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Article |
Shootin1: a protein involved in the organization of an asymmetric signal for neuronal polarization
Correspondence to Naoyuki Inagaki: ninagaki{at}bs.naist.jp
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Neurons have the remarkable ability to polarize even in symmetrical in vitro environments. Although recent studies have shown that asymmetric intracellular signals can induce neuronal polarization, it remains unclear how these polarized signals are organized without asymmetric cues. We describe a novel protein, named shootin1, that became up-regulated during polarization of hippocampal neurons and began fluctuating accumulation among multiple neurites. Eventually, shootin1 accumulated asymmetrically in a single neurite, which led to axon induction for polarization. Disturbing the asymmetric organization of shootin1 by excess shootin1 disrupted polarization, whereas repressing shootin1 expression inhibited polarization. Overexpression and RNA interference data suggest that shootin1 is required for spatially localized phosphoinositide-3-kinase activity. Shootin1 was transported anterogradely to the growth cones and diffused back to the soma; inhibiting this transport prevented its asymmetric accumulation in neurons. We propose that shootin1 is involved in the generation of internal asymmetric signals required for neuronal polarization.
K.B. Kim's present address is Laboratory of Cell Signal Transduction, School of Life Science and Biotechnology, Korea University, Seoul, 136-701 Korea.
Abbreviations used in this paper: 2DE, 2D electrophoresis; CMFDA, 5-chloromethylfluorescein diacetate; DIV, day in vitro; E, embryonic day; miRNA, microRNA; mRFP, monomeric red fluorescent protein; P, postnatal day; PI 3-kinase, phosphoinositide-3-kinase.
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Introduction |
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Recent studies have begun to define the signaling pathways involved in neuronal polarization. Esch et al. (1999) reported that the extracellular signals laminin and neuron-glia cell adhesion molecule can specify which neurite will become an axon. As effectors of spatial signals, rearrangements of the cytoskeleton are important, as actin filament instability (Bradke and Dotti, 1999) and tubulin assembly by collapsin response mediator protein-2 (Inagaki et al., 2001; Arimura and Kaibuchi, 2005) are reported to initiate axon formation. Recent work has shown that spatially localized intracellular signaling pathways, including phosphoinositide-3-kinase (PI 3-kinase), phosphatidylinositol (3,4,5) triphosphate, the mPar3–mPar6–aPKC complex (with the exception of some neurons in Drosophila melanogaster; Rolls and Doe, 2004), Cdc42, Rap1B, STEF/Tiam1, Rac, Akt, adenomatous polyposis coli, and glycogen synthase kinase-3ß, are involved in axon specification for neuronal polarity formation (Shi et al., 2003, 2004; Menager et al., 2004; Schwamborn and Puschel, 2004; Jiang et al., 2005; Nishimura et al., 2005; Yoshimura et al., 2005), and PI 3-kinase is implicated as an upstream molecule in these events (Shi et al., 2003; Arimura and Kaibuchi, 2005; Wiggin et al., 2005).
In spite of this progress, the mechanism and logic of how the polarized distribution of intracellular signals originates in the absence of external asymmetric cues remain elusive. During the polarization of cultured hippocampal neurons, undifferentiated neurites undergo competitive elongation with each other. When one of them exceeds the others by a critical length, it rapidly elongates to become an axon (Goslin and Banker, 1989). This observation led to the proposal that a positive feedback loop and negative regulation among neurites are necessary for neuronal polarization (Goslin and Banker, 1989; Andersen and Bi, 2000; Banker, 2003). A locally acting positive feedback loop may amplify a small stochastic increase in signals until it exceeds a threshold to induce an axon, and negative regulation may also be important to prevent the formation of surplus axons. However, little is known about the molecular basis of such regulation.
To approach this problem, we performed proteome analyses of cultured hippocampal neurons using highly sensitive large- gel 2D electrophoresis (2DE), which can detect 
11,000 protein spots over a dynamic range of 1–105 (Inagaki and Katsuta, 2004). We describe a novel brain-specific protein, named shootin1. Our data suggest that shootin1 organizes its own polarized distribution to break neuronal symmetry through the PI 3-kinase pathway.
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6,200 protein spots on 2DE gels and detected 277 that were consistently up-regulated during the transition from stage 2 to 3 (n 
3). The second analysis screened proteins enriched in axons (Fig. 1 B). Hippocampi dissected from embryonic day (E) 18 rat embryos were cut into 1-mm blocks and cultured on plastic dishes, where they formed complicated networks of radial axons in 2 wk. The explants' somatodendritic parts were then separated from the axon networks, and both were compared by 2DE. By screening 5,200 protein spots, we detected 200 spots enriched in the axon samples (n 3). A total of 23 spots were detected by both screenings. Tryptic digestion and mass spectrometry of one of them, located at a molecular mass of 60 kD and pI = 5.3 in gels (Fig. 1, A and B), identified 10 peptides whose sequences corresponded to the human cDNA sequence KIAA1598 encoding a 5'-truncated ORF of 446 amino acids. A BLAST search identified four human EST clones (BI598285, BG720033, BE568283, and BI457767) and suggested that 10 additional amino acids are present in the complete ORF. We then cloned the cDNAs for the rat and human ORFs and termed them shootin1.
Rat and human shootin1 encode proteins of 456 amino acids and predicted molecular masses of 52.4 and 52.6 kD, respectively (Fig. 1 C). Domain searching revealed that shootin1 contains three coiled-coil domains and a single proline-rich region (Fig. 1 D). It does not show significant homology to previously known polypeptides, however, suggesting that it belongs to a novel class of proteins. Database searches also identified a mouse orthologue of shootin1 (Fig. 1 C) and partial ORFs in Macaca fascicularis, chick, zebrafish, and Fugu rubripes. Invertebrate homologues of shootin1 were not found in the databases. Thus, shootin1 is probably a late addition to the genome during the evolution of animals.
Shootin1 is brain specific and highly up-regulated during polarization
We raised an antibody against recombinant shootin1. It recognized a 60-kD band, corresponding to the apparent Mr of native and recombinant shootin1, in immunoblots of rat cultured hippocampal neurons (Fig. 1 E, arrowhead). Consistent with the 2DE data for the metabolically labeled protein (Fig. 1 A), the level of shootin1 expression increased remarkably during stage 2/3 transition (14.4-fold increase; n = 4; P < 0.005) and remained high until day in vitro (DIV) 14, thereafter returning to a low level by DIV28 when expression of the presynaptic protein synaptophysin increased (Fig. 1 E). Immunoblot analysis of various rat tissues detected shootin1 in postnatal day (P) 4 and adult brains but not in other tissues, suggesting that shootin1 is a brain-specific protein (Fig. 1 F). Expression of shootin1 was relatively low on E15, peaked around P4, and decreased to a low level in the adult brain (Fig. 1 G). Thus, the expression of shootin1 is up-regulated, both in hippocampal neurons and in brain, during the period of axon formation and elongation.
Shootin1 accumulates in axonal growth cones during the stage 2/3 transition
Next, we examined the localization of shootin1 in cultured hippocampal neurons. Immunocytochemical analysis showed a faint and diffuse staining of endogenous shootin1 in early stage 2 neurons (18–24 h in culture; unpublished data). In late stage 2, moderate amounts of shootin1 appeared in some growth cones of minor processes (Fig. 1 H). We used a volume marker, 5-chloromethylfluorescein diacetate (CMFDA), to measure the relative concentration of shootin1: it was calculated by using CMFDA as an internal standard (shootin1 immunoreactivity/CMFDA staining). The relative concentration of shootin1 accumulated in the growth cones of late stage 2 neurons was 2–4 times higher than that in the cell body (Fig. 1 H, arrowheads). In stage 3, shootin1 accumulated strongly in axonal growth cones (Fig. 1 I, arrows): 100% of axonal growth cones showed accumulation (n = 19). The relative concentration of shootin1 in the axonal growth cones of stage 3 neurons was 10 times higher than that in the other regions. Notably, the accumulation seen at late stage 2 in minor processes mostly disappeared in stage 3 (Fig. 1 I, arrowheads), with only 12% of the processes showing accumulation (n = 68). Shootin1 concentration in the cell body remained low throughout stages 2 and 3 (Fig. 1, H and I, asterisks). The accumulation of shootin1 in axonal growth cones was observed until stage 5 (unpublished data).
In stage 2, shootin1 shows fluctuating accumulation in multiple growth cones, concurrent with neurite elongation
To analyze the localization of shootin1 in living neurons, we monitored fluorescent images of EGFP-shootin1 expressed in hippocampal neurons under the cytomegalovirus promoter every 5 min. Although relatively high levels of EGFP-shootin1 appeared in the soma, indicating that the expression exceeds the endogenous levels, its distribution in neurites was virtually identical to that of endogenous shootin1 (see the following paragraph). Consistent with the immunocytochemical data, we observed accumulation of EGFP-shootin1 in the growth cones of minor processes in late stage 2 neurons (Fig. 2 A).
As reported previously (Goslin and Banker, 1989), minor processes showed competitive extension and retraction before polarization. Surprisingly, "hotspots" of EGFP-shootin1 accumulation repeatedly appeared and disappeared in the growth cones of individual neurites (n = 11 cells; Fig. 2 A and Video 1, available at http://www.jcb.org/cgi/content/full/jcb.200604160/DC1). Most of the neurites elongated in conjunction with EGFP-shootin1 accumulation and, conversely, retracted as EGFP-shootin1 disappeared (Fig. 2 B). To measure relative concentration of EGFP-shootin1 in growth cones, we used the volume marker monomeric red fluorescent protein (mRFP): it was calculated by using mRFP as an internal standard (EGFP-shootin1/mRFP). By quantifying EGFP-shootin1 and mRFP in growth cones and neurite elongation speed, we found a clear dose dependency of neurite elongation rate on shootin1 concentration in the growth cones of stage 2 neurons (Fig. 2 C).
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Excess levels of shootin1 disturb its asymmetric distribution and induce formation of surplus axons
To examine whether the asymmetric accumulation of shootin1 in a single neurite is important for neuronal polarization, we overexpressed EGFP-shootin1 or myc-tagged shootin1 (myc-shootin1) in hippocampal neurons under the stronger ß-actin promoter. A high level of EGFP-shootin1 was detected in the soma, with its frequent transport from the soma to growth cones (Fig. 3 A, arrowheads; and Video 3, available at http://www.jcb.org/cgi/content/full/jcb.200604160/DC1).
This in turn resulted in more continuous accumulation of EGFP-shootin1 in multiple growth cones (Fig. 3 A, arrows; compared with the dynamic fluctuation of a lower level of EGFP-shootin1 in Fig. 2, A and B, and Video 1) and ectopic accumulation of myc-shootin1 in minor process growth cones in stage 3 neurons (Fig. 3 B, arrowheads). These results suggest that the limited amount of shootin1 is essential for its asymmetric accumulation in a single neurite.
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Repressing shootin1 expression inhibits neuronal polarization
We next suppressed shootin1 expression using a vector-based RNAi system that expresses microRNA (miRNA). To ensure a high level of expression of miRNA before polarization, hippocampal neurons prepared from E18 rat embryo and transfected with the expression vector of a miRNA designated against shootin1 or a control miRNA were plated on polystyrene plates without any coating. After 20 h for the induction of the miRNA expression, the cells were collected and cultured on coverslips coated with polylysine and laminin. The shootin1 miRNA reduced the level of neuronal shootin1 (Fig. 3 F, arrows), in comparison to control neurons (arrowheads) and neurons transfected with the control miRNA. Repression of shootin1 expression by the miRNA led to significant suppression of neuronal polarization at 50 and 70 h in culture, whereas the control miRNA had no such effect (Fig. 3 G). On the other hand, 100% of neurons transfected with the shootin1 miRNA (n = 25) became polarized on DIV7. As the 20-h delay in neuronal plating might affect time course of neuronal polarization after plating, we also performed similar experiments using E17 rat embryo. Essentially equivalent data were obtained with E17 rat embryo (Fig. 3 G). The significant suppression of neuronal polarization by shootin1 RNAi provides evidence that shootin1 is involved in neuronal polarization.
Shootin1 accumulation in growth cones stimulates neurite elongation during the stage 2/3 transition
As described, shootin1 showed fluctuating accumulation in growth cones concurrent with neurite elongation in stage 2 neurons, raising the possibility that shootin1 accumulation in growth cones stimulates neurite elongation. During the stage 2/3 transition, neurites of hippocampal neurons show dynamic elongation and retraction without a remarkable increase in total neurite length (Goslin and Banker, 1989). In addition, the stage 2/3 transition is a critical period of neuronal polarization. Therefore, we examined the effect of shootin1 overexpression and RNAi during this period (24 and 48 h in culture). In contrast to the data of DIV7 (Fig. 3 E) and DIV4, shootin1 overexpression induced a significant increase in total neurite length during this period (Fig. 3 H). Furthermore, repression of its level by RNAi resulted in a significant decrease in it (Fig. 3 I). Along with the time-lapse data, these results suggest that shootin1 accumulation in growth cones stimulates neurite elongation during the transition from stage 2 to 3.
Shootin1 is anterogradely transported to the growth cones with wave-like structures and diffuses back to the soma
We next asked how shootin1 accumulates asymmetrically in hippocampal neurons. As already noted (Fig. 3 A, arrowheads), the series of time-lapse imaging revealed active transport of shootin1 from the cell body to the growth cones in stages 2 and 3 neurons (Fig. 4 A).
The shootin1 transport was observed along minor processes and axons. Ruthel and Banker (1998, 1999) reported wave-like anterograde movement of growth cone–like structures along minor processes and axons of cultured hippocampal neurons. The transport rate of these "waves" was 3 µm/min, similar to that of slow axonal transport component b, which transports actin (Lasek, et al., 1984; Brown, 2003). In addition, waves were enriched in F-actin and their movement was reversibly blocked by the actin-disrupting agent cytochalasin. Therefore, Ruthel and Banker (1998, 1999) suggested that actin and other cytoskeletal components are transported as waves from the cell body to neurite tips via an actin-dependent mechanism. Shootin1 traveled as discrete boluses with growth cone–like structures at a mean rate of 1.0 ± 0.1 µm/min (n = 12), which is similar to the speed of wave transport. We occasionally observed transient retrograde transport of GFP-shootin1. However, as in the case of the wave, retrograde transport was rare and short-lived, quickly reverting to anterograde movement. In addition, the boluses of shootin1 were enriched for F-actin (Fig. 4 B) and the transport was arrested by the actin-disrupting agent cytochalasin D within 5 min (Fig. S3 A, available at http://www.jcb.org/cgi/content/full/jcb.200604160/DC1), as reported for the waves. Blebbistatin, an inhibitor of myosin II (Straight et al., 2003), also stopped shootin1 transport (Fig. 4 C). These results suggest that shootin1 is anterogradely transported with the wave-like structure by an actin- and myosin-dependent mechanism.
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Inhibition of shootin1 transport prevents its asymmetric accumulation in neurons
We next asked whether the anterograde transport of shootin1 is involved in its asymmetric accumulation in hippocampal neurons. As shown in Fig. 5 A and Fig. S3 C, cessation of shootin1 transport in stage 2 neurons by blebbistatin or cytochalasin D prevented accumulation of shootin1 in multiple growth cones.
Stage 2 neurons were cultured for 36 h in the presence of blebbistatin or cytochalasin D. As described, in control neurons, shootin1 accumulates asymmetrically in growth cones of nascent axons during this period. On the other hand, shootin1 did not accumulate in single neurites in the presence of these drugs (Fig. 5 B and Fig. S3 D). Cessation of shootin1 transport in already polarized stage 3 neurons also prevented accumulation of shootin1 in axonal growth cones, as described (Fig. 4 D and Fig. S3 B). These data indicate that the actin- and myosin-dependent anterograde transport of shootin1 is necessary for its asymmetric accumulation in single growth cones.
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We further examined the functions of shootin1 and PI 3-kinase within the cell polarity pathways. Inhibition of PI 3-kinase activity by LY294002 led to a reduction in the percentage of neurons with multiple axons induced by shootin1 overexpression (Fig. 6 H and Fig. S4 A). On the other hand, multiple axon formation by overexpression of constitutively active PI 3-kinase was not inhibited by shootin1 RNAi (Figs. 6 H and Fig. S4 B). Collectively, these results provide evidence that shootin1 functions upstream of PI 3-kinase and is required for spatially localized PI 3-kinase activity, which is essential for neuronal polarization (Shi et al., 2003).
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Shootin1 in generation of an asymmetric signal for neuronal polarization
How does the symmetric localization of shootin1 shift to asymmetric localization before polarization? Does shootin1 spontaneously organize its own polarized distribution without asymmetric cues? Two types of regulation have been postulated for a mechanism that accounts for spontaneous neuronal polarization (Craig and Banker, 1994; Andersen and Bi, 2000). One is a positive feedback loop acting locally in neurites, where a stochastic increase in signals is enhanced until their level exceeds a threshold to induce an axon. If accumulation of a particular molecule in growth cones stimulates neurite elongation and if its accumulation increases in proportion to neurite length, such a molecule can constitute the positive feedback loop to enhance its own signal for axon formation (Goslin and Banker, 1990; Craig and Banker, 1994). Our data suggest that shootin1 accumulation in growth cones stimulates neurite elongation during the stage 2/3 transition. In addition, shootin1 may accumulate in growth cones in a neurite length–dependent manner, as it is actively transported from the cell body to growth cones and its retrograde diffusion to the cell body should vary inversely with neurite length (Goslin and Banker, 1989). Consistent with this notion, inhibiting the anterograde transport of shootin1 disturbed its asymmetric accumulation in neurons. Thus, shootin1 is a good candidate molecule for the requisite positive feedback loop for axon induction (Fig. 7 A).
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Fig. 7 C shows our current model in which shootin1 generates an asymmetric signal for neuronal polarization. In this model, shootin1 up-regulation triggers the aforementioned positive and negative regulation, which shifts the symmetry of neurons from a stable to an unstable state. Eventually, shootin1 accumulates asymmetrically in a single neurite, through amplification of its stochastic signals and competitive accumulation among neurites, thereby leading to neuronal polarization.
Shootin1 and previously described mechanisms for neuronal polarization
Recent papers indicate that PI 3-kinase is located at an upstream position in signaling pathways for neuronal polarization involving many molecules, such as phosphatidylinositol (3,4,5) triphosphate, the mPar3–mPar6–aPKC complex, Cdc42, Rap1B, STEF/Tiam1, Rac, Akt, adenomatous polyposis coli, glycogen synthase kinase-3ß, and collapsin response mediator protein-2 (Shi et al., 2003; Arimura and Kaibuchi, 2005; Wiggin et al., 2005). We found that shootin1 interacts with PI 3-kinase and is required for spatially localized PI 3-kinase activity in hippocampal neurons. Furthermore, a series of overexpression and loss of function studies suggested that shootin1 functions upstream of PI 3-kinase in regulating neuronal polarity. Thus, shootin1 may be involved in the organization of polarized PI 3-kinase activity (Fig. 7 C), which is essential for neuronal polarization (Shi et al., 2003).
Recently, Jacobson et al. (2006) showed time-lapse imaging of the motor domain of kinesin-1 in cultured hippocampal neurons. As in the case of shootin1, the kinesin-1 motor domain transiently accumulated in different minor processes of stage 2 neurons and selectively and continuously accumulated in the nascent axon during the stage 2/3 transition, thereby indicating that it serves as a very early marker for the symmetry breaking event. The present data showed that shootin1 accumulation in growth cones was induced by the actin- and myosin-dependent wave-like transport and stimulated neurite elongation. On the other hand, the accumulation of the kinesin-1 motor domain may be dependent on microtubule and was not related to neurite elongation. It is intriguing to analyze how these molecules interact during polarization.
In addition to internal signals for polarization, additional external cues are likely to adjust the orientation of an axon and dendrites in situ. Although the identities of such cues in the brain are not yet clear, Esch et al. (1999) reported that the spatially asymmetric extracellular signals of laminin and neuron-glia cell adhesion molecule can specify which neurite will become an axon under experimental conditions. The present study does not rule out the possibility that shootin1 is modified by other molecules. By regulating the activity of shootin1, additional molecules might further adjust the orientation of an axon and dendrites in situ.
In conclusion, we have identified shootin1, a novel protein involved in neuronal polarization. Based on the present findings, we proposed a model in which shootin1 expression triggers the positive and negative regulation required for neuronal symmetry breaking. Although we cannot rule out the involvement of other potential mechanisms, our data provide an insight into how internal asymmetric signals are generated during neuronal polarization.
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1 mm), carefully washed to remove dissociated cells, and cultured on polylysine- and laminin-coated plastic dishes. The explants started to extend radial axons on the dishes within 12 h. On DIV14, the radial axons formed complicated networks around the explants. The axonal networks were usually devoid of cell bodies, dendrites, and nonneuronal cells (Fig. 1 B), although we occasionally observed migration of neurons from blocks onto axonal networks. Such cells were rigorously removed under a microscope using pipette tips. Explants containing somatodendritic parts were separated from radial axons by applying streams of medium to the explants with a pipette, and the explants were then collected in microcentrifuge tubes. Removal of the explants and dissociated cells from axon networks was verified by microscopy.
For quantitative 2DE, stages 2 and 3 neurons were metabolically labeled with the culture medium containing 13% of the normal levels of methionine and cysteine plus Pro-mix L-[35S] in vitro cell labeling mix (containing 70% L-[35S]methionine and 30% L-[35S]cysteine; GE Healthcare) for 4 h. Hippocampal explants were labeled with the same medium for 24 h.
Highly sensitive gel 2DE and protein identification by mass spectrometry
2DE was performed as reported previously (Oguri et al., 2002), using a 93- x 103-cm large-gel system (Inagaki and Katsuta, 2004). For differential 2DE, neurons or explants were metabolically labeled and protein spots separated by 2DE gels were visualized by autoradiography. For protein identification, unlabeled protein samples from 2-wk-old rat brains were separated by the 2DE gel and visualized by silver staining. The protein spots corresponding to the radio-labeled ones were then excised from gels and in-gel digested as described previously (Nomura et al., 2004). Matrix-assisted laser desorption/ionization mass spectrometry was performed using a Voyager Elite equipped with delayed extraction (Applied Biosystems). Database searches were conducted using the Mascot program (Matrix Science) and National Center for Biotechnology Information databases.
Cloning of shootin1
cDNA encoding KIAA1598 was provided by T. Nagase and O. Ohara (Kazusa DNA Research Institute, Chiba, Japan). Full-length cDNA of human shootin1 was obtained by PCR of KIAA1598 with the primers 5'-GCGGATCCATGAACAGCTCGGACGAAGAGAAGCAGCTGCAGCTCATTACCAGTCTGAAG and 5'-GCGGATCCCTACTGGGAGGCCAGTATTC. cDNA encoding rat shootin1 was amplified by PCR from a rat brain cDNA library (CLONTECH Laboratories, Inc.) with the primers 5'-CCGCTCGAGATGAACAGCTCGGACGAGGAGAAG and 5'-CCGCTCGAGTTACTGGGAGGCCAGGATTCCCTTCAG. The cDNAs were then subcloned into pCMV (Stratagene), pCAGGS with a ß-actin promoter (provided by J. Miyazaki, Osaka University, Osaka, Japan; Niwa et al., 1991), pEGFP (CLONTECH Laboratories, Inc.), pGEX (GE Healthcare), and pKaede-MC1 (MBL International Corporation) vectors.
Protein and antibody preparation
Recombinant shootin1 was expressed in Escherichia coli as a GST fusion protein and purified on a glutathione–Sepharose column (GE Healthcare), after which GST was removed from shootin1 by PreScission protease (GE Healthcare). Rabbit polyclonal anti-shootin1 antibody was raised against the recombinant shootin1 and affinity purified before use.
Immunocytochemistry, immunoblot, and immunoprecipitation
Immunocytochemistry, CMFDA staining, Rhodamine phalloidin staining, and immunoblot were performed as described previously (Inagaki et al., 2001). For immunoprecipitation, P4 or P5 rat brains were extracted by addition of lysis buffer (50 mM Tris-HCl, pH 8.0, 1 mM EDTA, 150 mM NaCl, 1% Triton X-100, 0.1% SDS, 0.1% sodium deoxycholate, 2 mM phenylmethylsulfonyl fluoride, 5 µg/ml leupeptin, 10 mM NaF, 1 mM Na3VO4, and 10 mM ß-glycerophosphate) and centrifuged at 100,000 g for 30 min at 4°C. The supernatants were incubated with antibodies overnight at 4°C, and immunocomplexes were then precipitated with protein G–Sepharose 4B (GE Healthcare). After washing out beads with RIPA buffer, immunocomplexes were analyzed by immunoblot.
Microscopy
Fluorescent and phase-contrast images of neurons were acquired at room temperature using a fluorescent microscope (Axioplan 2; Carl Zeiss MicroImaging, Inc.) equipped with a Plan-NEOFLUAR 40x, 0.75 NA, or 20x, 0.50 NA, objective, a charge-coupled device camera (AxioCam MRm; Carl Zeiss MicroImaging, Inc.), and imaging software (AxioVision 3; Carl Zeiss MicroImaging, Inc.). Time-lapse microscopy was performed at 37°C using a fluorescent microscope (Axiovert S100; Carl Zeiss MicroImaging, Inc.) equipped with a Plan-NEOFLUAR 40x, 1.3 NA oil iris objective, CSNAP, and Deltavision 2 (Applied Precision) software or Axiovert 200M (Carl Zeiss MicroImaging, Inc.) equipped with a Plan-NEOFLUAR 40x, 0.75 NA objective, LSM 510 scan module (Carl Zeiss MicroImaging, Inc.), and LSM 510 META software (Carl Zeiss MicroImaging, Inc.). The acquired images were analyzed with Multi Gauge (Fujifilm) or LSM510 META software.
Transfection and RNAi
Neurons or HEK293T cells were transfected with cDNA or RNA by the calcium phosphate method (Inagaki et al., 2001), Nucleofector (Amaxa), or Lipofectamine 2000 (Invitrogen) before or after plating. For vector-based RNAi analysis, we used BLOCK-iT Pol II miR RNAi expression vector kit (Invitrogen). The targeting mRNA sequence TGAAGCTGTTAAGAAACTGGA corresponds to nucleotides 138–158 in the coding region of rat shootin1, whereas the control vector pcDNA 6.2-GW/EmGFP-miR-neg encodes an mRNA not to target any known vertebrate gene.
Materials
Antibodies against myc, tau-1, synaptophysin, MAP-2, 
-tubulin, the p85 subunit of PI 3-kinase, and monoclonal (587F11) phospho-Akt (Ser473) were obtained from MBL International Corporation, Boehringer, Progen, Sigma-Aldrich, Sigma-Aldrich, Upstate Biotechnology, and Cell Signaling Technology, respectively. CMFDA, Rhodamine phalloidin, blebbistatin, cytochalasin D, and LY294002 were obtained from Invitrogen, Invitrogen, BIOMOL Research Laboratories, Inc., Calbiochem, and Calbiochem, respectively. cDNA encoding Myr-PI 3-K p110 was obtained from Upstate Biotechnology. mRFP was provided by R. Tsien (University of California, San Diego, La Jolla, CA).
Online supplemental material
Fig. S1 shows serial time-lapse images of EGFP-shootin1 accumulation in neurites 1 and 2 of Fig. 2 (D–G). Fig. S2 shows DIV7 hippocampal neurons overexpressing myc-shootin1, which are immunostained by anti-synaptophysin or anti–MAP-2 antibody. Fig. S3 shows the effects of cytochalasin D on shootin1 distribution in hippocampal neurons. Fig. S4 shows that inhibition of PI 3-kinase activity suppresses formation of shootin1-induced multiple axons, but repression of shootin1 expression by RNAi does not inhibit formation of PI 3-kinase–induced multiple axons. Video 1 is a time-lapse video of a stage 2 hippocampal neuron expressing EGFP-shootin1 as described in Fig. 2 A. Video 2 is a time-lapse video of a hippocampal neuron expressing EGFP-shootin1 taken from stages 2 to 3 as described in Fig. 2 (D–G). Video 3 is a time-lapse video of a hippocampal neuron overexpressing EGFP-shootin1 as described in Fig. 3 A. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200604160/DC1.
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Acknowledgments |
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This research was supported in part by Precursory Research for Embryonic Science and Technology at Japanese Science and Technology Corporation; the Ministry of Education, Sports, Culture, Science and Technology (18300107); and the Japan Society for the Promotion of Science KAKENHI (18016020 and 18022028), the Suntory Institute for Bioorganic Research, and the Osaka Medical Research Foundation for Incurable Diseases.
Submitted: 26 April 2006
Accepted: 18 September 2006
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References |
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Andersen, S.S., and G.Q. Bi. 2000. Axon formation: a molecular model for the generation of neuronal polarity. Bioessays. 22:172–179.[CrossRef][Medline]
Ando, R., H. Hama, M. Yamamoto-Hino, H. Mizuno, and A. Miyawaki. 2002. An optical marker based on the UV-induced green-to-red photoconversion of a fluorescent protein. Proc. Natl. Acad. Sci. USA. 99:12651–12656.
Arimura, N., and K. Kaibuchi. 2005. Key regulators in neuronal polarity. Neuron. 48:881–884.[CrossRef][Medline]
Banker, G. 2003. Pars, PI 3-kinase, and the establishment of neuronal polarity. Cell. 112:4–5.[CrossRef][Medline]
Bradke, F., and C.G. Dotti. 1999. The role of local actin instability in axon formation. Science. 283:1931–1934.
Brown, A. 2003. Axonal transport of membranous and nonmembranous cargoes: a unified perspective. J. Cell Biol. 160:817–821.
Craig, A.M., and G. Banker. 1994. Neuronal polarity. Annu. Rev. Neurosci. 17:267–310.[CrossRef][Medline]
Da Silva, J.S., and C.G. Dotti. 2002. Breaking the neuronal sphere: regulation of the actin cytoskeleton in neuritogenesis. Nat. Rev. Neurosci. 3:694–704.[CrossRef][Medline]
Dotti, C.G., and G.A. Banker. 1987. Experimentally induced alteration in the polarity of developing neurons. Nature. 330:254–256.[CrossRef][Medline]
Dotti, C.G., C.A. Sullivan, and G.A. Banker. 1988. The establishment of polarity by hippocampal neurons in culture. J. Neurosci. 8:1454–1468.[Abstract]
Esch, T., V. Lemmon, and G. Banker. 1999. Local presentation of substrate molecules directs axon specification by cultured hippocampal neurons. J. Neurosci. 19:6417–6426.
Goslin, K., and G. Banker. 1989. Experimental observations on the development of polarity by hippocampal neurons in culture. J. Cell Biol. 108:1507–1516.
Goslin, K., and G. Banker. 1990. Rapid changes in the distribution of GAP-43 correlate with the expression of neuronal polarity during normal development and under experimental conditions. J. Cell Biol. 110:1319–1331.
Horton, A.C., and M.D. Ehlers. 2003. Neuronal polarity and trafficking. Neuron. 40:277–295.[CrossRef][Medline]
Inagaki, N., and K. Katsuta. 2004. Large gel two-dimensional electrophoresis: improving recovery of cellular proteome. Current Proteomics. 1:35–39.
Inagaki, N., K. Chihara, N. Arimura, C. Menager, Y. Kawano, N. Matsuo, T. Nishimura, M. Amano, and K. Kaibuchi. 2001. CRMP-2 induces axons in cultured hippocampal neurons. Nat. Neurosci. 4:781–782.[CrossRef][Medline]
Jacobson, C., B. Schnapp, and G.A. Banker. 2006. A change in the selective translocation of the Kinesin-1 motor domain marks the initial specification of the axon. Neuron. 49:797–804.[CrossRef][Medline]
Jiang, H., W. Guo, X. Liang, and Y. Rao. 2005. Both the establishment and the maintenance of neuronal polarity require active mechanisms: critical roles of GSK-3ß and its upstream regulators. Cell. 120:123–135.[Medline]
Lasek, R.J., J.A. Garner, and S.T. Brady. 1984. Axonal transport of the cytoplasmic matrix. J. Cell Biol. 99:212s–221s.
Menager, C., N. Arimura, Y. Fukata, and K. Kaibuchi. 2004. PIP3 is involved in neuronal polarization and axon formation. J. Neurochem. 89:109–118.[CrossRef][Medline]
Nishimura, T., T. Yamaguchi, K. Kato, M. Yoshizawa, Y. Nabeshima, S. Ohno, M. Hoshino, and K. Kaibuchi. 2005. PAR-6-PAR-3 mediates Cdc42-induced Rac activation through the Rac GEFs STEF/Tiam1. Nat. Cell Biol. 7:270–277.[CrossRef][Medline]
Niwa, H., K. Yamamura, and J. Miyazaki. 1991. Efficient selection for high-expression transfectants with a novel eukaryotic vector. Gene. 108:193–199.[CrossRef][Medline]
Nomura, E., K. Katsuta, T. Ueda, M. Toriyama, T. Mori, and N. Inagaki. 2004. Acid-labile surfactant improves in-sodium dodecyl sulfate polyacrylamide gel protein digestion for matrix-assisted laser desorption/ionization mass spectrometric peptide mapping. J. Mass Spectrom. 39:202–207.[CrossRef][Medline]
Oguri, T., I. Takahata, K. Katsuta, E. Nomura, M. Hidaka, and N. Inagaki. 2002. Proteome analysis of rat hippocampal neurons by multiple large gel two-dimensional electrophoresis. Proteomics. 2:666–672.[CrossRef][Medline]
Rolls, M.M., and C.Q. Doe. 2004. Baz, Par-6 and aPKC are not required for axon or dendrite specification in Drosophila. Nat. Neurosci. 7:1293–1295.[CrossRef][Medline]
Ruthel, G., and G. Banker. 1998. Actin-dependent anterograde movement of growth-cone-like structures along growing hippocampal axons: a novel form of axonal transport? Cell Motil. Cytoskeleton. 40:160–173.[CrossRef][Medline]
Ruthel, G., and G. Banker. 1999. Role of moving growth cone-like "wave" structures in the outgrowth of cultured hippocampal axons and dendrites. J. Neurobiol. 39:97–106.[CrossRef][Medline]
Schwamborn, J.C., and A.W. Puschel. 2004. The sequential activity of the GTPases Rap1B and Cdc42 determines neuronal polarity. Nat. Neurosci. 7:923–929.[CrossRef][Medline]
Shi, S.H., L.Y. Jan, and Y.N. Jan. 2003. Hippocampal neuronal polarity specified by spatially localized mPar3/mPar6 and PI 3-kinase activity. Cell. 112:63–75.[CrossRef][Medline]
Shi, S.H., T. Cheng, L.Y. Jan, and Y.N. Jan. 2004. APC and GSK-3beta are involved in mPar3 targeting to the nascent axon and establishment of neuronal polarity. Curr. Biol. 14:2025–2032.[CrossRef][Medline]
Straight, A.F., A. Cheung, J. Limouze, I. Chen, N.J. Westwood, J.R. Sellers, and T.J. Mitchison. 2003. Dissecting temporal and spatial control of cytokinesis with a myosin II inhibitor. Science. 299:1743–1747.
Wiggin, G.R., J.P. Fawcett, and T. Pawson. 2005. Polarity proteins in axon specification and synaptogenesis. Dev. Cell. 8:803–816.[CrossRef][Medline]
Winckler, B., and I. Mellman. 1999. Neuronal polarity: controlling the sorting and diffusion of membrane components. Neuron. 23:637–640.[CrossRef][Medline]
Yoshimura, T., Y. Kawano, N. Arimura, S. Kawabata, A. Kikuchi, and K. Kaibuchi. 2005. GSK-3ß regulates phosphorylation of CRMP-2 and neuronal polarity. Cell. 120:137–149.[CrossRef][Medline]
Yoshimura, T., N. Arimura, Y. Kawano, S. Kawabata, S. Wang, and K. Kaibuchi. 2006. Ras regulates neuronal polarity via the PI3-kinase/Akt/GSK-3ß/CRMP-2 pathway. Biochem. Biophys. Res. Commun. 340:62–68.[CrossRef][Medline]
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