PlexinD1 signaling controls morphological changes and migration termination in newborn neurons
2018; Springer Nature; Volume: 37; Issue: 4 Linguagem: Inglês
10.15252/embj.201797404
ISSN1460-2075
AutoresMasato Sawada, Nobuhiko Ohno, Mitsuyasu Kawaguchi, Shihhui Huang, Takao Hikita, Youmei Sakurai, Huy Bang Nguyen, Truc Quynh Thai, Yuri Ishido, Yutaka Yoshida, Hidehiko Nakagawa, Akiyoshi Uemura, Kazunobu Sawamoto,
Tópico(s)Wnt/β-catenin signaling in development and cancer
ResumoArticle18 January 2018free access Transparent process PlexinD1 signaling controls morphological changes and migration termination in newborn neurons Masato Sawada orcid.org/0000-0002-8694-8526 Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Search for more papers by this author Nobuhiko Ohno Division of Neurobiology and Bioinformatics, National Institute for Physiological Sciences, Okazaki, Japan Department of Anatomy, Division of Histology and Cell Biology, Jichi Medical University, School of Medicine, Shimotsuke, Japan Search for more papers by this author Mitsuyasu Kawaguchi Department of Organic and Medicinal Chemistry, Nagoya City University Graduate School of Pharmaceutical Sciences, Nagoya, Japan Search for more papers by this author Shih-hui Huang Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Search for more papers by this author Takao Hikita Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Search for more papers by this author Youmei Sakurai Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Search for more papers by this author Huy Bang Nguyen Division of Neurobiology and Bioinformatics, National Institute for Physiological Sciences, Okazaki, Japan Search for more papers by this author Truc Quynh Thai Division of Neurobiology and Bioinformatics, National Institute for Physiological Sciences, Okazaki, Japan Search for more papers by this author Yuri Ishido Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Search for more papers by this author Yutaka Yoshida Division of Developmental Biology, Cincinnati Children's Hospital Medical Center, Cincinnati, OH, USA Search for more papers by this author Hidehiko Nakagawa Department of Organic and Medicinal Chemistry, Nagoya City University Graduate School of Pharmaceutical Sciences, Nagoya, Japan Search for more papers by this author Akiyoshi Uemura orcid.org/0000-0001-5574-5470 Department of Retinal Vascular Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Search for more papers by this author Kazunobu Sawamoto Corresponding Author [email protected] orcid.org/0000-0003-1984-5129 Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Division of Neural Development and Regeneration, National Institute for Physiological Sciences, Okazaki, Japan Search for more papers by this author Masato Sawada orcid.org/0000-0002-8694-8526 Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Search for more papers by this author Nobuhiko Ohno Division of Neurobiology and Bioinformatics, National Institute for Physiological Sciences, Okazaki, Japan Department of Anatomy, Division of Histology and Cell Biology, Jichi Medical University, School of Medicine, Shimotsuke, Japan Search for more papers by this author Mitsuyasu Kawaguchi Department of Organic and Medicinal Chemistry, Nagoya City University Graduate School of Pharmaceutical Sciences, Nagoya, Japan Search for more papers by this author Shih-hui Huang Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Search for more papers by this author Takao Hikita Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Search for more papers by this author Youmei Sakurai Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Search for more papers by this author Huy Bang Nguyen Division of Neurobiology and Bioinformatics, National Institute for Physiological Sciences, Okazaki, Japan Search for more papers by this author Truc Quynh Thai Division of Neurobiology and Bioinformatics, National Institute for Physiological Sciences, Okazaki, Japan Search for more papers by this author Yuri Ishido Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Search for more papers by this author Yutaka Yoshida Division of Developmental Biology, Cincinnati Children's Hospital Medical Center, Cincinnati, OH, USA Search for more papers by this author Hidehiko Nakagawa Department of Organic and Medicinal Chemistry, Nagoya City University Graduate School of Pharmaceutical Sciences, Nagoya, Japan Search for more papers by this author Akiyoshi Uemura orcid.org/0000-0001-5574-5470 Department of Retinal Vascular Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Search for more papers by this author Kazunobu Sawamoto Corresponding Author [email protected] orcid.org/0000-0003-1984-5129 Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan Division of Neural Development and Regeneration, National Institute for Physiological Sciences, Okazaki, Japan Search for more papers by this author Author Information Masato Sawada1, Nobuhiko Ohno2,3, Mitsuyasu Kawaguchi4, Shih-hui Huang1, Takao Hikita1, Youmei Sakurai1, Huy Bang Nguyen2, Truc Quynh Thai2, Yuri Ishido1, Yutaka Yoshida5, Hidehiko Nakagawa4, Akiyoshi Uemura6 and Kazunobu Sawamoto *,1,7 1Department of Developmental and Regenerative Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan 2Division of Neurobiology and Bioinformatics, National Institute for Physiological Sciences, Okazaki, Japan 3Department of Anatomy, Division of Histology and Cell Biology, Jichi Medical University, School of Medicine, Shimotsuke, Japan 4Department of Organic and Medicinal Chemistry, Nagoya City University Graduate School of Pharmaceutical Sciences, Nagoya, Japan 5Division of Developmental Biology, Cincinnati Children's Hospital Medical Center, Cincinnati, OH, USA 6Department of Retinal Vascular Biology, Nagoya City University Graduate School of Medical Sciences, Nagoya, Japan 7Division of Neural Development and Regeneration, National Institute for Physiological Sciences, Okazaki, Japan *Corresponding author. Tel: +81 52 853 8532; Fax: +81 52 851 1898; E-mail: [email protected] EMBO J (2018)37:e97404https://doi.org/10.15252/embj.201797404 PDFDownload PDF of article text and main figures. Peer ReviewDownload a summary of the editorial decision process including editorial decision letters, reviewer comments and author responses to feedback. ToolsAdd to favoritesDownload CitationsTrack CitationsPermissions ShareFacebookTwitterLinked InMendeleyWechatReddit Figures & Info Abstract Newborn neurons maintain a very simple, bipolar shape, while they migrate from their birthplace toward their destinations in the brain, where they differentiate into mature neurons with complex dendritic morphologies. Here, we report a mechanism by which the termination of neuronal migration is maintained in the postnatal olfactory bulb (OB). During neuronal deceleration in the OB, newborn neurons transiently extend a protrusion from the proximal part of their leading process in the resting phase, which we refer to as a filopodium-like lateral protrusion (FLP). The FLP formation is induced by PlexinD1 downregulation and local Rac1 activation, which coincide with microtubule reorganization and the pausing of somal translocation. The somal translocation of resting neurons is suppressed by microtubule polymerization within the FLP. The timing of neuronal migration termination, controlled by Sema3E-PlexinD1-Rac1 signaling, influences the final positioning, dendritic patterns, and functions of the neurons in the OB. These results suggest that PlexinD1 signaling controls FLP formation and the termination of neuronal migration through a precise control of microtubule dynamics. Synopsis During termination of neuronal migration, newborn neurons are found to transiently extend a protrusion termed filopodium-like lateral protrusion (FLP). PlexinD1 signaling controls the FLP formation and migration termination of newborn neurons through regulation of microtubule dynamics. Migrating neurons transiently extend an FLP during neuronal deceleration. The FLP formation is induced by PlexinD1 downregulation and local Rac1 activation. Microtubule polymerization within the FLP suppresses the somal translocation of newborn neurons. PlexinD1 signaling determines the final positioning, dendritic patterns, and function of new neurons in the olfactory bulb. Introduction Precise control of the maintenance and termination of neuronal migration is required for proper brain development and function. In the mammalian brain, newly born neurons migrate from the germinal zone toward their destinations (Cooper, 2013; Marin, 2013). During migration, the new neurons maintain an immature morphology and move in a saltatory manner, executed by repeated extension of the leading process and subsequent somal translocation (Bellion et al, 2005; Schaar & McConnell, 2005). The termination of new-neuron migration occurs when they arrive their final destinations, and is regulated by intrinsic and extrinsic mechanisms (Gongidi et al, 2004; Bortone & Polleux, 2009; Simo & Cooper, 2013; Ota et al, 2014). In addition, prominent changes in cell morphology, involving the extension of cellular protrusions, are observed during migration termination (Nadarajah et al, 2001) and lead to the development of dendrites and functional neuronal circuits. However, how the morphological changes that occur during neuronal migration termination are regulated is unclear. The migration of olfactory bulb (OB) interneurons, which are continuously generated from neural stem cells (NSCs) in the postnatal ventricular–subventricular zone (V-SVZ; Doetsch et al, 1999), provides an excellent model to study the morphological changes that accompany neuronal migration termination (Ghashghaei et al, 2007; Sawada et al, 2011). These new neurons tangentially migrate toward the OB core through the rostral migratory stream (RMS; Lois et al, 1996; Anton et al, 2004; Sawamoto et al, 2006; Snapyan et al, 2009; Kaneko et al, 2010; Garcia-Gonzalez et al, 2017) and then turn radially within the OB layers (Hack et al, 2002; Saghatelyan et al, 2004; Ng et al, 2005; Garcia-Gonzalez et al, 2017; Petri et al, 2017). In the OB, each new neuron terminates its migration at a specific depth in the OB layers: granule cells (GCs) travel to various depths of the granule cell layer (GCL), whereas periglomerular cells (PGCs) travel farther, to the glomerular layer (GL; Luskin, 1993; Lois & Alvarez-Buylla, 1994). The distinct dendritic arborization and synaptic connection patterns of these new neurons depend on their final positioning in the OB (Ota et al, 2014). Therefore, the final positioning of new neurons in the OB, determined by the regulation of migration termination, is critical for the development and maintenance of functional OB circuits. Here, we report a mechanism by which the termination of new-neuron migration is maintained in the postnatal OB. When the new neurons approach their destination, they transiently extend a protrusion from the proximal part of the leading process in the resting phase of migration, which we refer to as a filopodium-like lateral protrusion (FLP). Local Rac1 activation, the timing of which is controlled by the downregulation of repulsive Sema3E-PlexinD1 signaling, induces FLP formation. Microtubule (MT) polymerization within the FLP suppresses the somal translocation of new neurons, thereby maintaining the resting phase. The regulatory mechanism involving FLP formation links neuronal migration termination and the initiation of differentiation, contributing to the positioning and functions of new neurons in the postnatal OB. Results The FLP is formed in the resting phase of migration during neuronal deceleration To study the morphological changes of new neurons during the termination process of migration, we observed them by live imaging in cultured OB slices. As described previously, actively migrating new neurons showed repeated extension of the leading process followed by forward movement of the soma (Ota et al, 2014; Fig 1A; Movie EV1; Appendix Fig S1A). These new neurons occasionally formed protrusions that branched laterally from the leading process. Because the length of these protrusions showed a bimodal distribution (Fig 1B), we defined the type with longer protrusions (> 11 μm), which was distinct from the shorter type, as "filopodium-like lateral protrusions (FLPs)". FLP formation was observed only in the proximal leading process (Fig 1C). Furthermore, the frequency of FLP formation was significantly higher in the resting phase than in the migratory phase, while non-FLP-protrusions did not show an association with migration phase (Fig 1D). In addition, the FLPs were observed only in the late stage of radial migration, and not in the resting phase of actively migrating neurons in the RMS or in neurons in the initial stage of radial migration in the OB (Fig 1A, 0–60 min; Appendix Fig S1B). The duration of the resting phase of neurons with FLPs was significantly longer than that of neurons without FLPs (Fig 1E). To further investigate the timing of FLP formation, we performed time-lapse imaging of cultured new neurons expressing EGFP fused to the calponin homology domain of utrophin (EGFP-UtrCH), a fluorescent reporter of F-actin (Burkel et al, 2007). The transient extension of the FLP, labeled with a strong EGFP-UtrCH signal, was observed after the end of leading-process extension and before the beginning of somal translocation in cultured new neurons (Fig 1F and G; Movie EV2). FLPs were also observed in resting neurons generated in the embryonic lateral, medial, and caudal ganglionic eminences (Appendix Fig S2). Together, these results suggest that FLPs represent a distinct type of protrusion with unique characteristics in terms of length, position, and timing of formation in migrating new neurons. Figure 1. Characterization of the FLP in migrating neurons A. Time-lapse images of a tdTomato-labeled GC migrating in an OB slice. B. Distribution of protrusion lengths (229 events, 24 cells). Gray and black lines indicate fitted curves for short and long protrusions, respectively. FLPs, green. C. Cellular location of protrusions (24 cells). D. Protrusion formation in the resting and migratory phases (24 cells, ***P < 0.005, paired t-test). E. Duration of the resting phase (24 cells, *P < 0.05, paired t-test). F, G. Time-lapse images of a cultured migrating neuron expressing EGFP-UtrCH (green) and DsRed (red) (F) and FLP frequency (G) (15 cells, **P < 0.01, one-way ANOVA followed by Tukey–Kramer test). H, H'. Representative image of an FLP-bearing cultured neuron stained with phalloidin (green) and an α-tubulin (red) antibody. Magnified images of the boxed area in (H) are shown in (H'). I, I'. Representative images of a cultured neuron with or without an FLP, stained with anti-tyrosinated (Tyr, green) and anti-acetylated (Ac, magenta) tubulin antibodies. Magnified images of the boxed areas in (I) are shown in (I'). White arrowheads, loosened tubulin bundles. J. Classification of EB3-GFP+ (a marker of MT plus-ends) trajectories. K. EB3-GFP+ trajectories (white and orange arrows). Orange arrows indicate "traversing" trajectories classified in (J). Magnified image of the boxed area in (K) is shown in (K'). Green arrowheads indicate swelling. L, M. Proportion of EB3-GFP+ trajectory directions in the distal (L) and proximal (M) leading process. ***P < 0.005 [vs. FLP(−)], one-way ANOVA followed by Tukey–Kramer test. Parentheses, number of cells. N. EB3-GFP speed. *P < 0.05, **P < 0.01, ***P < 0.005 (vs. FLP(−) [no swelling]), §P < 0.05, §§§P < 0.005 (vs. FLP(−) [swelling]), ##P < 0.0005 (vs. FLP(+)-Distal LP), one-way ANOVA followed by Tukey–Kramer test. Parentheses, number of events. O, P. Transmission electron microscopy images of the proximal leading process of a migrating neuron without (O) or with (P) an FLP. Red, blue, and green indicate cell membrane, microtubules, and centrioles, respectively. Q–R''. SBF-SEM images (Q) and their 3D reconstruction (R–R'') of an FLP-bearing neuron (green). Arrows, mitochondria (Q). Red indicates the region of contact between the FLP and GCs (R''). Data information: Yellow arrows and red arrowheads indicate FLPs and non-FLP protrusions, respectively (A, F, H, I, K, and P). Scale bars: 10 μm (A, F, H), 5 μm (I, K), 1 μm (O–Q). Bars indicate mean ± SEM. Download figure Download PowerPoint To examine the cytoskeletal components of the FLPs, we analyzed the F-actin and MT content in cultured V-SVZ-derived new neurons. Whereas the shorter protrusions contained F-actin but lacked α-tubulin-positive structures (Fig 1H and H'), a common feature of filopodia (Dent et al, 2007), the FLPs contained both F-actin and MTs. To further investigate the MT network in the FLP-bearing neurons, we used anti-tyrosinated- and anti-acetylated-tubulin antibodies, which label dynamic and stable MTs, respectively (Umeshima et al, 2007). The FLPs predominantly contained tyrosinated tubulin, indicating that they are transient structures (Fig 1I). Furthermore, MT bundles were locally loosened in the proximal leading process and decreased in the perinuclear MT cage in the FLP-bearing neurons, compared with those in neurons without FLP (Fig 1I and I'). Consistent with these observations, MT plus-end imaging using EB3-GFP (Watanabe et al, 2015) revealed that MT polymerization traversed sideways across the longitudinal axis of the proximal leading process in neurons with FLPs but not in those without FLPs (Fig 1J–M; Movie EV3). Furthermore, the overall MT polymerization levels in the proximal leading process and soma of neurons with FLPs were significantly lower than in those regions of neurons without FLPs (Fig 1N; Movie EV3). On the other hand, active MT polymerization was observed in the FLP, at a level comparable to that in the distal leading process, which showed the highest MT polymerization activity in the neuron (Fig 1N). The traversing MTs in the proximal leading process were also observed in FLP-bearing neurons in vivo (Fig 1O and P; Appendix Fig S3). Collectively, these observations indicated that FLP formation always coincides with local cytoskeletal reorganization and decelerated migration. To characterize the FLP structure in detail, we performed three-dimensional ultrastructural analyses using serial block-face scanning electron microscopy (SBF-SEM). Unlike mature dendrites, the FLP contained no or very few mitochondria and made contact with the GC soma (Fig 1Q–R''; Movie EV4; Appendix Fig S4). The formed FLP extended further and developed into a stable structure (Fig 1A, 310–530 min). Furthermore, some of the mature GCs (18.8 ± 3.4%) had a dendrite with spine-like structures in the FLP-corresponding region of their primary dendrite (Appendix Fig S1C). These results suggested that the FLPs can eventually develop into a lateral dendrite after migration termination (Appendix Fig S1D). To investigate the FLP formation in new neurons that terminate their migration at different regions of the OB, we compared the migration of GCs and PGCs using doublecortin (Dcx)-GFP mice. We dissected different regions of the OB of these mice that contained migrating GC- or PGC-rich cell fractions and transplanted them into the V-SVZ of wild-type (WT) mice (Fig 2A). At 8 days post-transplantation (dpt), both GC-rich and PGC-rich cell fractions had migrated to the OB and exhibited FLPs (Fig 2B and C). In the GCL, the PGC-rich fraction contained a higher proportion of cells exhibiting a leading process and cytoplasmic swelling, typical morphological features of migratory neurons (Schaar & McConnell, 2005), than did the GC-rich fraction (Fig 2C). Notably, the proportion of FLP-bearing neurons in the GC-rich fraction was significantly higher than in the PGC-rich one (Fig 2C; GC-rich fraction, 82.8 ± 2.9%; PGC-rich fraction, 63.3 ± 2.4%; P = 0.0066, unpaired t-test). In addition, in the external plexiform layer (EPL) and GL, the proportion of FLP-bearing neurons in the migratory PGC-rich-fraction cells at 14 dpt was higher than that at 8 dpt (Fig 2D; 8 dpt, 72.1 ± 5.8%; 14 dpt, 92.4 ± 1.6%; P = 0.0014, unpaired t-test). These results suggested that neurons in both the GC- and the PGC-rich fractions form FLPs during neuronal deceleration, the timing of which is earlier in the GC-rich fraction than in the PGC-rich one. At 14 dpt, a higher percentage of the PGC-rich than the GC-rich fraction had reached the GL (Fig 2E and F). Together, these findings suggest that transplanted GCs and PGCs both have the potential to reach their targeted destinations, and that FLP formation might be related to the process of migration termination in GCs and PGCs. Figure 2. Neurons from GC- and PGC-rich fractions have the potential to reach their targeted destinations and form FLPs during neuronal deceleration Experimental scheme. Typical morphology of a transplanted cell with an FLP (arrow) observed in the GCL 8 days post-transplantation (dpt). Proportion of swelling-bearing cells with a leading process with or without an FLP from GC-rich (three mice) and PGC-rich (three mice) fractions of the GCL at 8 dpt. Proportion of swelling-bearing cells with a leading process with or without an FLP from PGC-rich fractions of the EPL and GL at 8 (three mice) and 14 (seven mice) dpt. Anti-GFP (green) and Hoechst 33342 (blue, nuclei) staining of OB sections at 14 dpt. Proportion of neurons in the OB layers from GC-rich (seven mice) and PGC-rich (seven mice) fractions at 14 dpt. Data information: GL, glomerular layer; EPL, external plexiform layer; MCL, mitral cell layer; IPL, inner plexiform layer; s/m/dGCL, superficial/middle/deep granule cell layer. Scale bars: 10 μm (B), 50 μm (E). *P < 0.05, **P < 0.01, ***P < 0.005 (unpaired t-test). Bars indicate mean ± SEM. Download figure Download PowerPoint Sema3E-PlexinD1-mediated inhibition of Rac1 activation suppresses the FLP formation in migrating neurons In search of transmembrane receptors involved in FLP formation, we found that PlexinD1 (Gu et al, 2005; Chauvet et al, 2007) was expressed specifically in the leading process of Dcx-positive (Dcx+) new neurons with a migratory morphology in the OB (Fig 3A; Appendix Fig S5A–J). To compare the expression levels of PlexinD1 in migrating GCs and PGCs, we quantified the PlexinD1 intensity in transplanted cells and found that it was higher in cells from the PGC-rich fraction than in cells from the GC-rich one (Appendix Fig S5C). Consistent with this finding, the PlexinD1 expression level in Dcx+ cells was higher in the EPL (which contains migrating PGCs) than in the GCL (which contains differentiating GCs and migrating GCs/PGCs; Fig 3A and B). Together, these results suggested that new neurons that migrate longer distances have higher levels of PlexinD1. Figure 3. Expression of Sema3E and PlexinD1 in the OB Coronal section of the OB in WT mice stained for PlexinD1 (red) and Dcx (green). Relative expression level of PlexinD1 in Dcx+ cells (GCL, n = 138 cells from three mice; EPL, n = 36 cells from three mice; ***P < 0.005, Mann–Whitney U-test). Bars indicate mean ± SEM. Coronal sections of the OB in Sema3E+/GFP mice stained for GFP (green), NeuN (red), and Dcx (blue). Data information: GL, glomerular layer; EPL, external plexiform layer; MCL, mitral cell layer; IPL, inner plexiform layer; GCL, granule cell layer. Scale bars: 10 μm (A), 50 μm (C). Download figure Download PowerPoint To examine the relationship between the subcellular distribution of PlexinD1 and FLP formation, we used a PlexinD1-GFP fusion protein, which recapitulated the distribution of endogenous PlexinD1 in the leading process of migrating new neurons (Appendix Fig S5A–O). Time-lapse imaging of cultured new neurons expressing PlexinD1-GFP revealed that PlexinD1 was locally downregulated at the membrane of the proximal leading process prior to FLP formation (Appendix Fig S5P–Q), suggesting that PlexinD1 suppresses FLP formation. Consistent with this observation, persistent PlexinD1 overexpression significantly decreased the FLP-formation frequency and increased the migration speed of new neurons in the GCL (Fig 4A and B; Movie EV1). Taken together, these results suggest that PlexinD1 suppresses FLP formation and promotes new-neuron migration. Figure 4. Sema3E-PlexinD1-mediated inhibition of Rac1 activation suppresses FLP formation in migrating neurons A. Time-lapse images of a tdTomato-labeled PlexinD1-overexpressing neuron migrating in the OB. B. Effect of PlexinD1 overexpression on the FLP-formation frequency and neuronal migration speed in the GCL. FLP formation, n = 78 (Control), 45 (PlexinD1) cells, ***P < 0.005, unpaired t-test; Speed, n = 114 (Control), 58 (PlexinD1) cells, ***P < 0.005, Mann–Whitney U-test. C. Time-lapse images of DsRed-labeled control and PlexinD1-KD neurons. D. Effect of Sema3E on leading-process length and FLP-formation frequency (n = 37, 37, 45, 40 cells; ***P < 0.005, unpaired t-test). E. Effect of Sema3E on neuronal migration in vitro (n = 42, 142, 44, 114 cells; *P < 0.05, unpaired t-test). F. Representative images of Dcx+ neurons (red) expressing EGFP (control) or EGFP-R-RasCA (green). G. Number of FLPs in control and R-RasCA-expressing neurons (n = 319, 345, 178, 231 cells, *P < 0.05, ***P < 0.005, Mann–Whitney U-test). H, H'. Time-lapse FRET ratiometric images of Rac1 activity (pseudocolors) in a cultured migrating neuron. Magnified images of the boxed areas in (H) are shown in (H'). I. Activation of Rho GTPases at the leading process before FLP formation [Rac1 (56 events, 10 cells), cdc42 (32 events, 11 cells), RhoA (27 events, 11 cells); ***P < 0.005, ****P < 0.0005, paired t-test (vs. baseline)]. J. Correlation between Rac1 activation and FLP-formation frequency (10 cells, *P < 0.05, Kruskal–Wallis test followed by Steel–Dwass test). K. Rac1 activation in the disappearing and stable FLP [disappearing (16 events, 10 cells); stable (54 events, 10 cells); *P < 0.05, ****P < 0.0005, paired t-test (vs. baseline); disappearing vs. stable, ***P < 0.005, unpaired t-test]. L. Effect of Sema3E on Rac1 activation at the leading tip or shaft in control (10 cells) and PlexinD1-KD (11 cells) neurons [Sema3E(−) vs. (+), ***P < 0.005, paired t-test; Control vs. PlexinD1-KD in Sema3E(+), ***P < 0.005, unpaired t-test]. M. Time-lapse FRET ratiometric images of Rac1 activity in a neuron migrating in a cultured OB slice. N. Rac1 activation before (pre) and during (FLP) FLP formation [seven events, six cells; *P < 0.05, paired t-test (vs. baseline)]. Data information: Numbers indicate minutes after the first imaging frame (A, H, M) or Sema3E addition (C). Arrows, FLPs. Red arrowheads, Rac1 activation. Scale bars: 10 μm. Bars indicate mean ± SEM. Download figure Download PowerPoint We next investigated the expression pattern of Sema3E, a ligand for PlexinD1 (Gu et al, 2005; Fig 3C; Appendix Fig S6). Sema3E expression was observed in NeuN+ mature GCs and PGCs but not in Dcx+ new neurons in the OB (Fig 3C; Appendix Fig S6A–E and H). Reelin+ mature projection neurons (mitral and tufted cells) also did not express Sema3E (Appendix Fig S6F). In addition, Sema3e mRNA localized to the GL, MCL, and GCL in the OB (Appendix Fig S6G). Together, these results suggested that neurons secrete Sema3E after they stop migrating and become mature olfactory interneurons. Sema3E protein significantly suppressed FLP formation and promoted cultured new-neuron migration without affecting the leading-process length (Fig 4C–E). Expression of a constitutively active form of R-Ras (R-RasCA), which is resistant to PlexinD1's GTPase-activating protein (GAP) activity (Uesugi et al, 2009), antagonized the Sema3E-mediated suppression of FLP formation (Fig 4F and G). These results suggested that Sema3E-PlexinD1 signaling suppresses FLP formation and maintains the migratory potential of new neurons. To investigate how Sema3E-PlexinD1 signaling suppresses FLP formation, we analyzed the activation of Rho GTPases, which function downstream of R-Ras, using FRET imaging (Yoshizaki et al, 2003; Ota et al, 2014; Fig 4H–N). The Rac1 activity was increased at the leading process before FLP formation and was sustained within the FLP during FLP formation, in both dissociated neurons (Fig 4H–K) and brain slices (Fig 4M and N). Sema3E significantly suppressed Rac1 activation at the shaft, but not the tip of the leading process in a PlexinD1-dependent manner (Fig 4L). Taken together, these results suggest that Sema3E-PlexinD1 signaling suppresses FLP formation via Rac1 inhibition. Microtubule polymerization within the FLP suppresses the somal translocation of resting neurons To study the role of Rac1 activity in FLP formation and in migration termination, we introduced a photoactivatable (PA)-Rac1 (Wu et al, 2009) into cultured new neurons. Rac1 inactivation in the FLP by a dominant-negative form of PA-Rac1 (PA-Rac1DN) caused rapid FLP retraction, suggesting that Rac1 activity is required for maintaining the FLP (Fig 5A and B; Movie EV5). Furthermore, this FLP retraction was followed by somal translocation, leading to an increased neuronal migration speed (Fig 5A and C; Movie EV5). We also studied
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