Skip to main content
NCBI home page
The EMBO Journal logoLink to The EMBO Journal
. 2018 Jan 18;37(4):e97404. doi: 10.15252/embj.201797404

PlexinD1 signaling controls morphological changes and migration termination in newborn neurons

Masato Sawada 1, Nobuhiko Ohno 2,3, Mitsuyasu Kawaguchi 4, Shih‐hui Huang 1, Takao Hikita 1, Youmei Sakurai 1, Huy Bang Nguyen 2, Truc Quynh Thai 2, Yuri Ishido 1, Yutaka Yoshida 5, Hidehiko Nakagawa 4, Akiyoshi Uemura 6, Kazunobu Sawamoto 1,7,✉
PMCID: PMC5813262  PMID: 29348324

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.

Keywords: microtubule, neuronal migration, olfactory bulb, photoactivation, postnatal neurogenesis

Subject Categories: Cell Adhesion, Polarity & Cytoskeleton; Neuroscience

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.

Figure 1

Open in a new tab
  • 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.

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.

Figure 2

Open in a new tab
  1. Experimental scheme.
  2. Typical morphology of a transplanted cell with an FLP (arrow) observed in the GCL 8 days post‐transplantation (dpt).
  3. 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.
  4. 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.
  5. Anti‐GFP (green) and Hoechst 33342 (blue, nuclei) staining of OB sections at 14 dpt.
  6. 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.

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 .

Figure 3

Open in a new tab
  1. Coronal section of the OB in WT mice stained for PlexinD1 (red) and Dcx (green).
  2. 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.
  3. 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).

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.

Figure 4

Open in a new tab
  • 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.

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 the role of Rac1 using a constitutively active form of PA‐Rac1 (PA‐Rac1CA). Transient Rac1 photoactivation using PA‐Rac1CA, but not C450A (a light‐insensitive mutant), induced FLP formation (Fig 5D and E), indicating that local Rac1 activation in the proximal reading process is sufficient for FLP formation. In addition, the neuronal migration speed was significantly decreased during FLP formation (Fig 5D and E; Movie EV5). Rac1 activation affects various biological responses, such as the oxidative response, in addition to cytoskeletal reorganization (Bedard & Krause, 2007). To investigate the downstream factors of activated Rac1, we tested the effects of Rac1‐effector knockdown (KD) on FLP formation and migration termination after PA‐Rac1CA‐induced Rac1 activation (Fig 5F–H; Appendix Fig S7). We first examined the role of p67phox, a component of NADPH oxidase, which produces reactive oxygen species (ROS) downstream of activated Rac1 (Koga et al, 1999; Lapouge et al, 2000; Dang et al, 2001). The KD of p67phox caused a decrease in ROS production in migrating new neurons (Appendix Fig S7), but did not affect the Rac1‐induced FLP formation or maintenance of migration termination (Fig 5G and H). We next examined IQGAP1, which reorganizes F‐actin and MT cytoskeletons in response to Rac1 activation (Fukata et al, 2002; Watanabe et al, 2004). IQGAP1‐KD prevented the PA‐Rac1CA‐induced FLP formation and maintenance of migration termination (Fig 5F and H; Appendix Fig S7). These results suggested that the Rac1‐induced FLP formation and maintenance of migration termination are controlled by IQGAP1‐mediated cytoskeletal reorganization.

Figure 5. FLP formation and suppression of somal translocation are mediated by IQGAP1 in new neurons.

Figure 5

Open in a new tab
  • A
    Time‐lapse images of PA‐Rac1DN‐associated FLP retraction. Magnified images of the FLP are shown in (A').
  • B
    FLP length following photoactivation.
  • C
    Migrating neuron speed (nine cells, ***P < 0.005, paired t‐test).
  • D
    Time‐lapse images of PA‐Rac1CA‐induced FLP formation. Magnified images of the FLP are shown in (D').
  • E
    Frequency of PA‐induced FLP formation [n = 3 (28 cells, C450A), 5 (43 cells, PA‐Rac1CA) cultures, ***P < 0.005, unpaired t‐test], and migrating neuron speed (five cells, ***P < 0.005, paired t‐test).
  • F, G
    Time‐lapse images of PA‐Rac1CA‐induced FLP formation in IQGAP1‐ (F) and p67phox‐ (G) KD neurons.
  • H
    Frequency of PA‐induced FLP formation [n = 3 (Control, 22 cells), 4 (IQGAP1‐KD, 39 cells), 4 (p67phox‐KD, 42 cells) cultures; **P < 0.01, one‐way ANOVA followed by Dunnett test].
  • I
    Trajectories (white and orange arrows) of EB3‐mCherry+ MT plus‐ends in a PA‐Rac1DN‐expressing cell. Orange arrows indicate “traversing” trajectories classified in (J). Dotted lines indicate cell membrane (I). Nuc, nucleus.
  • J, K
    Proportion of EB3‐mCherry+ trajectory directions in the proximal leading process (J; 5 cells; *P < 0.05, **P < 0.01, paired t‐test) and speed of EB3‐mCherry+ MT plus‐ends (K; 5 cells; proximal LP, *P < 0.05, Mann–Whitney U‐test; soma, ***P < 0.005, unpaired t‐test). Parentheses, number of events (K).
Data information: Numbers indicate minutes (A, D, F, G) or seconds (I) after photostimulation. Yellow arrows indicate FLPs. Circled area was photostimulated. Dotted lines indicate the cell rear (A, D, F, G). Scale bars: 10 μm. Bars indicate mean ± SEM.

Considering the active MT polymerization observed in the FLP (Fig 1N), and the promotion of directional MT polymerization in the proximal leading process and somal translocation following FLP retraction (Fig 5I–K), we hypothesized that MT polymerization in the FLP contributes to the maintenance of migration termination. To test this possibility, we used a photoswitchable inhibitor of MT polymerization, photostatin‐1 (PST‐1), which is activated and inactivated by violet (405 nm) and green (514 nm) laser illumination, respectively (Borowiak et al, 2015; Appendix Fig S8A and B). We illuminated the FLP with a violet laser to inhibit MT polymerization and illuminated the proximal leading process with a green laser to suppress activated PST‐1 that might diffuse from the FLP. This illumination procedure inhibited the MT polymerization in the FLP without affecting the FLP structure and caused a reinitiation of somal translocation (Fig 6A–D). These results suggested that MT polymerization in the FLP suppresses the somal translocation of new neurons, thereby maintaining the resting phase.

Figure 6. MT polymerization in the FLP suppresses the somal translocation of resting neurons.

Figure 6

Open in a new tab
  • A, B
    Time‐lapse images of the inhibition of MT polymerization in the FLP by PST‐1. Circled areas were photostimulated using a 405‐nm (activating) (A) and 514‐nm (deactivating) (B) laser. Rectangular areas were photostimulated using a 514‐nm laser. Magnified images of the FLP in (A) are shown in (A'). (A') Disappearance of EB3‐GFP+ signals (green arrowheads) by PST‐1 activation in the FLP.
  • C
    Density of EB3‐GFP+ dots (MT plus‐ends) in the proximal leading process and FLP (four cells; *P < 0.05, paired t‐test).
  • D
    Migrating neuron speed [No laser vs. Photoswitching in 405 nm, *P < 0.05, paired t‐test; 405 nm (five cells) vs. 514 nm (six cells) in Photoswitching, *P < 0.05, unpaired t‐test].
  • E, F
    Time‐lapse images of the inhibition of somal translocation by PST‐1 after FLP retraction. Circled areas were photostimulated using a 405‐ (E) and 514‐ (F) nm laser.
  • G
    Migrating neuron speed [405 nm (six cells) vs. 514 nm (five cells) in Photoswitching, *P < 0.05, unpaired t‐test; No laser vs. Photoswitching in 514 nm, *P < 0.05, paired t‐test].
  • H
    Model.
Data information: Numbers indicate seconds (A, B, E, and F) before and after photostimulation. Yellow arrows indicate FLPs. Dotted lines indicate the cell rear. Scale bars: 10 μm. Bars indicate mean ± SEM.

To investigate whether the somal translocation occurring after FLP retraction requires directional MT polymerization in the proximal leading process, we illuminated the proximal leading process of PST‐1‐treated neurons with a violet or green laser after FLP retraction. We found that the somal movement after FLP retraction was slower in the presence of activated PST‐1 (Fig 6E–G; Appendix Fig S8C and D), suggesting that directional MT polymerization is required for somal translocation. Taken together, these results suggest that MT polymerization within the FLP, whose formation is regulated by Rac1 and IQGAP1, suppresses the somal translocation of resting neurons (Fig 6H).

Sema3E‐PlexinD1 signaling determines final positioning, dendritic patterns, and function of new neurons in the OB

To study whether the morphological regulation of migrating neurons by Sema3E‐PlexinD1 signaling affects their final positioning in the OB, we injected lentiviral vectors expressing either WT PlexinD1 or PlexinD1 mutants containing defective GAP domains (Worzfeld et al, 2014) into the V‐SVZ and analyzed the infected neuron distribution in the OB 10 days later (Fig 7A). The migration of neurons toward the GL was increased by overexpressing WT PlexinD1 (Fig 7B; Appendix Fig S9A–C), but not the PlexinD1 GAP domain mutants (Fig 7B), suggesting that PlexinD1's GAP activity is involved in promoting new‐neuron migration and determining their final positions in the OB. Next, we analyzed Sema3E KO and PlexinD1‐cKO mice, in which Cre‐expressing lentivirus was injected into the V‐SVZ of PlexinD1 fl/fl mice. The neuron migration toward the GL was inhibited in both Sema3E KO (Fig 7C) and PlexinD1‐cKO (Fig 7D) mice. To analyze whether the migration of the transplanted PGC‐rich cell fraction toward the GL (Fig 2E and F) was also regulated by PlexinD1, the migrating PGC‐rich cell fraction was infected with lentivirus expressing PlexinD1‐shRNA in vitro and transplanted into the V‐SVZ of WT mice. We found that PlexinD1‐KD inhibited migration of the PGC‐rich cell fraction toward the GL (Appendix Fig S9D–L). The PlexinD1 deficiency had no effect on neurogenesis or fate specification in the V‐SVZ, neuronal migration in the RMS, or interneuron identity (Appendix Fig S10). Collectively, these results suggested that Sema3E‐PlexinD1 signaling is essential for proper neuronal positioning in the OB.

Figure 7. Sema3E‐PlexinD1 signaling determines final positioning, dendritic patterns, and function of new neurons in the OB .

Figure 7

Open in a new tab
  • A
    Schematic illustration of new‐neuron positioning and dendritic patterns in the OB.
  • B–D
    Proportion of neurons in the OB layers in control (five mice), PlexinD1 (five mice), PlexinD1GAP1RA (six mice), and PlexinD1∆RBD (five mice) groups (B; one‐way ANOVA followed by Tukey–Kramer test), control (seven mice) and Sema3E KO (six mice) mice (C; unpaired t‐test), and control (six mice) and PlexinD1‐cKO (six mice) mice (D; unpaired t‐test) at 10 days postinfection (dpi). *P < 0.05, **P < 0.01, ***P < 0.005.
  • E
    Traces of neuronal morphologies in control and PlexinD1‐cKO mice at 28 dpi. Arrows, lateral dendrites.
  • F
    Proportion of neurons, classified by their dendritic patterns, in control (six mice) and PlexinD1‐cKO (five mice) groups. *P < 0.05, unpaired t‐test.
  • G
    Anti‐GABA (upper panels, green) and NeuN (lower panels, green) staining of OB sections from control and Ad‐Cre; PlexinD1‐cKO mice at 28 dpi. Nuclei were stained with Hoechst 33342 (blue).
  • H
    Numbers of GABA+ PGCs and NeuN+ GCs in control (six mice) and Ad‐Cre;PlexinD1‐cKO (six mice) mice. *P < 0.05, ***P < 0.005, unpaired t‐test.
  • I
    Proportions of PGCs and GCs in the OB interneurons (PGCs + GCs) in control (six mice) and Ad‐Cre;PlexinD1‐cKO (six mice) mice. *P < 0.05, unpaired t‐test.
  • J
    Olfactory habituation and discrimination in control (seven mice) and Ad‐Cre; PlexinD1‐cKO (seven mice) mice. ### P < 0.005 (vs. first session), *P < 0.05, repeated measure ANOVA followed by Bonferroni test. Gray lines represent individual mouse data.
Data information: GL, glomerular layer; s/dEPL, superficial/deep external plexiform layer; MCL, mitral cell layer; IPL, inner plexiform layer; s/m/dGCL, superficial/middle/deep granule cell layer; RMS, rostral migratory stream. Scale bars: 20 μm. Bars indicate mean ± SEM.

Mature neurons extend distinct dendrite patterns based on their position in the OB. While PGCs are located in the GL and extend their dendrites within the glomerulus, deep and superficial GCs extend dendrites to the deep and superficial EPLs, where they connect to mitral and tufted cell dendrites, respectively (Shepherd et al, 2004; Fig 7A). To study the dendritic patterns of abnormally positioned neurons, we injected Cre‐expressing lentivirus into the V‐SVZ of PlexinD1 fl/fl mice and analyzed the dendritic morphology of the infected neurons 28 days later. The proportions of PGCs and superficial dendrite‐bearing GCs were significantly decreased and that of deep dendrite‐bearing GCs was significantly increased in PlexinD1‐cKO mice (Fig 7E and F; Appendix Fig S11A and B). PlexinD1‐cKO neurons adapted their dendritic patterns appropriately, according to their position (Appendix Fig S11C–F). These results suggested that abnormally positioned neurons maintained their potential for dendritic development based on their position.

Since mitral and tufted cells project axons to different targets in the olfactory cortex and exhibit distinct electrophysiological properties upon olfactory stimulation (Nagayama et al, 2004; Igarashi et al, 2012), alterations in the dendritic patterning of new neurons in the OB might cause functional deficits. To analyze the effects of the abnormal positioning of PlexinD1‐deficient neurons on olfactory behaviors, we knocked out PlexinD1 specifically in postnatal NSCs by adenoviral targeting. The intraventricular injection of Cre‐encoding adenovirus (Ad‐Cre) into postnatal day 0 (P0) PlexinD1 fl/fl mice, resulting in Ad‐Cre; PlexinD1‐cKO animals, enabled the specific and efficient targeting of periventricular NSCs (Appendix Fig S12A–C) and caused abnormal OB interneuron positioning (Fig 7G and H). The proportion of PGCs in the OB interneurons (PGCs + GCs) was decreased by the PlexinD1 deficiency (Fig 7I). Using four different behavioral analyses, we found that these mice displayed impaired odor habituation and discrimination (Fig 7J; Appendix Fig S12D–F). To test whether the elimination of postnatally born OB interneurons also caused these olfactory deficits, we generated Ad‐Cre; NSE‐DTA mice by intraventricularly injecting Ad‐Cre into P0 NSE‐DTA mice (Kobayakawa et al, 2007; Imayoshi et al, 2008). Expression of diphtheria toxin fragment A under the control of the neuron‐specific enolase gene promoter eliminated postnatally generated OB interneurons, but did not cause these olfactory deficits (Appendix Fig S12G–I), as reported in the adult brain previously (Imayoshi et al, 2008). Together, these results suggested that the appropriate positioning of new neurons controlled by Sema3E‐PlexinD1 signaling contributes to olfactory function.

Discussion

In this study, we examined the mechanism for the termination of new‐neuron migration in the postnatal OB. FLP formation was observed at the proximal leading process of new neurons in the resting phase of migration. A downregulation of Sema3E‐PlexinD1 signaling locally increased the Rac1 activity in the proximal leading process of new neurons, which promoted FLP formation and the maintenance of migration termination, thus contributing to the positioning, dendritic patterns, and functions of new neurons.

Although various morphological changes accompanying the termination of neuronal migration have been described (Nadarajah et al, 2001), our observations using light and electron microscopies suggest that the FLP is a unique protrusion that is spatiotemporally regulated and distinct from any other protrusions, including filopodia. FLPs were observed not only in the V‐SVZ‐derived new neurons (Fig 1) but also in other migrating neurons generated in the embryonic ganglionic eminences (Appendix Fig S2). All of these neurons extend a long leading process, in which proximal‐region cytoskeletal changes drive somal translocation (Shinohara et al, 2012; Yang et al, 2012; Nishimura et al, 2014; Ota et al, 2014; Trivedi et al, 2017), and distal‐region cytoskeletal changes determine directionality (Martini et al, 2009; Lysko et al, 2011, 2014). Since FLPs can develop into lateral dendrites after neuronal migration termination (Appendix Fig S1C and D), the mechanism for FLP formation, which is controlled by Sema3E‐PlexinD1 signaling, could regulate migration termination and the initiation of differentiation, simultaneously and rapidly, providing an efficient strategy for neuronal development.

After Rac1‐induced FLP formation, we observed MT reorganization in the proximal leading process, disruption of the perinuclear cage, and suppression of somal translocation. However, it was possible that the photoactivation of PA‐Rac1CA in the proximal leading process altered the cytoskeletal organization or other unknown cell machineries, to affect neuronal migration independently of the FLP. Moreover, even if we carefully adjusted the laser illumination to restrict the photoactivation within the FLP, the activated Rac1DN protein could diffuse and directly affect the endogenous Rac1 activity in the leading process. On the other hand, our experiments using PST‐1, which inhibited the MT polymerization independently of Rac1 inhibition and without retracting the FLP structure, suggested that MT polymerization within the FLP suppressed the somal translocation of new neurons, thereby maintaining the resting phase. It remains unknown whether FLP formation causes the local MT reorganization in the proximal leading process or vice versa. However, we found that inhibiting the MT polymerization in the FLP caused a recovery of directional MT polymerization in the proximal leading process, while local MT reorganization in the proximal leading process (Fig 6E) did not induce FLPs. Therefore, it is unlikely that the FLP forms as a result of MT reorganization in the proximal leading process. It has been reported that branched structures of the leading process are related to slowed neuronal migration (Koizumi et al, 2006; Guerrier et al, 2009; Belvindrah et al, 2017) and that branched processes develop into apical dendrites (Nadarajah et al, 2001; O'Dell et al, 2015) after migration termination in the developing cerebral cortex. It is possible that these branches are controlled by similar mechanisms as the FLPs in the OB.

The somal translocation of migrating neurons is achieved by the coordinated regulation of RhoA‐actomyosin signaling (Bellion et al, 2005; Schaar & McConnell, 2005; Tsai et al, 2007; Godin et al, 2012; Shinohara et al, 2012; Ota et al, 2014) and dynamic MT organization (Koizumi et al, 2006; Tsai et al, 2007; Umeshima et al, 2007; Belvindrah et al, 2017). We found that the somal translocation occurring after FLP retraction required directional MT polymerization in the proximal leading process. Consistent with these findings, it is reported that the MT network in the proximal leading process and soma (Umeshima et al, 2007; Belvindrah et al, 2017) and MT‐related motor regulators such as dynein, LIS1, and Dcx (Koizumi et al, 2006; Tsai et al, 2007) contribute to somal translocation by providing a scaffold on which dynein motors can pull the nucleus.

Postnatal NSCs in different regions of the V‐SVZ produce different types of olfactory interneurons (Kelsch et al, 2007; Merkle et al, 2007, 2014). Therefore, it is possible that the timing of PlexinD1 downregulation, which controls FLP formation and neuronal migration termination in the OB, is intrinsically programmed in each new neuron or its parental NSC in the V‐SVZ. The timing of migration termination and the subsequent integration of cortical interneurons are also intrinsically determined by cellular age (Bortone & Polleux, 2009; Inamura et al, 2012; Southwell et al, 2012). In contrast, interestingly, our transplantation study showed that the duration and distance of migration are not strictly programmed in each V‐SVZ‐derived new neuron (Fig 2). Instead, transplanted GCs and PGCs could maintain their migratory potential until they reached their final destinations, indicating that the migratory potential has plasticity. These findings suggest that the V‐SVZ‐derived new neurons adjust their migratory potential according to their environment. Furthermore, the Sema3E‐PlexinD1 signaling‐mediated final positioning of new neurons affects their dendritic patterns and functions, consistent with previous studies in the developing cerebral cortex (Oishi et al, 2016) and postnatal OB (Ota et al, 2014). Therefore, it is possible that not only intrinsic programs (Kelsch et al, 2007; Merkle et al, 2007, 2014) but also positional cues such as layer‐specific centrifugal inputs (Kaneko et al, 2006; Garcia‐Gonzalez et al, 2017) contribute to the migration and maturation processes of new neurons, which have plastic potentials in the postnatal OB.

Sema3E and PlexinD1 serve as a guidance signal for axonal extension and synaptic formation in the central nervous system (Chauvet et al, 2007; Pecho‐Vrieseling et al, 2009; Ding et al, 2012; Deck et al, 2013; Fukuhara et al, 2013). Moreover, Sema3E‐PlexinD1 signaling reduces the motogenic potential of migrating Cajal‐Retzius cells (Bribian et al, 2014) and enhances the survival of gonadotropin‐releasing hormone neurons (Cariboni et al, 2015). In this study, we demonstrated that Sema3E maintains the neuronal migratory morphology by suppressing Rac1‐induced FLP formation, in a PlexinD1‐dependent manner. Unlike another member of the Plexin family, PlexinB2, which is reported to control neurogenesis and cell‐fate specifications in the V‐SVZ (Saha et al, 2012), PlexinD1 was not detectable in NSCs or neural progenitor cells, and its deletion did not affect neurogenesis or fate specification. It is possible that mature interneurons start to secrete Sema3E after they stop migrating, to help maintain the migration capacity of later born neurons. Furthermore, considering that new neurons that migrated longer distances had higher levels of PlexinD1, and that deleting and overexpressing PlexinD1 in migrating new neurons altered their final positioning, the signals inducing a local decrease of PlexinD1 in migrating new neurons and Sema3E secretion from mature interneurons could determine the interneuron distribution in the postnatal OB. Although Sema3E KO mice did not show any visible disorganization of the gross OB layer structures (Appendix Fig S6I), we could not exclude the possibility that the migration defects were partly caused by developmental defects in the brain of the Sema3E KO mice (Chauvet et al, 2007; Cariboni et al, 2015). FLP formation could be observed in cell types with a highly elongated morphology such as migrating neurons derived from the postnatal V‐SVZ (Fig 1) and embryonic ganglionic eminences (Appendix Fig S2). It is possible that this mechanism we discovered in migrating neurons also acts as a general regulator of the migration of highly polarized cells such as neural and vascular endothelial cells (Ono et al, 1997; Gerhardt et al, 2003; Bellion et al, 2005; Schaar & McConnell, 2005; Abraham et al, 2015; Tsai et al, 2016).

This study suggests that local Rac1 activation induces FLPs to promote the morphological transition of migrating immature neurons to mature neurons, an important step in the development and maintenance of neuronal circuits under physiological conditions. Under pathological conditions, PlexinD1 mutations in Möbius syndrome patients and in mice cause similar disturbances in neuronal migration (Tomas‐Roca et al, 2015), suggesting that a disruption in the morphological regulation of migrating neurons by PlexinD1 may be involved in neurological disorders. After brain injury, V‐SVZ‐derived new neurons migrate toward the lesion and differentiate into mature neurons in the striatum and cerebral cortex (Massouh & Saghatelyan, 2010; Sawada et al, 2011; Kaneko et al, 2017), where Sema3E is expressed during the postnatal period (Watakabe et al, 2006), suggesting that Sema3E‐PlexinD1 signaling contributes to endogenous neuronal regeneration. In conclusion, this study suggests that MT polymerization within the FLP, the formation of which is controlled by Sema3E‐PlexinD1‐Rac1 signaling, enables the maintenance of new‐neuron migration termination and highlights the importance of these mechanisms for the proper positioning of cells in neuronal circuits.

Materials and Methods

Animals

Wild‐type (WT) C57BL/6J mice were purchased from Japan SLC. Dcx‐EGFP mice (Gong et al, 2003) were provided by the Mutant Mouse Research Resource Center (MMRRC). PlexinD1‐flox mice (Zhang et al, 2009) and CAG‐MerCreMer mice (Egawa et al, 2009) were described previously. Sema3E KO mice (Gu et al, 2005) were provided by Dr. Fanny Mann (Institut de Biologie du Developpement de Marseille) and Dr. Christopher E. Henderson (Columbia University). Tamoxifen solution (Sigma, 20 mg/ml) was intraperitoneally injected into CAG‐MerCreMer; PlexinD1 fl/fl mice (200 μg/g body weight) once a day for two consecutive days (Appendix Fig S5H), or once at embryonic day 16 (Appendix Fig S10A–E). All of the animals used in this study were maintained within seven mice per cage and on a 12‐h light/dark cycle with ad libitum access to food and water. All of the experiments involving live animals were performed in accordance with the guidelines and regulations of Nagoya City University.

Transplantation

The PGC‐rich and GC‐rich regions of the OB from postnatal day 0–2 (P0‐2) Dcx‐GFP mice were dissected and then dissociated with trypsin–EDTA (Invitrogen). The cells were suspended in L‐15 medium (Invitrogen) and immediately transplanted into the V‐SVZ of P1 WT mice. In the PlexinD1‐KD experiment (Appendix Fig S9J–L), the dissociated cells were infected with lentivirus carrying control‐ or PlexinD1‐shRNA#1 for 1 h at 4°C and then injected into the V‐SVZ of P1 WT mice. Eight or 14 days after transplantation, the morphology and distribution of the transplanted cells in the OB layers of the recipient mice were analyzed.

Slice culture imaging

Imaging of migrating neurons in OB slice cultures was performed as described previously (Ota et al, 2014), with some modifications. Migrating neurons were lentivirally labeled with viruses injected into the V‐SVZ of P0‐1 mice. Six to seven days after virus injection, 200‐μm‐thick sagittal brain slices were prepared using a vibratome (VT‐1200, Leica) and then cultured on a filter membrane (Millipore) in Neurobasal medium (Gibco), supplemented with 20% fetal bovine serum, 2% NeuroBrew‐21 (Invitrogen), 2 mM l‐glutamine (Gibco), and 50 U/ml penicillin–streptomycin (Gibco), in a stage‐top chamber at 37°C in a 5% CO2 incubation system (Tokai Hit). Time‐lapse images were captured at 10‐min intervals for 12–24 h using an LSM710 laser‐scanning confocal microscope (Carl Zeiss). The neuronal migration speed and frequency of FLP formation were quantified using the ImageJ manual tracking tool. The maximal length and position of protrusions were quantified using ZEN software (Carl Zeiss). The resting phase was defined as a migration speed less than 12 μm/h. To determine the effect of endocytosis inhibitors on PlexinD1 expression, 200‐μm‐thick sagittal brain slices were prepared 6 days after PlexinD1‐GFP‐expressing lentivirus injection and cultured on a filter membrane with 30 μM myristyl trimethyl ammonium bromides (MiTMAB) or 50 μM monodansylcadaverine (MDC) at 37°C for 16 h. The cultured slices were then fixed with 4% paraformaldehyde (PFA) in 0.1 M phosphate buffer (PB) (pH 7.4) and subjected to immunohistochemistry using an anti‐GFP antibody (1:200, MBL).

In vitro culture of migrating neurons

The V‐SVZ was dissected from P0‐1 mice and dissociated with trypsin–EDTA (Invitrogen). The lateral, medial, and caudal ganglionic eminences were dissected from embryonic day 15 (E15) embryos. The cells were washed in L‐15 medium (Invitrogen) containing 40 μg/ml DNase I (Roche) and transfected with 2.0–2.25 μg plasmid DNA using the Amaxa Nucleofector II system (Lonza). The transfected cells were recovered in RPMI‐1640 medium (Thermo Fisher Scientific), allowed to aggregate, and then embedded in 50% Matrigel (BD Biosciences) in L‐15 medium. The cell aggregates were cultured in Neurobasal medium containing 2% NeuroBrew‐21 (Invitrogen), 2 mM l‐glutamine (Gibco), and 50 U/ml penicillin–streptomycin (Gibco) for 48–72 h. Please also see the Appendix Materials and Methods.

FRET imaging

FRET imaging of the Rho family of small GTPases in V‐SVZ‐derived migrating neurons in vitro was performed as described previously (Ota et al, 2014). The FRET probes for Rac1, cdc42, and RhoA (Raichu‐1011X, 1054X, and 1237X, respectively) were kind gifts from Dr. Michiyuki Matsuda (Kyoto University) (Yoshizaki et al, 2003). For the FRET imaging in brain slices, Riachu‐1011X was introduced into the V‐SVZ cells by in vivo electroporation as reported previously (Ota et al, 2014). Time‐lapse imaging of the FRET probe‐expressing migrating neurons was performed using an LSM710 laser‐scanning confocal microscope (Carl Zeiss) and a 40× water‐immersion objective lens. The cyan fluorescent protein (CFP) channel was excited using a 458‐nm argon laser. Two emission channels were split using a 545‐nm dichroic mirror, and a 475–525‐nm bandpass filter for CFP and a 530‐nm longpass filter for yellow fluorescent protein (YFP). The FRET ratio (intensity of FRET/CFP) was calculated, and their final images were generated with the MetaMorph software ratio image function (Molecular Probes) using eight ratios and 32 intensities. The baseline FRET activity was calculated by averaging the basal activities in the leading shaft over a 180‐min time period, and normalizing the average in each cell to 1.0. The extent of activation of each of the Rho family GTPases (FRETactivation) in the leading tip and shaft in a circular region of interest (ROI) was measured by the MetaMorph software Region measurements function, and normalized to the baseline activity in each frame (activity increase = FRETactivation − 1). Rac1 activation per hour in the leading tip or shaft of each cell is shown as ∑(activity increase)/h. At least three independent experiments were performed. In Fig 4K, FLPs that had disappeared or still existed 7 min later were defined as disappearing or stable FLPs, respectively. In the brain slice experiment, all of the probe‐expressing neurons with a leading process migrating over 40 min were analyzed.

Rac1 photoactivation

The photoactivatable (PA)‐Rac1 probes (pTriEx‐mVenus‐PA‐Rac1, pTriEx‐mVenus‐PA‐Rac1T17N, and pTriEx‐mVenus‐PA‐Rac1‐C450A, Addgene plasmids #22007, #22017, and #22021, respectively) were gifts from Dr. Klaus Hahn (University of North Carolina; Wu et al, 2009). Time‐lapse imaging of PA‐Rac1 probe‐expressing migrating neurons was performed using an LSM710 laser‐scanning confocal microscope (Carl Zeiss) and a 63× oil‐immersion objective lens. A circular ROI was set at the proximal leading process or FLP of the neurons and photoactivated using a 458‐nm (0.28 mW) argon laser at 30‐s (Fig 5A–E), 4‐min (Fig 5E, left), 15‐s (Fig 5F–H), or 5‐s (Fig 5I–K) intervals. The efficiency of photoactivation‐induced FLP formation was defined as the number of cells exhibiting photoactivation‐induced FLPs/the number of photoactivated cells. Neuronal migration speeds were calculated using the ImageJ manual tracking tool. For the FLP retraction analysis (Fig 5A–C, I–K), FLP retraction was defined as a > 50% decrease in the FLP length. At least three independent experiments were performed.

Photoswitching of photostatin‐1

Time‐lapse imaging of 2 μM PST‐1‐treated migrating neurons was performed using an LSM710 laser‐scanning confocal microscope (Carl Zeiss) with a 63× oil‐immersion objective lens. PST‐1 was photoswitched using 405‐nm (1.55 mW) and 514‐nm (1.29 mW) lasers at 15‐ to 25‐s intervals in a circular ROI set at the proximal leading process or FLP, respectively (Fig 6A–G; Appendix Fig S8C and D). PST‐1 was inactivated by 514‐nm laser illumination in a square ROI set at the proximal leading process (Fig 6A–D). Inhibition of MT polymerization in the FLP was monitored using EB3‐GFP (a marker of MT plus‐ends) by analyzing the density of EB3‐GFP+ dots (Fig 6A' and C). At least three independent experiments were performed.

Statistical analysis

Sample sizes were not predetermined, but were chosen based on previous studies. No statistical calculation was used to estimate the sample size. The experiments were not randomized, and the investigator was not blinded except for the KO studies (Fig 7C, D and H) and behavioral tests. The data are presented as the mean ± SEM. The data distribution was analyzed by the Kolmogorov–Smirnov test and/or Shapiro–Wilk test. The equality of variance between groups was analyzed by the F test. All of the data were two‐tailed. Comparisons between two groups were analyzed by paired t‐test, unpaired t‐test, or Mann–Whitney U‐test. Comparisons among multiple groups were analyzed by one‐way ANOVA, repeated‐measures ANOVA, or Kruskal–Wallis test followed by a post hoc Tukey–Kramer, Dunnett, Bonferroni, or Steel–Dwass test. Gaussian fitting curves for the protrusion distribution (Fig 1B) were created based on the minimum value of Akaike's information criterion (AIC) using the Solver add‐in tool (Excel, Microsoft) (bimodal distribution, AIC = 509.24). P < 0.05 were considered to be statistically significant.

Additional Materials and Methods are included in the Appendix.

Author contributions

MS, MK, HN, and KS designed the experiments and wrote the manuscript. MS, NO, MK, S‐hH, TH, YS, HBN, TQT, YI, YY, HN, AU, and KS performed the experiments and analyzed the data.

Conflict of interest

The authors declare that they have no conflict of interest.

Supporting information

Appendix

Click here for additional data file. (21.3MB, pdf)

Movie EV1

Click here for additional data file. (1.7MB, zip)

Movie EV2

Click here for additional data file. (199.5KB, zip)

Movie EV3

Click here for additional data file. (816.8KB, zip)

Movie EV4

Click here for additional data file. (8MB, zip)

Movie EV5

Click here for additional data file. (1.5MB, zip)

Review Process File

Click here for additional data file. (456.2KB, pdf)

Acknowledgements

We thank M. Kengaku and T. Kawauchi for critical reading of the manuscript; F. Mann, C. E. Henderson, M. Yamaguchi, S. Itohara, T. Ikeda, R. Kageyama, M. Götz, H. Miyoshi, T. Toyofuku, I. Oinuma, M. Matsuda, D. J. Solecki, W. Bement, K. Hahn, K. Kaibuchi, K. Yoshikawa, M. Hattori, and the MMRRC for materials; H. Takase, T. Miyamoto, I. Miyoshi, N. Dohi, K. Ikenaka, M. Furuse, M. Takahashi, and M. Takagishi for technical support; and J.M. García‐Verdugo, V. Herranz‐Pérez, A. Cebrián‐Silla, and Sawamoto laboratory members for discussions. This work was supported by research grants from NEXT (LS104), MEXT KAKENHI (22122004, 17H05750, and 17H05512) (to K.S.), JSPS KAKENHI (26250019, 17H01392, and JP16H06280 [to K.S.], 25890017, and 26830014 [to M.S.]), JSPS Program for Advancing Strategic International Networks to Accelerate the Circulation of Talented Researchers (S2704 [to M.S., M.K., H.N., and K.S.]), Grant‐in‐Aid for Research at Nagoya City University (to K.S. and H.N.), NIH grants NS093002 (to Y.Y.), the Novartis Foundation (Japan) for the Promotion of Science, the Hori Sciences and Arts Foundation (to M.S.), the Takeda Science Foundation (to K.S. and M.S.), and the Cooperative Study Programs of National Institute for Physiological Sciences (to K.S.).

The EMBO Journal (2018) 37: e97404 29348324

References

  1. Abraham S, Scarcia M, Bagshaw RD, McMahon K, Grant G, Harvey T, Yeo M, Esteves FO, Thygesen HH, Jones PF, Speirs V, Hanby AM, Selby PJ, Lorger M, Dear TN, Pawson T, Marshall CJ, Mavria G (2015) A Rac/Cdc42 exchange factor complex promotes formation of lateral filopodia and blood vessel lumen morphogenesis. Nat Commun 6: 7286 [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Anton ES, Ghashghaei HT, Weber JL, McCann C, Fischer TM, Cheung ID, Gassmann M, Messing A, Klein R, Schwab MH, Lloyd KC, Lai C (2004) Receptor tyrosine kinase ErbB4 modulates neuroblast migration and placement in the adult forebrain. Nat Neurosci 7: 1319–1328 [DOI] [PubMed] [Google Scholar]
  3. Bedard K, Krause KH (2007) The NOX family of ROS‐generating NADPH oxidases: physiology and pathophysiology. Physiol Rev 87: 245–313 [DOI] [PubMed] [Google Scholar]
  4. Bellion A, Baudoin JP, Alvarez C, Bornens M, Metin C (2005) Nucleokinesis in tangentially migrating neurons comprises two alternating phases: forward migration of the Golgi/centrosome associated with centrosome splitting and myosin contraction at the rear. J Neurosci 25: 5691–5699 [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Belvindrah R, Natarajan K, Shabajee P, Bruel‐Jungerman E, Bernard J, Goutierre M, Moutkine I, Jaglin XH, Savariradjane M, Irinopoulou T, Poncer JC, Janke C, Francis F (2017) Mutation of the alpha‐tubulin Tuba1a leads to straighter microtubules and perturbs neuronal migration. J Cell Biol 216: 2443–2461 [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Borowiak M, Nahaboo W, Reynders M, Nekolla K, Jalinot P, Hasserodt J, Rehberg M, Delattre M, Zahler S, Vollmar A, Trauner D, Thorn‐Seshold O (2015) Photoswitchable inhibitors of microtubule dynamics optically control mitosis and cell death. Cell 162: 403–411 [DOI] [PubMed] [Google Scholar]
  7. Bortone D, Polleux F (2009) KCC2 expression promotes the termination of cortical interneuron migration in a voltage‐sensitive calcium‐dependent manner. Neuron 62: 53–71 [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Bribian A, Nocentini S, Llorens F, Gil V, Mire E, Reginensi D, Yoshida Y, Mann F, del Rio JA (2014) Sema3E/PlexinD1 regulates the migration of hem‐derived Cajal‐Retzius cells in developing cerebral cortex. Nat Commun 5: 4265 [DOI] [PubMed] [Google Scholar]
  9. Burkel BM, von Dassow G, Bement WM (2007) Versatile fluorescent probes for actin filaments based on the actin‐binding domain of utrophin. Cell Motil Cytoskeleton 64: 822–832 [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. Cariboni A, Andre V, Chauvet S, Cassatella D, Davidson K, Caramello A, Fantin A, Bouloux P, Mann F, Ruhrberg C (2015) Dysfunctional SEMA3E signaling underlies gonadotropin‐releasing hormone neuron deficiency in Kallmann syndrome. J Clin Invest 125: 2413–2428 [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Chauvet S, Cohen S, Yoshida Y, Fekrane L, Livet J, Gayet O, Segu L, Buhot MC, Jessell TM, Henderson CE, Mann F (2007) Gating of Sema3E/PlexinD1 signaling by neuropilin‐1 switches axonal repulsion to attraction during brain development. Neuron 56: 807–822 [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Cooper JA (2013) Cell biology in neuroscience: mechanisms of cell migration in the nervous system. J Cell Biol 202: 725–734 [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Dang PM, Cross AR, Babior BM (2001) Assembly of the neutrophil respiratory burst oxidase: a direct interaction between p67PHOX and cytochrome b558. Proc Natl Acad Sci USA 98: 3001–3005 [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Deck M, Lokmane L, Chauvet S, Mailhes C, Keita M, Niquille M, Yoshida M, Yoshida Y, Lebrand C, Mann F, Grove EA, Garel S (2013) Pathfinding of corticothalamic axons relies on a rendezvous with thalamic projections. Neuron 77: 472–484 [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Dent EW, Kwiatkowski AV, Mebane LM, Philippar U, Barzik M, Rubinson DA, Gupton S, Van Veen JE, Furman C, Zhang J, Alberts AS, Mori S, Gertler FB (2007) Filopodia are required for cortical neurite initiation. Nat Cell Biol 9: 1347–1359 [DOI] [PubMed] [Google Scholar]
  16. Ding JB, Oh WJ, Sabatini BL, Gu C (2012) Semaphorin 3E‐Plexin‐D1 signaling controls pathway‐specific synapse formation in the striatum. Nat Neurosci 15: 215–223 [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Doetsch F, Caille I, Lim DA, Garcia‐Verdugo JM, Alvarez‐Buylla A (1999) Subventricular zone astrocytes are neural stem cells in the adult mammalian brain. Cell 97: 703–716 [DOI] [PubMed] [Google Scholar]
  18. Egawa G, Osawa M, Uemura A, Miyachi Y, Nishikawa S (2009) Transient expression of ephrin b2 in perinatal skin is required for maintenance of keratinocyte homeostasis. J Invest Dermatol 129: 2386–2395 [DOI] [PubMed] [Google Scholar]
  19. Fukata M, Watanabe T, Noritake J, Nakagawa M, Yamaga M, Kuroda S, Matsuura Y, Iwamatsu A, Perez F, Kaibuchi K (2002) Rac1 and Cdc42 capture microtubules through IQGAP1 and CLIP‐170. Cell 109: 873–885 [DOI] [PubMed] [Google Scholar]
  20. Fukuhara K, Imai F, Ladle DR, Katayama K, Leslie JR, Arber S, Jessell TM, Yoshida Y (2013) Specificity of monosynaptic sensory‐motor connections imposed by repellent Sema3E‐PlexinD1 signaling. Cell Rep 5: 748–758 [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Garcia‐Gonzalez D, Khodosevich K, Watanabe Y, Rollenhagen A, Lubke JHR, Monyer H (2017) Serotonergic projections govern postnatal neuroblast migration. Neuron 94: 534–549 [DOI] [PubMed] [Google Scholar]
  22. Gerhardt H, Golding M, Fruttiger M, Ruhrberg C, Lundkvist A, Abramsson A, Jeltsch M, Mitchell C, Alitalo K, Shima D, Betsholtz C (2003) VEGF guides angiogenic sprouting utilizing endothelial tip cell filopodia. J Cell Biol 161: 1163–1177 [DOI] [PMC free article] [PubMed] [Google Scholar]
  23. Ghashghaei HT, Lai C, Anton ES (2007) Neuronal migration in the adult brain: are we there yet? Nat Rev Neurosci 8: 141–151 [DOI] [PubMed] [Google Scholar]
  24. Godin JD, Thomas N, Laguesse S, Malinouskaya L, Close P, Malaise O, Purnelle A, Raineteau O, Campbell K, Fero M, Moonen G, Malgrange B, Chariot A, Metin C, Besson A, Nguyen L (2012) p27(Kip1) is a microtubule‐associated protein that promotes microtubule polymerization during neuron migration. Dev Cell 23: 729–744 [DOI] [PubMed] [Google Scholar]
  25. Gong S, Zheng C, Doughty ML, Losos K, Didkovsky N, Schambra UB, Nowak NJ, Joyner A, Leblanc G, Hatten ME, Heintz N (2003) A gene expression atlas of the central nervous system based on bacterial artificial chromosomes. Nature 425: 917–925 [DOI] [PubMed] [Google Scholar]
  26. Gongidi V, Ring C, Moody M, Brekken R, Sage EH, Rakic P, Anton ES (2004) SPARC‐like 1 regulates the terminal phase of radial glia‐guided migration in the cerebral cortex. Neuron 41: 57–69 [DOI] [PubMed] [Google Scholar]
  27. Gu C, Yoshida Y, Livet J, Reimert DV, Mann F, Merte J, Henderson CE, Jessell TM, Kolodkin AL, Ginty DD (2005) Semaphorin 3E and plexin‐D1 control vascular pattern independently of neuropilins. Science 307: 265–268 [DOI] [PubMed] [Google Scholar]
  28. Guerrier S, Coutinho‐Budd J, Sassa T, Gresset A, Jordan NV, Chen K, Jin WL, Frost A, Polleux F (2009) The F‐BAR domain of srGAP2 induces membrane protrusions required for neuronal migration and morphogenesis. Cell 138: 990–1004 [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Hack I, Bancila M, Loulier K, Carroll P, Cremer H (2002) Reelin is a detachment signal in tangential chain‐migration during postnatal neurogenesis. Nat Neurosci 5: 939–945 [DOI] [PubMed] [Google Scholar]
  30. Igarashi KM, Ieki N, An M, Yamaguchi Y, Nagayama S, Kobayakawa K, Kobayakawa R, Tanifuji M, Sakano H, Chen WR, Mori K (2012) Parallel mitral and tufted cell pathways route distinct odor information to different targets in the olfactory cortex. J Neurosci 32: 7970–7985 [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Imayoshi I, Sakamoto M, Ohtsuka T, Takao K, Miyakawa T, Yamaguchi M, Mori K, Ikeda T, Itohara S, Kageyama R (2008) Roles of continuous neurogenesis in the structural and functional integrity of the adult forebrain. Nat Neurosci 11: 1153–1161 [DOI] [PubMed] [Google Scholar]
  32. Inamura N, Kimura T, Tada S, Kurahashi T, Yanagida M, Yanagawa Y, Ikenaka K, Murakami F (2012) Intrinsic and extrinsic mechanisms control the termination of cortical interneuron migration. J Neurosci 32: 6032–6042 [DOI] [PMC free article] [PubMed] [Google Scholar]
  33. Kaneko N, Okano H, Sawamoto K (2006) Role of the cholinergic system in regulating survival of newborn neurons in the adult mouse dentate gyrus and olfactory bulb. Genes Cells 11: 1145–1159 [DOI] [PubMed] [Google Scholar]
  34. Kaneko N, Marin O, Koike M, Hirota Y, Uchiyama Y, Wu JY, Lu Q, Tessier‐Lavigne M, Alvarez‐Buylla A, Okano H, Rubenstein JL, Sawamoto K (2010) New neurons clear the path of astrocytic processes for their rapid migration in the adult brain. Neuron 67: 213–223 [DOI] [PMC free article] [PubMed] [Google Scholar]
  35. Kaneko N, Sawada M, Sawamoto K (2017) Mechanisms of neuronal migration in the adult brain. J Neurochem 141: 835–847 [DOI] [PubMed] [Google Scholar]
  36. Kelsch W, Mosley CP, Lin CW, Lois C (2007) Distinct mammalian precursors are committed to generate neurons with defined dendritic projection patterns. PLoS Biol 5: e300 [DOI] [PMC free article] [PubMed] [Google Scholar]
  37. Kobayakawa K, Kobayakawa R, Matsumoto H, Oka Y, Imai T, Ikawa M, Okabe M, Ikeda T, Itohara S, Kikusui T, Mori K, Sakano H (2007) Innate versus learned odour processing in the mouse olfactory bulb. Nature 450: 503–508 [DOI] [PubMed] [Google Scholar]
  38. Koga H, Terasawa H, Nunoi H, Takeshige K, Inagaki F, Sumimoto H (1999) Tetratricopeptide repeat (TPR) motifs of p67(phox) participate in interaction with the small GTPase Rac and activation of the phagocyte NADPH oxidase. J Biol Chem 274: 25051–25060 [DOI] [PubMed] [Google Scholar]
  39. Koizumi H, Higginbotham H, Poon T, Tanaka T, Brinkman BC, Gleeson JG (2006) Doublecortin maintains bipolar shape and nuclear translocation during migration in the adult forebrain. Nat Neurosci 9: 779–786 [DOI] [PubMed] [Google Scholar]
  40. Lapouge K, Smith SJ, Walker PA, Gamblin SJ, Smerdon SJ, Rittinger K (2000) Structure of the TPR domain of p67phox in complex with Rac.GTP. Mol Cell 6: 899–907 [DOI] [PubMed] [Google Scholar]
  41. Lois C, Alvarez‐Buylla A (1994) Long‐distance neuronal migration in the adult mammalian brain. Science 264: 1145–1148 [DOI] [PubMed] [Google Scholar]
  42. Lois C, Garcia‐Verdugo JM, Alvarez‐Buylla A (1996) Chain migration of neuronal precursors. Science 271: 978–981 [DOI] [PubMed] [Google Scholar]
  43. Luskin MB (1993) Restricted proliferation and migration of postnatally generated neurons derived from the forebrain subventricular zone. Neuron 11: 173–189 [DOI] [PubMed] [Google Scholar]
  44. Lysko DE, Putt M, Golden JA (2011) SDF1 regulates leading process branching and speed of migrating interneurons. J Neurosci 31: 1739–1745 [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Lysko DE, Putt M, Golden JA (2014) SDF1 reduces interneuron leading process branching through dual regulation of actin and microtubules. J Neurosci 34: 4941–4962 [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Marin O (2013) Cellular and molecular mechanisms controlling the migration of neocortical interneurons. Eur J Neurosci 38: 2019–2029 [DOI] [PubMed] [Google Scholar]
  47. Martini FJ, Valiente M, Lopez Bendito G, Szabo G, Moya F, Valdeolmillos M, Marin O (2009) Biased selection of leading process branches mediates chemotaxis during tangential neuronal migration. Development 136: 41–50 [DOI] [PubMed] [Google Scholar]
  48. Massouh M, Saghatelyan A (2010) De‐routing neuronal precursors in the adult brain to sites of injury: role of the vasculature. Neuropharmacology 58: 877–883 [DOI] [PubMed] [Google Scholar]
  49. Merkle FT, Mirzadeh Z, Alvarez‐Buylla A (2007) Mosaic organization of neural stem cells in the adult brain. Science 317: 381–384 [DOI] [PubMed] [Google Scholar]
  50. Merkle FT, Fuentealba LC, Sanders TA, Magno L, Kessaris N, Alvarez‐Buylla A (2014) Adult neural stem cells in distinct microdomains generate previously unknown interneuron types. Nat Neurosci 17: 207–214 [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Nadarajah B, Brunstrom JE, Grutzendler J, Wong RO, Pearlman AL (2001) Two modes of radial migration in early development of the cerebral cortex. Nat Neurosci 4: 143–150 [DOI] [PubMed] [Google Scholar]
  52. Nagayama S, Takahashi YK, Yoshihara Y, Mori K (2004) Mitral and tufted cells differ in the decoding manner of odor maps in the rat olfactory bulb. J Neurophysiol 91: 2532–2540 [DOI] [PubMed] [Google Scholar]
  53. Ng KL, Li JD, Cheng MY, Leslie FM, Lee AG, Zhou QY (2005) Dependence of olfactory bulb neurogenesis on prokineticin 2 signaling. Science 308: 1923–1927 [DOI] [PubMed] [Google Scholar]
  54. Nishimura YV, Shikanai M, Hoshino M, Ohshima T, Nabeshima Y, Mizutani K, Nagata K, Nakajima K, Kawauchi T (2014) Cdk5 and its substrates, Dcx and p27kip1, regulate cytoplasmic dilation formation and nuclear elongation in migrating neurons. Development 141: 3540–3550 [DOI] [PubMed] [Google Scholar]
  55. O'Dell RS, Cameron DA, Zipfel WR, Olson EC (2015) Reelin prevents apical neurite retraction during terminal translocation and dendrite initiation. J Neurosci 35: 10659–10674 [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Oishi K, Nakagawa N, Tachikawa K, Sasaki S, Aramaki M, Hirano S, Yamamoto N, Yoshimura Y, Nakajima K (2016) Identity of neocortical layer 4 neurons is specified through correct positioning into the cortex. Elife 5: e10907 [DOI] [PMC free article] [PubMed] [Google Scholar]
  57. Ono K, Yasui Y, Rutishauser U, Miller RH (1997) Focal ventricular origin and migration of oligodendrocyte precursors into the chick optic nerve. Neuron 19: 283–292 [DOI] [PubMed] [Google Scholar]
  58. Ota H, Hikita T, Sawada M, Nishioka T, Matsumoto M, Komura M, Ohno A, Kamiya Y, Miyamoto T, Asai N, Enomoto A, Takahashi M, Kaibuchi K, Sobue K, Sawamoto K (2014) Speed control for neuronal migration in the postnatal brain by Gmip‐mediated local inactivation of RhoA. Nat Commun 5: 4532 [DOI] [PubMed] [Google Scholar]
  59. Pecho‐Vrieseling E, Sigrist M, Yoshida Y, Jessell TM, Arber S (2009) Specificity of sensory‐motor connections encoded by Sema3e‐Plxnd1 recognition. Nature 459: 842–846 [DOI] [PMC free article] [PubMed] [Google Scholar]
  60. Petri R, Pircs K, Jonsson ME, Akerblom M, Brattas PL, Klussendorf T, Jakobsson J (2017) let‐7 regulates radial migration of new‐born neurons through positive regulation of autophagy. EMBO J 36: 1379–1391 [DOI] [PMC free article] [PubMed] [Google Scholar]
  61. Saghatelyan A, de Chevigny A, Schachner M, Lledo PM (2004) Tenascin‐R mediates activity‐dependent recruitment of neuroblasts in the adult mouse forebrain. Nat Neurosci 7: 347–356 [DOI] [PubMed] [Google Scholar]
  62. Saha B, Ypsilanti AR, Boutin C, Cremer H, Chedotal A (2012) Plexin‐B2 regulates the proliferation and migration of neuroblasts in the postnatal and adult subventricular zone. J Neurosci 32: 16892–16905 [DOI] [PMC free article] [PubMed] [Google Scholar]
  63. Sawada M, Huang S, Hirota Y, Kaneko N, Sawamoto K (2011) Neuronal Migration in the Adult Brain In Neurogenesis in the Adult Brain I, Neurobiology. Seki T, Sawamoto K, Parent JM, Alvarez‐Buylla A. (eds), pp 337–355. Tokyo: Springer; [Google Scholar]
  64. Sawamoto K, Wichterle H, Gonzalez‐Perez O, Cholfin JA, Yamada M, Spassky N, Murcia NS, Garcia‐Verdugo JM, Marin O, Rubenstein JL, Tessier‐Lavigne M, Okano H, Alvarez‐Buylla A (2006) New neurons follow the flow of cerebrospinal fluid in the adult brain. Science 311: 629–632 [DOI] [PubMed] [Google Scholar]
  65. Schaar BT, McConnell SK (2005) Cytoskeletal coordination during neuronal migration. Proc Natl Acad Sci USA 102: 13652–13657 [DOI] [PMC free article] [PubMed] [Google Scholar]
  66. Shepherd GM, Chen WR, Greer CA (2004) Olfactory bulb In The Synaptic Organization of the Brain. Shepherd GM. (ed), pp 165–216. New York: Oxford; [Google Scholar]
  67. Shinohara R, Thumkeo D, Kamijo H, Kaneko N, Sawamoto K, Watanabe K, Takebayashi H, Kiyonari H, Ishizaki T, Furuyashiki T, Narumiya S (2012) A role for mDia, a Rho‐regulated actin nucleator, in tangential migration of interneuron precursors. Nat Neurosci 15: 373–380 [DOI] [PubMed] [Google Scholar]
  68. Simo S, Cooper JA (2013) Rbx2 regulates neuronal migration through different cullin 5‐RING ligase adaptors. Dev Cell 27: 399–411 [DOI] [PMC free article] [PubMed] [Google Scholar]
  69. Snapyan M, Lemasson M, Brill MS, Blais M, Massouh M, Ninkovic J, Gravel C, Berthod F, Gotz M, Barker PA, Parent A, Saghatelyan A (2009) Vasculature guides migrating neuronal precursors in the adult mammalian forebrain via brain‐derived neurotrophic factor signaling. J Neurosci 29: 4172–4188 [DOI] [PMC free article] [PubMed] [Google Scholar]
  70. Southwell DG, Paredes MF, Galvao RP, Jones DL, Froemke RC, Sebe JY, Alfaro‐Cervello C, Tang Y, Garcia‐Verdugo JM, Rubenstein JL, Baraban SC, Alvarez‐Buylla A (2012) Intrinsically determined cell death of developing cortical interneurons. Nature 491: 109–113 [DOI] [PMC free article] [PubMed] [Google Scholar]
  71. Tomas‐Roca L, Tsaalbi‐Shtylik A, Jansen JG, Singh MK, Epstein JA, Altunoglu U, Verzijl H, Soria L, van Beusekom E, Roscioli T, Iqbal Z, Gilissen C, Hoischen A, de Brouwer AP, Erasmus C, Schubert D, Brunner H, Perez Aytes A, Marin F, Aroca P et al (2015) De novo mutations in PLXND1 and REV3L cause Mobius syndrome. Nat Commun 6: 7199 [DOI] [PMC free article] [PubMed] [Google Scholar]
  72. Trivedi N, Stabley DR, Cain B, Howell D, Laumonnerie C, Ramahi JS, Temirov J, Kerekes RA, Gordon‐Weeks PR, Solecki DJ (2017) Drebrin‐mediated microtubule‐actomyosin coupling steers cerebellar granule neuron nucleokinesis and migration pathway selection. Nat Commun 8: 14484 [DOI] [PMC free article] [PubMed] [Google Scholar]
  73. Tsai JW, Bremner KH, Vallee RB (2007) Dual subcellular roles for LIS1 and dynein in radial neuronal migration in live brain tissue. Nat Neurosci 10: 970–979 [DOI] [PubMed] [Google Scholar]
  74. Tsai HH, Niu J, Munji R, Davalos D, Chang J, Zhang H, Tien AC, Kuo CJ, Chan JR, Daneman R, Fancy SP (2016) Oligodendrocyte precursors migrate along vasculature in the developing nervous system. Science 351: 379–384 [DOI] [PMC free article] [PubMed] [Google Scholar]
  75. Uesugi K, Oinuma I, Katoh H, Negishi M (2009) Different requirement for Rnd GTPases of R‐Ras GAP activity of Plexin‐C1 and Plexin‐D1. J Biol Chem 284: 6743–6751 [DOI] [PMC free article] [PubMed] [Google Scholar]
  76. Umeshima H, Hirano T, Kengaku M (2007) Microtubule‐based nuclear movement occurs independently of centrosome positioning in migrating neurons. Proc Natl Acad Sci USA 104: 16182–16187 [DOI] [PMC free article] [PubMed] [Google Scholar]
  77. Watakabe A, Ohsawa S, Hashikawa T, Yamamori T (2006) Binding and complementary expression patterns of semaphorin 3E and plexin D1 in the mature neocortices of mice and monkeys. J Comp Neurol 499: 258–273 [DOI] [PubMed] [Google Scholar]
  78. Watanabe T, Wang S, Noritake J, Sato K, Fukata M, Takefuji M, Nakagawa M, Izumi N, Akiyama T, Kaibuchi K (2004) Interaction with IQGAP1 links APC to Rac1, Cdc42, and actin filaments during cell polarization and migration. Dev Cell 7: 871–883 [DOI] [PubMed] [Google Scholar]
  79. Watanabe T, Kakeno M, Matsui T, Sugiyama I, Arimura N, Matsuzawa K, Shirahige A, Ishidate F, Nishioka T, Taya S, Hoshino M, Kaibuchi K (2015) TTBK2 with EB1/3 regulates microtubule dynamics in migrating cells through KIF2A phosphorylation. J Cell Biol 210: 737–751 [DOI] [PMC free article] [PubMed] [Google Scholar]
  80. Worzfeld T, Swiercz JM, Senturk A, Genz B, Korostylev A, Deng S, Xia J, Hoshino M, Epstein JA, Chan AM, Vollmar B, Acker‐Palmer A, Kuner R, Offermanns S (2014) Genetic dissection of plexin signaling in vivo . Proc Natl Acad Sci USA 111: 2194–2199 [DOI] [PMC free article] [PubMed] [Google Scholar]
  81. Wu YI, Frey D, Lungu OI, Jaehrig A, Schlichting I, Kuhlman B, Hahn KM (2009) A genetically encoded photoactivatable Rac controls the motility of living cells. Nature 461: 104–108 [DOI] [PMC free article] [PubMed] [Google Scholar]
  82. Yang T, Sun Y, Zhang F, Zhu Y, Shi L, Li H, Xu Z (2012) POSH localizes activated Rac1 to control the formation of cytoplasmic dilation of the leading process and neuronal migration. Cell Rep 2: 640–651 [DOI] [PubMed] [Google Scholar]
  83. Yoshizaki H, Ohba Y, Kurokawa K, Itoh RE, Nakamura T, Mochizuki N, Nagashima K, Matsuda M (2003) Activity of Rho‐family GTPases during cell division as visualized with FRET‐based probes. J Cell Biol 162: 223–232 [DOI] [PMC free article] [PubMed] [Google Scholar]
  84. Zhang Y, Singh MK, Degenhardt KR, Lu MM, Bennett J, Yoshida Y, Epstein JA (2009) Tie2Cre‐mediated inactivation of plexinD1 results in congenital heart, vascular and skeletal defects. Dev Biol 325: 82–93 [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Appendix

Click here for additional data file. (21.3MB, pdf)

Movie EV1

Click here for additional data file. (1.7MB, zip)

Movie EV2

Click here for additional data file. (199.5KB, zip)

Movie EV3

Click here for additional data file. (816.8KB, zip)

Movie EV4

Click here for additional data file. (8MB, zip)

Movie EV5

Click here for additional data file. (1.5MB, zip)

Review Process File

Click here for additional data file. (456.2KB, pdf)

Articles from The EMBO Journal are provided here courtesy of Nature Publishing Group

RESOURCES