Huganir RL. The Cell Biology of Synaptic Plasticity: AMPA Receptor Trafficking. Annu Rev Cell Dev Biol. 2007;23:613–643. doi: 10.1146/annurev.cellbio.23.090506.123516. [DOI] [PubMed] [Google Scholar]
  70. Sin WC, Haas K, Ruthazer ES, Cline HT. Dendrite growth increased by visual activity requires NMDA receptor and Rho GTPases. Nature. 2002;419(6906):475–480. doi: 10.1038/nature00987. [DOI] [PubMed] [Google Scholar]
  71. Snider WD, Zhang L, Yusoof S, Gorukanti N, Tsering C. Interactions between dorsal root axons and their target motor neurons in developing mammalian spinal cord. J Neurosci. 1992;12(9):3494–3508. doi: 10.1523/JNEUROSCI.12-09-03494.1992. [DOI] [PMC free article] [PubMed] [Google Scholar]
  72. Stegenga SL, Kalb RG. Developmental Regulation of N-methyl-D-aspartate - and Kainate-Type Glutamate Receptor Expression in the Rat Spinal Cord. Neuroscience. 2001;105:499–507. doi: 10.1016/s0306-4522(01)00143-9. [DOI] [PubMed] [Google Scholar]
  73. Stuart G, Spruston N, Hausser M. Dendrites. Oxford: Oxford University Press; 1999. [Google Scholar]
  74. Thyagarajan A, Ting AY. Imaging activity-dependent regulation of neurexin-neuroligin interactions using trans-synaptic enzymatic biotinylation. Cell. 2010;143(3):456–469. doi: 10.1016/j.cell.2010.09.025. [DOI] [PMC free article] [PubMed] [Google Scholar] [Retracted]
  75. Toyooka K, Iritani S, Makifuchi T, Shirakawa O, Kitamura N, Maeda K, et al. Selective reduction of a PDZ protein, SAP-97, in the prefrontal cortex of patients with chronic schizophrenia. J Neurochem. 2002;83(4):797–806. doi: 10.1046/j.1471-4159.2002.01181.x. [DOI] [PubMed] [Google Scholar]
  76. Vandenberghe W, Robberecht W, Brorson JR. AMPA receptor calcium permeability, GluR2 expression, and selective motoneuron vulnerability. J Neurosci. 2000;20(1):123–132. doi: 10.1523/JNEUROSCI.20-01-00123.2000. [DOI] [PMC free article] [PubMed] [Google Scholar]
  77. Vaughn JE. Fine structure of synaptogenesis in the vertebrate central nervous system. Synapse. 1989;3:255–285. doi: 10.1002/syn.890030312. [DOI] [PubMed] [Google Scholar]
  78. Vaughn JE, Barber RP, Sims TJ. Dendritic development and preferential growth into synaptogenic fields: a quantitative study of Golgi-impregnated spinal motor neurons. Synapse. 1988;2(1):69–78. doi: 10.1002/syn.890020110. [DOI] [PubMed] [Google Scholar]
  79. Walton KD, Harding S, Anschel D, Harris YT, Llinas R. The Effects of Microgravity on the Development of Surface Righting in Rats. J Physiol. 2005;565:593–608. doi: 10.1113/jphysiol.2004.074385. [DOI] [PMC free article] [PubMed] [Google Scholar]
  80. Walton KD, Lieberman D, Llinas A, Begin M, Llinas RR. Identification of a critical period for motor development in neonatal rats. Neuroscience. 1992;51:763–767. doi: 10.1016/0306-4522(92)90517-6. [DOI] [PubMed] [Google Scholar]
  81. Wayman GA, Impey S, Marks D, Saneyoshi T, Grant WF, Derkach V, et al. Activity-dependent dendritic arborization mediated by CaM-kinase I activation and enhanced CREB-dependent transcription of Wnt-2. Neuron. 2006;50(6):897–909. doi: 10.1016/j.neuron.2006.05.008. [DOI] [PubMed] [Google Scholar]
  82. Wernig A, Muller S, Nanassy A, Cagol E. Laufband therapy based on “rules of spinal locomotion” is effective in spinal cord injured persons. Eur J Neurosci. 1995;7:823–829. doi: 10.1111/j.1460-9568.1995.tb00686.x. [DOI] [PubMed] [Google Scholar]
  83. Wernig A, Nanassy A, Muller S. Maintainance of locomotor ability following Laufband (treadmill) therapy in para- and tetraplegia persons: follow-up studies. Spinal Cord. 1998;36:744–749. doi: 10.1038/sj.sc.3100670. [DOI] [PubMed] [Google Scholar]
  84. Willatt L, Cox J, Barber J, Cabanas ED, Collins A, Donnai D, et al. 3q29 microdeletion syndrome: clinical and molecular characterization of a new syndrome. Am J Hum Genet. 2005;77(1):154–160. doi: 10.1086/431653. [DOI] [PMC free article] [PubMed] [Google Scholar]
  85. Wirz M, Colombo G, Dietz V. Long term effects of locomotor training in spinal humans. J Neurol Neurosurg Psychiatry. 2001;71(1):93–96. doi: 10.1136/jnnp.71.1.93. [DOI] [PMC free article] [PubMed] [Google Scholar]
  86. Wu GY, Cline HT. Stabilization of dendritic arbor structure in vivo by CaMKII. Science. 1998;279:222–225. doi: 10.1126/science.279.5348.222. [DOI] [PubMed] [Google Scholar]
  87. Wu GY, Deisseroth K, Tsien RW. Spaced stimuli stabilize MAPK pathway activation and its effects on dendritic morphology. Nat Neurosci. 2001;4(2):151–158. doi: 10.1038/83976. [DOI] [PubMed] [Google Scholar]
  88. Wu HH, Williams CV, McLoon SC. Involvement of nitric oxide in the elimination of a transient retinotectal projection in development. Science. 1994;265:1593–1596. doi: 10.1126/science.7521541. [DOI] [PubMed] [Google Scholar]
  89. Yu X, Malenka RC. Beta-catenin is critical for dendritic morphogenesis. Nat Neurosci. 2003;6(11):1169–1177. doi: 10.1038/nn1132. [DOI] [PubMed] [Google Scholar]
  90. Zamanillo D, Sprengel R, Hvalby O, Hensen V, Burnashev N, Rozov A, et al. Importance of AMPA receptors for hippocampal synaptic plasticity but not spatial learning. Science. 1999;284:1805–1811. doi: 10.1126/science.284.5421.1805. [DOI] [PubMed] [Google Scholar]
  91. Zhang L, Schessl J, Werner M, Bonnemann C, Xiong G, Mojsilovic-Petrovic J, et al. Role of GluR1 in activity-dependent motor system development. J Neurosci. 2008;28(40):9953–9968. doi: 10.1523/JNEUROSCI.0880-08.2008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  92. Zhang LI, Tao HW, Poo M. Visual input induces long-term potentiation of developing retinotectal synapses. Nat Neurosci. 2000;3(7):708–715. doi: 10.1038/76665. [DOI] [PubMed] [Google Scholar]
  93. Zhou W, Zhang L, Guoxiang X, Mojsilovic-Petrovic J, Takamaya K, Sattler R, et al. GluR1 controls dendrite growth through its binding partner, SAP97. J Neurosci. 2008;28(41):10220–10233. doi: 10.1523/JNEUROSCI.3434-08.2008. [DOI] [PMC free article] [PubMed] [Google Scholar]
  94. Zou DJ, Cline HT. Postsynaptic calcium/calmodulin-dependent protein kinase II required to limit elaboration of presynaptic and postsynaptic neuronal arbors. J Neurosci. 1999;19:8909–8919. doi: 10.1523/JNEUROSCI.19-20-08909.1999. [DOI] [PMC free article] [PubMed] [Google Scholar]

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