New Results

Rap2 and TNIK control Plexin-dependent synaptic tiling in C. elegans

Xi Chen, Akihiro C. E. Shibata, Nicholas L Weilinger, Ardalan Hendi, Mizuki Kurashina, Ethan Fortes, Brian MacVicar, Hideji Murakoshi, Kota Mizumoto
doi: https://doi.org/10.1101/238352

Abstract

During development, neurons form extensive synaptic connectivity with their fate-determined targets. Ectopic synapse formation with aberrant targets underlie various neurological disorders including autism. While we are beginning to elucidate the underlying mechanisms by which extracellular ligand-receptor interactions enhance synapse specificity by inhibiting synaptogenesis, our knowledge about their intracellular mechanisms remains limited. Here we show that Rap2 GTPase (rap-2) and its effector, TNIK (mig-15), act downstream of Plexin (plx-1) signaling to restrict presynaptic assembly and to form tiled synaptic innervation of two cholinergic motor neurons (DA8 and DA9) in C. elegans. Both constitutively GTP- and GDP-forms of rap-2 mutants exhibit similar synaptic tiling defects as plx-1 mutants, suggesting that cycling of the RAP-2 nucleotide state is critical for synapse inhibition. Consistently, RAP-2 activity is suppressed in a short segment of axon lacking synapses where PLX-1 is enriched. Excessive ectopic synapse formation in mig-15 mutants causes expansion of the synaptic domains in DA8 and DA9, which induces a severe synaptic tiling defect. Conversely, overexpression of mig-15 strongly inhibited synapse formation, which suggests that mig-15 is a negative regulator of synapse formation. These results reveal that subcellular regulation of small GTPase activity by Plexin shapes proper synapse patterning in vivo.

Introduction

During nervous system development, various instructive and repulsive signaling cues cooperatively direct neurons to form chemical synapses with their appropriate targets. Studies have identified some molecules and elucidated their downstream mechanisms that instruct synaptogenesis such as FGF, Ephrin/Eph, Ig-family of cell adhesion molecules (IgCAMs) and synaptic cell adhesion molecules (SynCAMs) (Shen and Bargmann, 2003; Shen et al., 2004). Several axon guidance cues and their receptors also play critical roles to inhibit synapse formation (Inaki et al., 2007). Semaphorins (Sema) and their receptors, Plexins, are two conserved families of molecules that have a well-established function to repel axons during development (Kolodkin et al., 1993; 1992; Negishi et al., 2005; Tran et al., 2007) and prominent roles contributing to immune system, cardiovascular development and cancer regulation (Epstein et al., 2015; Neufeld et al., 2005; Takamatsu and Kumanogoh, 2012).

In addition to its function as a long-range axon guidance cue during neuronal development, Sema/Plexin signaling plays a critical role as a negative regulator of synapse formation. The role of Sema/Plexin signaling in inhibiting synapse formation was first observed in Drosophila, where ectopic expression of Sema-2a causes elimination of specific neuromuscular junctions (Matthes et al., 1995). In mammals, Sema3E/PlexinD1 specifies sensory-motor connections (Pecho-Vrieseling et al., 2009). Secreted Sema3F locally inhibits spine development through its receptors PlexinA3 and Neuropilin-2 in hippocampal granule cells (Tran et al., 2009), and Sema5A/PlexinA2 signaling inhibits excitatory synapse formation in dentate granule cells (Duan et al., 2014). Sema5B diminishes synaptic connections in cultured hippocampal neurons (O’Connor et al., 2009). However, little is known about the intracellular mechanisms through which Sema/Plexin signaling inhibits synapse formation.

The cytoplasmic domain of Plexin contains a GAP (GTPase-activating protein) domain that inactivates small GTPases (Oinuma et al., 2004; Rohm et al., 2000). Upon activation by Semaphorins, Plexins repel axon outgrowth by inhibiting R-Ras (Negishi et al., 2005). Recent biochemical and structural analyses demonstrated the GAP domain of mammalian PlexinA3 is specific for Rap GTPases, which belong to the Ras family of GTPases and regulate actin cytoskeleton (Wang et al., 2012; 2013). PlexinA3 dimerization by Semaphorin activates its GAP domain, thereby inhibiting Rap1 from inducing neurite retraction. Drosophila PlexA and zebrafish PlexinA1 promote remodeling of epithelial cells by inhibiting Rap1 GTPase during wound healing (Yoo et al., 2016). Another Rap GTPase, Rap2, can inhibit neurite outgrowth (Kawabe et al., 2010). Similar to Sema/Plexin signaling, Rap GTPases regulate synapse formation and function. Rap2 negatively regulate spine number in cultured hippocampal neurons (Fu et al., 2007). Rap1 and Rap2 regulate synaptic activity by removing AMPA receptors from spines during long-term depression and depotentiation, respectively (J. J. Zhu et al., 2002; Y. Zhu et al., 2005) While the GAP domain of Plexin is critical to inhibit synapse formation (Duan et al., 2014; Mizumoto and Shen, 2013a), we still do not know whether Plexin regulates synapse patterning via Rap GTPases at presynaptic sites.

In Caenorhabditis elegans, Sema/Plexin signaling functions in vulva formation and male ray development (Dalpé et al., 2005; 2004; Fujii et al., 2002; Ikegami et al., 2004; Liu et al., 2005; Nakao et al., 2007; Nukazuka et al., 2008; 2011). Using this model system, we previously reported that Sema/Plexin signaling in the nervous system mediates a critical inter-axonal interaction for the tiled synaptic innervation of two DA-class cholinergic motor neurons (DA8 and DA9) (Mizumoto and Shen, 2013a). Cell bodies of nine DA neurons in C. elegans reside in the ventral nerve cord, sending dendrites ventrally and axons dorsally to form en passant synapses onto the dorsal body wall muscles. Even though axons of DA neurons show significant overlap, each motor neuron forms synapses onto muscles within specific sub-axonal domains, which do not overlap with those from neighboring DA neurons. This unique synaptic innervation creates tiled synaptic patterns along the nerve cord (White et al., 1986).

Tiled synaptic innervation occurs within most motor neuron classes and may contribute to the sinusoidal locomotion pattern of C. elegans (White et al., 1986). Using a combination of two fluorescent proteins (GFP and mCherry) fused with the presynaptic vesicle protein, RAB-3, and two tissue specific promoters (Figure 1) (Mizumoto and Shen, 2013a), we can visualize this synaptic tiling between DA8 and DA9 neurons. We reported that PLX-1 localizes at the anterior edge of the DA9 synaptic domain in axon-axon interactions in a Semaphorin-dependent manner, where it locally inhibits formation of the presynaptic specialization via its GAP domain. Loss of Semas or plx-1 causes anterior expansion of DA9 synaptic domain and posterior expansion of DA8 synaptic domain. This result indicates loss of inter-axonal interactions between DA8 and DA9 neurons. However, Tran et al., also observed excess dendritic spine formation, specifically within the region close to the cell body, in the plexin knockout mouse (Tran et al., 2009). These findings suggest a conserved mechanism by which Sema/Plexin locally inhibits synapse formation.

Figure 1.
Figure 1. Gain- and loss-of-function rap-2 mutants show synaptic tiling defects

(A and B) Representative image of synaptic tiling of wild type (A) and plx-1 mutant (B) animals. Images show wyIs446 marker to label synapses in DA8 (GFP::RAB-3) and DA9 (GFP::RAB-3+mCherry::RAB-3). Dotted box represents the magnified images from above of the synaptic tiling border. Schematics of DA8 (green) and DA9 (magenta) neurons with parameters for analysis shown on the right. (C - F) Representative images of wyIs446 strains with the following genotypes: rap-2(G12V) overexpression in DA neurons (C), rap-2 G12V (miz18) (D), rap-2 null (gk11) (E) and rap-2 S17A (miz19) (F). Synaptic domains from DA8 and DA9 are highlighted with green and magenta lines, respectively. Asterisks: DA9 cell body. Arrows: dorsal commissure of DA9. Scale bars: 20μm. (G) Quantification of overlap between DA8 and DA9 synaptic domains. Each dot represents a single animal. Blue bars indicate mean ± SEM. n.s.: not significant; ***: p < 0.001.

We previously reported that let-60/KRas gain-of-function mutants showed mild synaptic tiling defects. Since mammalian Plexin acts as a RapGAP and very mild synaptic tiling defects of let-60(gf) mutants, we hypothesized that Rap GTPase is the major downstream effector of PLX-1 to regulate synaptic tiling of DA neurons. Here, we report that rap-2, a C. elegans ortholog of human Rap2A, and its effector kinase mig-15 (TNIK: Traf2- and Nck-interacting kinase) act downstream of PLX-1 to regulate synaptic tiling. plx-1 delineates the border of synaptic tiling by locally inhibiting rap-2 along the DA9 axon and mig-15 inhibits synapse formation. Our results reveal the molecular mechanism underlying Plexin signaling to form fine synaptic map connectivity.

Results

rap-2 functions downstream of plx-1 to regulate synaptic tiling

Three Rap genes exist in the C. elegans genome (rap-1, rap-2 and rap-3). To delineate which Rap GTPase functions downstream of PLX-1 in synaptic tiling between DA8 and DA9 neurons, we first examined the expression patterns of all three rap genes (Figures S1). Among them, only rap-2, an ortholog of mammalian Rap2a, was expressed in motor neurons including DA8 and DA9, while rap-1 and rap-3 were not expressed in these cells (Figures S1A-C). In wild type animals, synaptic domains of DA8 and DA9 neurons did not show significant overlap, creating tiled synaptic innervation (Figures 1A and 1G). In the plx-1(nc36) null mutant, synaptic domains of DA8 and DA9 expanded posteriorly and anteriorly, respectively. As a result, synaptic domains of these neurons overlapped significantly (Figures 1B and 1G).

Since the intracellular domain of Plexin contains a RapGAP domain, we hypothesized that RAP-2 preferentially exists in a GTP-bound form in the plx-1 mutants. As the DA and VA classes of cholinergic neurons express unc-4, over-expression of a constitutively GTP-bound form of rap-2(G12V) under the unc-4 promoter elicited a similar synaptic tiling defect as plx-1 mutants (Figures 1C and 1G). We then generated rap-2(G12V) mutants using CRISPR/Cas9 genome editing. We observed the same synaptic tiling defects in three independent rap-2(G12V) mutant alleles (miz16, miz17 and miz18) as in plx-1 mutants (Figures 1D, 1G and S2). We found a comparable level of gene expression among all three rap-2(G12V) mutants to wild type rap-2 using RT-qPCR. These results indicate that the rap-2(G12V) mutation itself, not changes in gene expression, underlie the synaptic tiling defect in rap-2(G12V) mutants (Figure S2B).

Surprisingly, the null mutant of rap-2(gk11) also showed the same synaptic tiling defect, as did the constitutively GTP-form of rap-2(G12V) mutants (Figures 1E and 1G). This result suggests that synaptic tiling requires both the GTP- and GDP-bound forms of RAP-2. Indeed, the constitutively GDP-bound form of rap-2 mutants (S17A: miz19, miz20) showed the identical synaptic tiling defect as rap-2(G12V) and rap-2(gk11) mutants (Figures 1F, 1G and Figure S2). These results suggest that the cycling between GTP- and GDP-forms of RAP-2 is critical to regulate the spatial patterning of synapses.

Similar to plx-1 mutants, the synaptic tiling defect in all rap-2 mutants is caused by both the posterior expansion of DA8 synaptic domain and the anterior expansion of the DA9 synaptic domain (Figures S2C and S2D). However, none of rap-2(G12V), rap-2(S17A) and rap-2(gk11) mutants enhanced or suppressed the synaptic tiling defect in plx-1 mutants (Figure 1G). These results suggest that plx-1 and rap-2 function in the same genetic pathway. Consistent with the expression patterns of rap-1 and rap-3, neither rap-1(pk2082) nor rap-3(gk3975) null mutants showed significant synaptic tiling defects by themselves and did not enhance the synaptic tiling defect in rap-2(gk11) null mutants (Figures S1D-G). All mCherry::RAB-3 puncta co-localized with an active zone marker, CLA-1 (Xuan et al., 2017) in the DA9 neurons of plx-1(nc36) and rap-2(gk11) mutants, suggesting that RAB-3 puncta represent functional synapses (Figure S3). Taken together, these data indicate that rap-2 is a Rap GTPase that acts downstream of plx-1 for synaptic tiling in DA neurons.

RAP-2 functions cell autonomously in DA neurons

We next determined the cellular location for rap-2 function. Since rap-2(gk11) null mutants showed a synaptic tiling defect, we conducted tissue specific rescue experiments using tissue specific promoters as previously described (Mizumoto and Shen, 2013a). Expression of rap-2 in the post-synaptic body wall muscle cells under the hlh-1 promoter or in another class of cholinergic motor neurons in the dorsal nerve cord (DB neurons) under the truncated unc-129 promoter did not rescue the synaptic tiling defect in rap-2(gk11) animals (Figures 2A, 2B and 2G). However, DA neuron-specific expression using the unc-4c promoter strongly rescued the synaptic tiling defect (Figures 2C and 2G). DA9-specific expression of rap-2 under the mig-13 promoter partially rescued the synaptic tiling defect (Figures 2E and 2G). DA9-specific expression of rap-2 rescued the phenotype of anterior expansion of the DA9 synaptic domain but not the posterior expansion of DA8 synaptic domain (Figures 2H and 2I). These results suggest that rap-2 regulates synapse patterning in a cell-autonomous manner. In contrast to the DA9-specific rescue experiment in rap-2 mutants, DA9-specific expression of plx-1 cDNA was sufficient to rescue synaptic defects in both DA9 and DA8 (Mizumoto and Shen, 2013a) (see discussion).

Figure 2.
Figure 2. rap-2 functions in DA neurons

(A - F) Representative images of rap-2; wyIs446 animals expressing Phlh-1::rap-2 (A), Punc-129::rap-2 (B), Punc-4c::rap-2 (C), Punc-4c::Rap2a (human) (D), Pmig-13::rap-2 (E) and Punc-4c;;rap-1 (F). Synaptic domains of DA8 and DA9 are highlighted with green and magenta lines, respectively. Asterisks: DA9 cell body. Arrows: dorsal commissure of DA9. Scale bars: 20μm. (G - I) Quantification of DA8/DA9 overlap (G), DA9 synaptic domain (H) and DA8 asynaptic domain (I). Each dot represents a single animal. Blue bars indicate mean ± SEM. n.s.: not significant; ***: p < 0.001; ** : p < 0.01; * : p < 0.05.

We also observed that expression of human Rap2a in DA neurons rescued the synaptic tiling defect of rap-2 mutants, suggesting the function of rap-2 in synapse patterning is conserved across species (Figures 2D and 2G). Previous work suggested a partial functional redundancy between rap-1 and rap-2 in C. elegans (Pellis-van Berkel et al., 2005). However, we found that rap-1 expression in DA neurons did not rescue the synaptic tiling defect of rap-2 mutants, suggesting functional diversity between rap-1 and rap-2 (Figures 2F and 2G). Taken together, we conclude that rap-2 functions cell autonomously in DA neurons to regulate synaptic tiling.

RAP-2 activity is spatially regulated by PLX-1

Previously, we demonstrated that PLX-1::GFP is localized at the anterior edge of the DA9 synaptic domain, where it negatively regulates synapse formation through its cytoplasmic GAP domain (Figure 3A) (Mizumoto and Shen, 2013a). In the rap-2(gk11) mutant background, we observed no change in PLX-1::GFP localization but did observe ectopic synapses in the axonal region anterior to the PLX-1::GFP domain (Figure 3B). This result is consistent with our hypothesis that rap-2 acts downstream of plx-1 to regulate synaptic tiling. Together with our findings that synaptic tiling requires both GTP- and GDP-bound forms of RAP-2, we speculate that PLX-1 acting at the anterior edge of the DA9 synaptic domain regulates the spatial activity of RAP-2 along the axon.

Figure 3.
Figure 3. PLX-1 locally inhibits Rap2 activity in DA9

(A and B) Representative image of PLX-1::GFP alone (left) and PLX-1::GFP with mCherry::RAB-3 (right) labeled with wyIs320 in DA9 of wildtype (A) and rap-2 (gk11) (B). Bracket indicates PLX-1::GFP localized at the anterior edge of the DA9 synaptic domain. Arrowheads indicate ectopic synapses formed anterior to the PLX-1:: GFP patch. Scale bars: 20μm. (C) Schematic of Rap2 FRET sensor system. Binding of mRFP-RalGDS(RBD)-mRFP to EGFP-Rap2 induces FRET from EGFP to mRFP, leading to decreased EGFP fluorescence lifetime. (D) Representative epifluorescence (top) and fluorescence lifetime images (bottom) of HeLa cells expressing Rap2 FRET sensor (mRFP-RalGDS(RBD)-mRFP) with either constitutively GTP(G12V)- or GDP(S17A)-forms of human Rap2a. (E) Quantification of the binding fraction of Rap2 mutants. Binding fractions were measured from fluorescence decay curves of individual cells (G12V, n=25; S17A, n=25), as described previously (Murakoshi et al., 2011). Data are shown as mean ± SEM. Asterisks denote statistical significance (p<0.05, student’s t-test). (F) Representative images of fluorescence lifetime (top) and epifluorescence (bottom) of mizIs19 in wild type (left) and plx-1 mutants (right). White arrowheads indicate the position of the putative synaptic tiling border, as judged by the slight dorsal shift of the DA9 axon, as reported previously (Mizumoto and Shen, 2013a). (G) Quantification of the difference in binding fraction of GTP-Rap2a and RalGDS(RBD) between synaptic region (dotted boxes in F) and anterior asynaptic region (solid boxes in F). The binding fraction measured in synaptic region (dotted rectangle) was subtracted by that in non-synaptic region (solid rectangle). Five micrometers along the axon line from the synaptic tiling border were used for quantification. Data are presented as mean ± SEM (wild type, n=22; plx-1, n=17). (H) Representative images of fluorescence lifetime (top) and epifluorescence (bottom) of mizIs19 in unc-104 (left) and unc-104; plx-1 double mutants (right). (I) Quantification of the difference in binding fraction of GTP-Rap2a and RalGDS(RBD) between synaptic region (dotted boxes in H) and anterior asynaptic region (solid boxes in H). The binding fraction measured in synaptic region (dotted rectangle) was subtracted by that in non-synaptic region (solid rectangle). Five micrometers along the axon line from the synaptic tiling border were used for quantification. Data are presented as mean ± SEM (unc-104, n=21; unc-104;plx-1, n=23).

So, we sought to determine the spatial distribution of GTP-RAP-2 in DA9 axons. We conducted Fluorescence Lifetime Imaging Microscopy (FLIM)-based FRET (Förster Resonance Energy Transfer) measurements using EGFP-Rap2A (human) and mRFP-RalGDS(RBD: Ras Binding Domain)-mRFP (Yasuda et al., 2006). As RalGDS-RBD specifically binds to GTP-Rap2 but not GDP-Rap2 (Ohba et al., 2000), FRET from eGFP-Rap2a to mRFP-RalGDS(RBD)-mRFP can be used as a readout of Rap2 activity. We detected FRET signal as a change of GFP fluorescence lifetime (Figure 3C). In HeLa cells, we observed a shorter lifetime of constitutively bound GTP construct EGFP-Rap2A(G12V) compared to GDP-bound EGFP-Rap2A(S17A), indicating that the FRET sensor can detect the nucleotide state of Rap2a (Figures 3D and 3E). Due to the low expression of C. elegans RAP-2 constructs in HeLa cells, we were not able to test whether the mammalian FRET sensor can detect C. elegans RAP-2 activity (data not shown).

We then expressed EGFP-Rap2a and mRFP-RalGDS(RBD)-mRFP FRET sensors in DA9 neurons in C. elegans. As human Rap2a rescued the synaptic tiling defect of rap-2(gk11) mutants (Figure 2G), we reasoned that the activity pattern of human Rap2a should recapitulate that of endogenous RAP-2. We indeed observed lower Rap2a activity at the anterior edge of the DA9 synaptic domain compared to within the synaptic domain (Figures 3F and 3G). This observation is consistent with the localization of PLX-1::GFP at the anterior edge of DA9 synaptic domain (Figure 3A) (Mizumoto and Shen, 2013a). Local inhibition of Rap2a activity was strongly diminished in the plx-1 mutant background (Figures 3F and 3G). Previously, we showed that the PLX-1::GFP localization was unaffected in unc-104/Kif1A mutants, while synapses are absent from all axons including DA9 (Mizumoto and Shen, 2013a). We observed the same local inhibition of Rap2A activity at the putative synaptic tiling border in wild type, but not in plx-1(nc36), animals (Figure 3H and 3I). These results strongly suggest that Plexin localized at the anterior edge of the DA9 synaptic domain locally inactivates Rap2 GTPase to delineate the synaptic tiling border in DA9.

mig-15(TNIK) regulates synaptic tiling

In mammals, TNIK (Traf2 and Nck1-interacting kinase) acts with Rap2 to regulate neurite extension and AMPA receptor trafficking in hippocampal neurons and microvilli formation in intestinal cells (Hussain et al., 2010; Kawabe et al., 2010). In C. elegans, mig-15 is the sole ortholog of mammalian TNIK and its paralog MINK1 (Misshapen-like kinase 1), which is also an effector of Rap GTPase (Nonaka et al., 2008). mig-15 can regulate various cellular processes, such as axon guidance and cell migration (Chapman et al., 2008; Poinat et al., 2002; Shakir et al., 2006; Teulière et al., 2011). We found that mig-15(rh148) hypomorphic mutants showed a severe synaptic tiling defect (Figures 4A and 4D). Similar to plx-1 and rap-2 mutants, the synaptic tiling defect followed the anterior expansion of the DA9 synaptic domain and the posterior expansion of the DA8 synaptic domain (Figures 4E and 4F). All RAB-3 puncta in mig-15(rh148) mutants are co-localized with an active zone marker, CLA-1 (Figure S3), suggesting that these RAB-3 puncta represent synapses. We observed axon guidance defects (23%, n=100) or ectopic branch formation (56%, n=100) in DA9 of mig-15 mutant animals (Figure S4). These were excluded from our analysis of synaptic tiling phenotypes. Whereas only half of the mig-15 mutant animals showed guidance defects or ectopic branch formation, the synaptic tiling defect of mig-15 mutants was almost fully penetrant (Figure 4D). These data suggest that the synaptic tiling defect is not a secondary effect of axon outgrowth and guidance.

Figure 4.
Figure 4. mig-15(TNIK) mutants show a severe synaptic tiling defect

(A-C) Representative images of synaptic tiling marker (wyIs446) in mig-15(rh148) (A), rap-2(gk11); mig-15(rh148) (B) and plx-1(nc36); mig-15(rh148) (C) mutants. Synaptic domains of DA8 and DA9 are highlighted with green and magenta lines, respectively. Asterisks: DA9 cell body. Arrows: dorsal commissure of DA9. Scale bars: 20μm. (D-F) Quantification of overlap between DA8 and DA9 synaptic domains (D), DA9 synaptic domain (E) and DA8 asynaptic domain (F) in respective mutant backgrounds. Each dot represents measurement from single animal. Blue bars indicate mean ± SEM. n.s.: not significant; *** : p < 0.001.

The other two nonsense alleles (rh326: Q439Stop, rh80: W898Stop) also showed identical synaptic tiling defects as mig-15(rh148) (Figures S5A-5E). mig-15(rh80) has a nonsense mutation within the highly conserved CNH (citron/NIK homology) domain, which is required to interact with Rap2 in both mammals and C. elegans (Taira et al., 2004). This suggests a physical interaction between RAP-2 and MIG-15 for synaptic tiling. plx-1 or rap-2 mutants did not enhance the synaptic tiling defect in mig-15 mutants (Figures 4B-4F). This result is consistent with the hypothesis that mig-15 acts in the same genetic pathway as plx-1 and rap-2. Interestingly, the degree of overlap between DA8 and DA9 synaptic domains was even larger in mig-15 mutants than those observed in plx-1 and rap-2 mutants (compare Figures 1G and 4D). Taken together, these results suggest that mig-15 also acts downstream of additional signaling pathways.

mig-15 functions in DA neurons

We then sought to determine in which cells mig-15 functions by conducting tissue specific rescue experiments. Since several mig-15 isoforms (wormbase and data not shown) exist, we used the mig-15 genomic sequence for the rescue experiments. Expression of mig-15 under the DA neuron specific promoter (Punc-4c) strongly rescued the synaptic tiling defect of mig-15(rh148) mutants (Figures 5A and 5F), consistent with our hypothesis that mig-15 acts in the same genetic pathway as plx-1 and rap-2.

Figure 5.
Figure 5. mig-15 functions in DA neurons

(A-D) Representative images of synaptic tiling marker (wyIs446) in the following backgrounds: mig-15(rh148); Ex[Punc-4c::mig-15] (A), mig-15(rh148); Ex[Pmig-13::mig-15] (B), wild type (C) and wild type animals expressing dominant-negative mig-15(K50A) in DA neurons (D). Synaptic domains of DA8 and DA9 are highlighted with green and magenta lines, respectively. Asterisks: DA9 cell body. Arrows: dorsal commissure of DA9. Scale bars: 20μm. (E) Amino acid alignment of amino-terminal region of MIG-15 and TNIK. A kinase-dead mutation in TNIK (K54A) and corresponding mutation in MIG-15 (K50A) are highlighted in red. (F-I) Quantification of overlap between DA8 and DA9 synaptic domains (F and I), DA9 synaptic domain (G), DA8 asynaptic domain (H) in respective mutant backgrounds. Each dot represents measurements from a single animal. Blue bars indicate mean ± SEM. n.s.: not significant; *** : p < 0.001; * : p < 0.05.

Expression of mig-15 in both DA8 and DA9 rescued both posterior expansion of the DA8 synaptic domain and anterior expansion of the DA9 synaptic domain (Figures 5G and 5H). DA9 specific expression of mig-15 under the mig-13 promoter rescued anterior expansion of the DA9 synaptic domain, suggesting that mig-15 functions cell autonomously in DA9 (Figures 5B and 5G). We observed that Pmig-13::mig-15 weakly rescued the posterior expansion of the DA8 synaptic domain (Figure 5H). This is likely due to the leaky expression of mig-15 in DA8, as the mig-15 genomic fragment without promoter showed slight rescue of the synaptic tiling defect in mig-15(rh148) mutants (Figure 5F). Kinase dead TNIK mutants act as a dominant-negative (Mahmoudi et al., 2009). Expression of mutant mig-15(kd), which carries the same mutation at the corresponding amino acid of the dominant-negative TNIK (Figure 5E), in DA neurons caused a severe synaptic tiling defect (Figures 5C, 5D and 5I). Based on these results, we conclude that mig-15 functions cell autonomously in DA neurons.

mig-15 inhibits synapse formation

We observed that DA9-specific expression of mig-15 under the mig-13 promoter in mig-15 mutants often exhibited a shorter synaptic domain compared to wild type (Figures 5B and 5G). So, we speculated that an excess amount of mig-15 inhibits synapse formation. We tested the effect of mig-15 overexpression in the wild type background. Strikingly, DA9-specific mig-15 overexpression in wild type (mig-15(OE)) significantly reduced synapse number compared to wild type (Figures 6A, 6C, 6D and S6). This reduction occurred without affecting overall morphology of the DA9 neuron (Figure S4). Conversely, DA9 synapse number was significantly increased in the mig-15(rh148) mutants (Figures 6B, 6D and S6). Similarly, mig-15 overexpression significantly reduced synapse number in DD-type GABAergic motor neurons (Figure S7). These results indicate that mig-15 is a negative regulator of synapse formation. However, pan-neuronal expression of mig-15 under the rab-3 promoter caused severe uncoordinated locomotion in wildtype animals (Figure S8). These locomotor defects occurred concomitant with significantly reduced GFP::RAB-3 intensity in the dorsal nerve cord in mig-15 over-expressing animals and without causing significant axon guidance defects (Figure S8). Taken together, these data indicate that reduced synapse number by mig-15 overexpression disrupted the proper functioning of the motor circuit.

Figure 6.
Figure 6. mig-15 negatively regulates synapse number

(A-C) Representative images of DA9 presynaptic marker (wyIs85) in wild type (A), mig-15(rh148) (B) and mig-15 overexpressing animals (C). Brackets represent DA9 synaptic domain. Asterisks: DA9 cell body. Arrows: dorsal commissure of DA9. Scale bars: 20μm. (D) Quantification of DA9 synapse number. Each dot represents measurements from a single animal. Blue bars indicate mean ± SEM. n.s.: not significant; *** : p < 0.001; * : p < 0.05. (E-H) Representative images of synaptic branched F-actin labeled with GFP::utCH (wyIs329) in wild type (E), rap-2(gk11) (F), mig-15(rh148) (G) and mig-15 overexpressing animals (H). (I) Quantification of the length of GFP::utCH. Distance from the dorsal commissure to the most anterior and brightest GFP spot was measured. Blue bars indicate mean ± SEM. n.s.: not significant; ***: p < 0.001; **: p < 0.01, * : p < 0.05.

Rap GTPase and TNIK are well-known actin cytoskeleton regulators (Lin et al., 2010; 2008; Taira et al., 2004). Previous studies demonstrated that ARP2/3-dependent branched F-actin is required for presynaptic development (P. Chia et al., 2012; P. H. Chia et al., 2014). Branched F-actin visualized by GFP::ut-CH (utrophin calponin homology domain) is enriched within the DA9 synaptic domain (Figure 6E) (P. Chia et al., 2012; Mizumoto and Shen, 2013a). We predicted that mig-15 negatively regulates synapse formation by re-organizing the branched F-actin in the synaptic region. We did observe longer synaptic F-actin distribution in rap-2(gk11) and mig-15(rh148) mutants (Figures 6F, 6G and 6I). Conversely, overexpression of mig-15 in DA9 significantly decreased the length of synaptic F-actin (Figures 6H and 6I). Overexpression of mig-15 also appeared to decrease the overall amount of synaptic F-actin (Figure 6H). These results suggest that mig-15 inhibits synapse formation by negatively regulating the formation of synaptic F-actin.

plx-1 and rap-2 can coordinate the position of synaptic tiling border and synapse number

mig-15(OE) caused a reduction of synapse number in DA9. As a result, the length of DA9 synaptic domain was significantly reduced in mig-15(OE) animals than in wild type (Figures 7A and 7E). Despite this, synaptic tiling is maintained without a significant gap between DA8 and DA9 synaptic domains in mig-15(OE) animals, suggesting that the position of synaptic tiling border has shifted posteriorly in mig-15(OE) animals. Indeed, the length of the posterior asynaptic domain of DA8 was significantly shorter in mig-15(OE) animals, indicating that the DA8 synaptic domain expanded posteriorly (Figure 7D). This result strongly suggests that the PLX-1/RAP-2 signaling pathway can specify the position of synaptic tiling border according to the available number of synapses in each DA neuron. It is therefore likely that synaptic tiling is a mechanism to maintain the uniform distribution of the synapses from one class of motor neuron in the nerve cord. Consistently, DA8 synaptic domain did not shift posteriorly when mig-15 was overexpressed in DA9 of the synaptic tiling mutants, plx-1 or rap-2 (Figures 7B-7D). This result suggests that DA8 no longer senses the reduction of DA9 synapse number in the synaptic tiling mutants.

Figure 7.
Figure 7. PLX-1/RAP-2 signaling coordinates synapse number and synaptic tiling border

(A-C) Representative images of synaptic tiling marker (wyIs446) overexpressing mig-15 in DA9 from wild type (A), plx-1(nc36) (B) and rap-2(gk11) (C). Synaptic domains of DA8 and DA9 are highlighted with green and magenta lines, respectively. Asterisks: DA9 cell body. Arrows: dorsal commissure of DA9. Scale bars: 20μm. (D) Quantification of the DA8 asynaptic domain. (E) Quantification of the DA9 synaptic domain. Each dot represents measurements from a single animal. Blue bars indicate mean ± SEM. n.s.: not significant; ***: p < 0.001. (F) Schematic illustration of synapse distribution in wild type (left), animals overexpressing mig-15 in DA9 (middle) and synaptic tiling mutants overexpressing mig-15 in DA9 (right). Arrows indicate the synaptic tiling border, and colored arrows represent expanded or shortened synaptic domain from DA8 (green) and DA9 (magenta) synaptic domains.

In summary, we demonstrate that synaptic tiling is a mechanism to maintain a uniform distribution of synapses from one class of motor neurons along the nerve cord. Further, our results indicate plx-1 and rap-2 play critical roles in this process by coordinating the position of the synaptic tiling border (Figure 7F).

Discussion

While much is known about the morphogenic triggers for axon guidance and patterned synapse formation, the downstream sequelae of these intracellular effectors has remained unclear. We discovered the role of Rap2 GTPase and TNIK in synapse pattern formation in C. elegans. Since Sema/Plexin signaling processes to inhibit synapse formation are well conserved across species, we propose that Sema/Plexin also utilize Rap2 and TNIK to regulate synapse patterning in mammals as well.

Cell autonomous and non-autonomous functions of Sema/Plexin signaling components

Previously we showed that both smp-1 and plx-1 are necessary and sufficient in DA9, which suggests that smp-1 and plx-1 act cell-autonomously in DA9 and non-autonomously in DA8 to determine the synaptic tiling border. We proposed that Sema/PLX-1 in DA9 send a retrograde signal to DA8 through an unidentified signaling molecule (X) to induce the synaptic tiling pattern in DA8 (Mizumoto and Shen, 2013a). However, we found that both rap-2 and mig-15 act cell autonomously, since our DA9-specific rescue experiment only rescued the DA9 phenotype, but not the DA8 phenotype. This conclusion is further supported since the synaptic tiling defects of these mutants were fully rescued when both neurons express functional rap-2 cDNA or mig-15 genomic DNA. We propose that each neuron utilizes a different set of cell surface proteins but share common intracellular mechanisms to specify synapse patterning. Diverse signaling and cell adhesion molecules, such as atrial natriuretic peptide receptor (NPR) and GPCRs, regulate Rap activity (Birukova et al., 2008; Gloerich and Bos, 2011; Weissman et al., 2004). Screening for these potential Rap regulators should identify novel molecules that interact with Sema/Plexin and act in DA9.

Cycling of Rap GTPase activity in synaptic tiling

We showed that both GDP- and GTP-forms of RAP-2 are required for proper synapse patterning. Considering that PLX-1 regulates the spatial distribution of RAP-2 activity and mig-15 acts downstream of rap-2 in synaptic tiling, RAP-2 may also locally regulate MIG-15(TNIK). While we did not observe a subcellular localization of GFP-MIG-15 in DA9 (data not shown), PLX-1 and RAP-2 may instead regulate MIG-15 activity rather than its spatial localization.

Small GTPase activity is regulated by GAP and GEF (Guanine nucleotide exchange factor) proteins. Yet, we did not observe significant synaptic tiling defects in mutants of putative RAP-2 GEFs, which include pxf-1(RAPGEF2/6) and epac-1(RAPGEF3/4/5) (data not shown) (Frische et al., 2007; Pellis-van Berkel et al., 2005). We speculate that multiple RapGEFs act redundantly to activate RAP-2 in synaptic tiling.

mig-15(TNIK) may integrate multiple inhibitory cues during synapse formation

mig-15 mutants show a greater degree of overlap between DA8 and DA9 synaptic domains than plx-1 or rap-2 mutants. This effect partially occurs from excess synaptogenesis in the posterior asynaptic domain of both DA8 and DA9 neurons. Previously, we demonstrated that Wnt morphogens and their receptors, Frizzled, instruct synaptic topographic patterning by locally inhibiting synapse formation. Indeed, synaptic tiling defects in mig-15 mutants was somewhat similar to the combined effect of plx-1 and wnt mutants (Mizumoto and Shen, 2013b). TNIK can act as a positive regulator of the canonical Wnt signaling pathway in colorectal cancer cells (Mahmoudi et al., 2009). While we do not know whether the canonical Wnt signaling pathway contributes to local inhibition of synapse formation, we propose that TNIK integrates multiple signaling pathways for precise synapse pattern formation.

The exact mechanisms of synaptic actin regulation by TNIK remain undetermined. TNIK could activate JNK kinase pathway (Taira et al., 2004). The MIG-15/JNK-1 signaling pathway inhibits axonal branch formation in sensory neurons in C. elegans (Crawley et al., 2017). In contrast to these well-established role of MIG-15/TNIK as an activator of the JNK pathway, we did not observe any synaptic tiling defect in jnk-1 mutant animals (Figure S5F and S5G). Our result suggests mig-15 does not inhibit synapse formation through the JNK pathway. Further genetic studies of mig-15 in synaptic tiling will elucidate the molecular mechanisms that underlie the role of MIG-15/TNIK in synapse pattern formation.

Plexin signaling and diseases

Aberrant neuronal wiring underlies many neurological disorders. Not surprisingly, Semaphorin and Plexin genes are associated with various neurodevelopmental disorders and intellectual disabilities, including autism spectrum disorders (ASD) and schizophrenia (Mah et al., 2006). For example, PLXNB1, SEMA3A, SEMA4D and SEMA6C are significantly upregulated in the prefrontal cortices of schizophrenic patients (Eastwood et al., 2003). However, non-synonymous variations in the Sema3D gene had a significant protective effect against developing schizophrenia (Fujii et al., 2011). More recent work showed that loss of Sema5A/PlexA2 signaling induces excess excitatory synapse formation in granule cells, which causes ASD-like behavioural defects in mice (Duan et al., 2014).

Similar to Sema/Plexin signaling, TNIK is also associated with various neurological disorders, including schizophrenia and intellectual disabilities (Anazi et al., 2016; Potkin et al., 2010). TNIK can also physically bind and act with DISC1 (Disrupted in Schizophrenia 1) to regulate synaptic composition (Q. Wang et al., 2011). So, we propose that the Sema/Plexin/Rap2/TNIK signaling pathway plays a critical role to precisely define synaptic connections and its disruption may induce serious neurological disorders.

Interestingly, SNPs in Plexin genes are also associated with extremely high IQ. Recent work suggests that loss of PlexinA1 confers better motor control in rodents, due to increased synaptic connectivity in the corticospinal cord (Gu et al., 2017; Spain et al., 2016). Further studies on the Plexin/Rap2/TNIK signaling pathway in synapse map formation, as presented here, will likely reveal the genetic basis of these disorders and conditions.

Methods

Strains

All C. elegans strains were derived from Bristol N2 and raised on OP50 Escherichia coli-seeded nematode growth medium (NGM) plates at 20 C and maintained as described previously (Brenner, 1974). The following mutants were used in this study: unc-104(e1265)II, plx-1(nc36)IV, rap-1(pk2082)IV, rap-3(gk3975)TV, jnk-1(gk7)IV rap-2(gk11)V, rap-2(miz16)V, rap-2(miz17)V, rap-2(miz18)V, rap-2(miz19)V. rap-2(miz20)V, mig-15(rh148)X, mig-15(rh80)X, mig-15(rh326)X.

CRISPR/Cas9 genome editing

rap-2(miz16)V, rap-2(miz17)V, rap-2(miz18)V, rap-2(miz19)V. rap-2(miz20)V were generated using Co-CRISPR method (Kim et al., 2014). unc-22 or dpy-10 co-CRISPR markers were used for selecting candidate animals (Arribere et al., 2014; Kim et al., 2014). Vectors for sgRNA and Cas9 were obtained from Addgene (Plasmid ID: 46169 and 46168, respectively) (Friedland et al., 2013). The rap-2 guide RNA sequence (5’ – gTAGTGGAGGTGTCGGAAAAT-3’) was designed using MIT CRISPR design tool (crispr.mit.edu:8079) and inserted into sgRNA vector using Q5 Site-Directed Mutagenesis kit (NEB). Repair templates with either G12V (miz17 and miz18) and S17A (miz19 and miz20) mutation were generated by PCR with primer sets carrying corresponding mutations (see supplemental Experimental Procesures). Synonymous mutations were also introduced in the sgRNA recognition sequence to avoid Cas9 recruitment to the edited genome. PCR products were cloned into EcoRI site of the pBluescript SK(+) vector. Synthesized double-stranded DNA (GeneArt, Thermo Fisher) was used as a repair template for generating rap-2(miz16) mutant.

Plasmid constructions

C. elegans expression clones were made in a derivative of pPD49.26 (A. Fire), the pSM vector (kind gift from S. McCarroll and C. I. Bargmann). Primer sets used in this study are listed in the Supplemental Experimental Procedures. The following constructs were used and transgenes were generated using standard microinjection method (Mello et al., 1991): wyIs446 (Punc-4::2xGFP-rab-3; Pmig-13::mCherry-rab-3; Podr-1::RFP), wyIs85 (Pitr-1::GFP-rab-3; Podr-1::RFP), wyIs442 (Pflp-13::2xGFP-rab-3; Pplx-2::mCherry-rab-3; Podr-1::RFP), wyIs320 (Pitr-1::plx-1::GFP; Pmig-13::mCherry;;rab-3; Podr-1::GFP), wyIs329 (Pmig-13::GFP-utCH; Pmig-13::mCherry::rab-3; Podr-1::GFP), wyIs524 (Punc-4::2xGFP-rab-3; Pmig-13::mCherry-rab-3; Podr-1::RFP), wyIs685 (Pmig-13::mCherry::rab-3; Pmig-13::GFPnovo2::cla-1; Podr-1::GFP) mizIs1 (Pitr-1::GFPnovo2-CAAX; Pvha-6::zif-1; Pitr-1::mCherry::rab-3; Podr-1::GFP), mizIs19 (Pmig-13::eGFP::hRap2a; Pmig-13::mRFP-RalGDS(RBD)-mRFP; Podr-1::GFP), mizIs33 (Prab-3::mig-15; Podr-1::GFP), jsIs682 (Prab-3::GFP::rab-3; lin-15(+)), wyEx5445 (Prap-1::GFP; Punc-4::myr-mCherry; Podr-1::RFP), wyEx5464 (Prap-2::GFP; Punc-4::myr-mCherry; Podr-1::RFP), mizEx194 (Prap-3::GFP; Pmig-13::myr-mCherry; Podr-1::RFP), mizEx165 (Phlh-1::rap-2; Podr-1::GFP), mizEx164 (Punc-129::rap-2; Podr-1::GFP), mizEx174 (Punc-4c::rap-2; Podr-1::GFP), mizEx157 (Pmig-13::rap-2; Podr-1::GFP), mizEx156 (Punc-4c::hRap2a; Podr-1::GFP), mizEx177 (Punc-4c::rap-1; Podr-1::GFP), mizEx151 (Pmig-13::mig-15; Podr-1::GFP), mizEx147 (Punc-4c::mig-15; Podr-1::GFP), mizEx153 (DpSMmig-15; Podr-1::GFP), mizEx178 (Punc-4c::mig-15(K50A); Podr-1::GFP), mizEx173 (Punc-4::rap-2(G12V); Podr-1::GFP), mizEx179 (Pflp-13::mig-15; Podr-1::GFP), mizEx170 (Pmig-13::mig-15; Podr-1::GFP), mizEx197 (Pmig-13::mig-15; Podr-1::GFP), mizEx210 (Pmig-13::mig-15; Podr-1::RFP), mizEx257 (Prab-3::GFP; Prab-3::mCherry::rab-3, Podr-1::mScarlet::CAAX)

Cloning of rap-1, rap-2 and mig-15

cDNAs of rap-1 and rap-2 were obtained from cDNA library prepared from N2 RNA. Trizol (Invitrogen) was used to purify total RNA from N2, and the SuperScript III First-Strand Synthesis System for RT-PCR (Invitrogen) was used for the reverse-transcription. mig-15 genomic DNA was amplified from the N2 genomic DNA purified using GeneJET Genomic DNA Purification Kit (Thermo Scientific). Phusion (NEB) or Q5 (NEB) DNA polymerases were used for all PCR reactions for amplifying cDNA and genomic DNA fragments. Amplified fragments were cloned into the AscI and KpnI sites of pSM vector using SLiCE method (Motohashi, 2015), Gibson assembly (Gibson et al., 2009) or T4 ligase (NEB). List of primers used in this study is available in the Supplemental Experimental Procedures.

Confocal Microscopy

Images of fluorescently tagged fusion proteins were captured in live C. elegans using a Zeiss LSM800 confocal microscope (Carl Zeiss, Germany). Worms were immobilized on 2% agarose pad using a mixture of 7.5 mM levamisole (Sigma-Aldrich) and 0.225M BDM (2,3-butanedione monoxime) (Sigma-Aldrich). Images were analyzed with Zen software (Carl Zeiss) or Image J (NIH, USA). Definition of each parameter is as follows (Mizumoto and Shen, 2013a): DA8/9 overlap: a distance between the most anterior DA9 synapse and the most posterior DA8 synapse, DA8 asynaptic domain: a distance from commissure to the most posterior DA8 synapse, DA9 synaptic domain: a distance between the most anterior and posterior DA9 synapses. Middle L4 (judged by the stereotyped shape of developing vulva) animals were used for quantification. Averages were taken from at least 20 samples. For GFP::Utrophin-CH, we measured the length from the posterior end of dorsal axon to the anterior end of GFP::Utrophin-CH domain. For each marker strain, the same imaging setting (laser power, gain pinhole) and image processing were used for comparing different genotypes.

Two-photon FLIM experiment

Expression vector for cultured cells (pCI-eGFP-hRap2a, pCI-eGFP-RAP-1, pCI-eGFP-RAP-2, pCI-eGFP-RAP-2(G12V), pCI-eGFP-RAP-2(S17A)) were generated by replacing Ras in pCI-eGFP-Ras (Yasuda et al., 2006) with hRap2a and rap-2 cDNAs with XhoI and BamHI. pCI-mRFP-RalGDS-mRFP plasmid is a kind gift from Dr. Yasuda. Rap2 and FRET sensor plasmids were mixed in 1:2 ratio and transfected into HeLa cells using Lipofectamine 3000 (ThermoFisher). FLIM was conducted 24 hours after transfection. For expression of FLIM markers in the DA9 neuron, each fusion protein constructs were cloned into AscI and KpnI sites of the pSM vector containing mig-13 promoter using SLiCE method.

A custom-made two-photon fluorescence lifetime imaging microscope was used as described elsewhere(Murakoshi et al., 2011). Briefly, EGFP-Rap2a was excited with a Ti-sapphire laser (Mai Tai; Spectra-Physics) tuned to 920 nm. The X and Y scanning galvano mirrors (6210H; Cambridge Technology) were controlled with Scanlmage software(Pologruto et al., 2003). EGFP photon signals were collected an objective lens (60×, 1.0 NA; Olympus) and a photomultiplier tube (H7422-40p; Hamamatsu) placed after a dichroic mirror (FF553-SDi01; Semrock) and emission filter (FF01-625/90; Semrock). A fluorescence lifetime curve was recorded by a time-correlated single-photon-counting board (SPC-150; Becker & Hickl) controlled with custom software. For construction of a fluorescence lifetime image, the mean fluorescence lifetime values (τm) in each pixel were translated into a color-coded image. We quantified free EGFP-Rap2a and EGFP-Rap2a undergoing FRET (binding fraction) as described elsewhere (Yasuda et al., 2006). Briefly, we calculated the proportion of EGFP undergoing FRET in individual ROIs using the following formula: Embedded Image where τfree and τFRET are the fluorescence lifetime of free EGFP and EGFP undergoing FRET, respectively.

Statistics

Prism (GraphPad) software was used for statistical analysis. one-way ANOVA was done and corrected for multiple comparisons with posthoc Tukey’s multiple comparisons tests done between all genotypes. Student’s t-test was used for pairwise comparison. Sample numbers were predetermined before conducting statistical analyses.

Author contribution

KM conceived the project. XC and KM designed and conducted the experiments, collected and interpreted the data. AS, NLW and HM conducted FLIM experiment. AH, MK and EF generated some C. elegans strains and observed phenotypes. MDL conducted co-IP experiment. KM wrote the manuscript with the feedbacks from all authors. KM, HM and BM acquired funding.

Acknowledgements

We are grateful to Donald Moerman and his lab members for generating rap-3 mutant strain, providing other mutant strains, sharing reagents and for general discussions. We also thank Ryohei Yasuda for FRET sensor plasmids, Hiroshi Kawabe for human Rap2a cDNA, Peri Kurshan for sharing unpublished strain, Richard Ikegami for plx-2 promoter construct, Lisa Fernando for the technical support, all Mizumoto lab members and general discussions, Kang Shen, Shaul Yogev and Maulik Patel for comments on the manuscript. Some strains used in this study were obtained from the CGC, which is funded by NIH Office of Research Infrastructure Programs (P40 OD010440) and C. elegans gene knock out consortium. Mizumoto Lab is funded by HFSP (CDA-00004/2014), CIHR (PJT-148667), NSERC (RGPIN-2015-04022), CFI-JELF (34722) and Tomizawa Jun-ichi and Keiko Fund for Young Scientist. KM is a recipient of Canada Research Chair and Michael Smith Foundation for Health Research Scholar.

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Posted December 22, 2017.
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Rap2 and TNIK control Plexin-dependent synaptic tiling in C. elegans
Xi Chen, Akihiro C. E. Shibata, Nicholas L Weilinger, Ardalan Hendi, Mizuki Kurashina, Ethan Fortes, Brian MacVicar, Hideji Murakoshi, Kota Mizumoto
bioRxiv 238352; doi: https://doi.org/10.1101/238352
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