CORRIGENDUM
Development 138, 385 (2011) doi:10.1242/dev.062976
© 2011. Published by The Company of Biologists Ltd
MID1 and MID2 are required for Xenopus neural tube closure through the regulation of
microtubule organization
Makoto Suzuki, Yusuke Hara, Chiyo Takagi, Takamasa S. Yamamoto and Naoto Ueno
There was an error published in Development 137, 2329-2339.
In the RT-PCR and in situ hybridization section on p. 2330, the primer pair for xMID1 was incorrect. The correct primer pair is
shown below.
5¢-AGTGTGGTTTCCTATGAGCTA-3¢ and 5¢-TGTATAATGGTTCTGTTTGAT-3¢
DEVELOPMENT
The authors apologise to readers for this mistake.
RESEARCH ARTICLE 2329
Development 137, 2329-2339 (2010) doi:10.1242/dev.048769
© 2010. Published by The Company of Biologists Ltd
MID1 and MID2 are required for Xenopus neural tube
closure through the regulation of microtubule organization
Makoto Suzuki1,2, Yusuke Hara1,2, Chiyo Takagi1, Takamasa S. Yamamoto1 and Naoto Ueno1,2,*
SUMMARY
Closure of the neural tube requires both the change and maintenance of cell shape. The change occurs mainly through two
coordinated morphogenetic events: cell elongation and apical constriction. How cytoskeletal elements, including microtubules,
are regulated in this process in vivo is largely unknown. Here, we show that neural tube closure in Xenopus depends on
orthologs of two proteins: MID1, which is responsible for Opitz G/BBB syndrome in humans, and its paralog MID2. Depletion of
the Xenopus MIDs (xMIDs) by morpholino-mediated knockdown disrupted epithelial morphology in the neural plate, leading to
neural tube defects. In the xMID-depleted neural plate, the normal epithelial organization was perturbed without affecting
neural fate. Furthermore, the xMID knockdown destabilized and caused the disorganization of microtubules, which are normally
apicobasally polarized, accounting for the abnormal phenotypes. We also found that the xMIDs and their interacting protein
Mig12 were coordinately required for microtubule stabilization during remodeling of the neural plate. Finally, we showed that
the xMIDs are required for the formation of multiple epithelial organs. We propose that similar MID-governed mechanisms
underlie the normal morphogenesis of epithelial tissues and organs, including the tissues affected in patients with Opitz G/BBB
syndrome.
INTRODUCTION
In vertebrates, the neural tube is the primary luminal structure of
early development. It is the anlage of the central nervous system
and forms from a flat neuroepithelial sheet called the neural plate.
The lateral edges of the neural plate form ridges, called neural
folds, along the dorsal surface, parallel to the anterior-posterior
axis. The neural folds continue to rise and eventually meet at the
dorsal midline, where they fuse to form the luminal structure of the
neural tube (Colas and Schoenwolf, 2001; Copp et al., 2003;
Davidson and Keller, 1999). Failure of neural tube closure causes
congenital malformations, collectively called neural tube defects
(NTDs), including anencephaly and spina bifida (Colas and
Schoenwolf, 2001; Copp et al., 2003).
During neural tube closure, the neuroepithelial cells undergo
dynamic changes in shape, including apicobasal elongation and
apical constriction, which cause the tissue to bend to form the
neural tube (Colas and Schoenwolf, 2001; Davidson and Keller,
1999). The apicobasal elongation changes the cuboidal
neuroepithelial cells into columnar cells (Burnside, 1973). Apical
constriction minimizes the apical surface of selected cells in the
neural plate (located at hinge points), causing them to adopt wedgelike rather than columnar shapes (Schoenwolf and Franks, 1984).
To achieve the complex morphological changes required for tube
formation, these cellular changes must be tightly controlled in time
and space.
1
Division of Morphogenesis, Department of Developmental Biology, National
Institute for Basic Biology, Nishigonaka 38, Myodaiji, Okazaki 444-8585, Aichi,
Japan. 2Department of Basic Biology, School of Life Science, the Graduate University
for Advanced Studies (SOKENDAI), Nishigonaka 38, Myodaiji, Okazaki 444-8585,
Aichi, Japan.
*Author for correspondence ([email protected])
Accepted 17 May 2010
It is widely accepted that regulation of the neuroepithelial
cytoskeleton is fundamental to cellular morphogenesis during
neural tube closure (Colas and Schoenwolf, 2001; Copp et al.,
2003; Pilot and Lecuit, 2005; Quintin et al., 2008). In particular,
regulation of the actin cytoskeleton has been extensively studied,
and analyses in mice, chick and Xenopus show that actin-binding
proteins and their regulators, including Shroom3, MARCKS, Rap1,
ROCKs, p190 RhoGAP and RhoA, positively regulate apical
constriction via myosin activity (Copp et al., 2003; Haigo et al.,
2003; Hildebrand, 2005; Kinoshita et al., 2008; Nishimura and
Takeichi, 2008). In addition, cell adhesion molecules such as Ncadherin and Nectin contribute to apical constriction by regulating
cortical actin assembly (Morita et al., 2010; Nandadasa et al.,
2009).
By contrast, the roles and regulatory mechanisms of
microtubules in neural tube closure have been elusive. During
cell elongation, microtubules polymerize and assemble along the
apicobasal axis (Burnside, 1973; Handel and Roth, 1971;
Karfunkel, 1971). In chick and Xenopus, microtubule
polymerization inhibitors induce aberrant cell morphologies and
defects in neural tube closure (Handel and Roth, 1971;
Karfunkel, 1971). In addition, non-centrosomal -tubulin,
indirectly recruited to the apical side by Shroom3, participates
in the assembly of microtubule arrays and apicobasal cell
elongation (Lee et al., 2007). Thus, microtubules appear to be
important in the cellular morphogenesis required for neural tube
closure.
Here, we show that the Xenopus orthologs of human MID1
(also known as FXY, RNF59, TRIM18) and of MID2, an MID1
paralog (also known as FXY2, RNF60, TRIM1) are crucial for
epithelial remodeling in neural tube closure. In humans, MID1 is
responsible for X-linked Opitz G/BBB syndrome (OS), listed as
OMIM 30000 (Buchner et al., 1999; Quaderi et al., 1997; Robin
et al., 1995). OS is characterized by midline malformations,
including hypertelorism, hypospadias, cleft lip/palate,
DEVELOPMENT
KEY WORDS: Neural tube closure, Microtubule, MID1, MID2, Opitz syndrome, Epithelial remodeling, Xenopus
2330 RESEARCH ARTICLE
MATERIALS AND METHODS
Cloning of Xenopus MID1 and MID2
Xenopus laevis MID1 was identified as a cDNA clone, XL082d10, in
our EST database (XDB3, http://xenopus.nibb.ac.jp). Since this clone
contains a 103 bp internal non-coding sequence, we isolated the entire
coding region from neurula cDNA by PCR using UTR sequence-specific
primers (5⬘-ggaattcGCACGAGGCTGGATTTTGCTTAC-3⬘ and 5⬘TGTGCATTGCAATGGATTCCCAATGGC-3⬘), and cloned it into
pCS2p+. Similarly, a partial cDNA of Xenopus laevis MID2 was
obtained by PCR from neurula cDNA using primers based on the
genomic sequence of Xenopus tropicalis (5⬘-GAATGGAACAGCCCTGTCTCATTCT-3⬘ and 5⬘-ACCTTCAAGCAATTTCTTCTCTCTG-3⬘) and cloned into pBluescript SK+. This clone contained
the 5⬘UTR and 1.1 kb of the coding region, and the last 89 bp exhibited
high homology (98.9%) with the 5⬘ region of another cDNA clone,
xlk74e03ex (deposited in NBRP Xenopus, http://www.shigen.nig.ac.jp/
xenopus/top.jsp), indicating that the entire coding region of xMID2 was
spanned by these two cDNA clones. We then isolated the entire coding
region from tailbud cDNA by PCR using UTR sequence-specific
primers (5⬘-GAATGGAACAGCCCTGTCTCATTCT-3⬘ and 5⬘-GATTTCCCATCCAAGTCCTTTGCTG-3⬘) and cloned it into pCS2p+.
Phylogenetic analysis was performed using MEGA4 software (Tamura
et al., 2007). GenBank accession numbers are GU362929 (xMID1) and
GU362930 (xMID2).
Morpholinos, plasmids and mRNA preparation
Antisense morpholino oligonucleotides (Mo) were obtained from Gene
Tools. The Mo sequences were as follows: xMID-Mo, 5⬘-CAGTTCAGACTCCAGTGTTTCCATC-3⬘; 5mis-xMID-Mo, 5⬘-CACTTGAGACTACAGTCTTTCGATC-3⬘; standard control-Mo, 5⬘-CCTCTTACCTCAGTTACAATTTATA-3⬘. The Mig12-Mo was reported previously (Hayes
et al., 2007). Each Mo was injected at 13-17 ng per blastomere unless
otherwise stated. Because neither the 5mis-xMID-Mo nor the standard
control-Mo affected normal Xenopus development, we describe these Mos
as ‘control-Mo’ in this study.
Full-length or truncated forms of xMID1, xMID2, Mig12 (Hayes et al.,
2007), human -tubulin and human tau (MAPT) (Lu and Kosik, 2001)
were subcloned into pCS2p+ with or without Venus or EGFP. For rescue
constructs, silent mutations were introduced into the Mo recognition sites
of the xMIDs. Mig12-GFP (Hayes et al., 2007) and Flag--globin
(Hemmati-Brivanlou et al., 1994; Ohkawara et al., 2003) were reported
previously. Capped mRNAs were synthesized with the mMESSAGE
mMACHINE Kit (Ambion) and purified on a NICK column (Pharmacia).
Embryo manipulation and microinjection
Capped mRNAs or Mos were injected into the appropriate region of twoor four-cell embryos. The injected embryos were cultured in 3%
Ficoll/0.1⫻ Steinberg’s Solution to stage 9, then washed and cultured in
0.3⫻ Marc’s Modified Ringer’s (MMR) until the appropriate stage
(Nieuwkoop and Faber, 1967). Morphogenetic defects in the morphants
were analyzed at stage 16-17 unless otherwise stated. In animal cap
elongation assays, 0.5 pg activin mRNA was injected into the animal pole
of two-cell embryos. The animal cap was dissected at stage 9 and cultured
in Steinberg’s Solution until the sibling embryos reached stage 17.
RT-PCR and in situ hybridization
RT-PCR and in situ hybridization were performed as described (Goda et
al., 2009). For RT-PCR with dissected tissues, the neural plate and ventral
epidermis at stage 14 were separated from the underlying mesoderm in
Danilchik’s For Amy Medium (DFA) (Sater et al., 1993). Ten explants
were used for each experiment. The following primers were used: xMID1,
5⬘-GTTGTCTTCTCTGTTGAATAA-3⬘ and 5⬘-TGTATAATGGTTCTGTTTGAT-3⬘; xMID2, 5⬘-GTCATGAAGTTAAGAAAACTTGCTC-3⬘
and 5⬘-ACCTTCAAGCAATTTCTTCTCTCTG-3⬘; NCAM, 5⬘-GCCTGTAGAATTACAATGCTG-3⬘ and 5⬘-AGCATCTTGGCTGCTGGCATT-3⬘;
Sox2, 5⬘-GAGGATGGACACTTATGCCCAC-3⬘ and 5⬘-GGACATGCTGTAGGTAGGCGA-3⬘; Epidermal keratin I, 5⬘-CGGTTGAAGGTAACCTGA-3⬘ and 5⬘-CAACCTTCCCATCAACCA-3⬘; ODC, 5⬘-CAGCTAGCTGTGGTGTGG-3⬘ and 5⬘-CAACATGGAAACTCACACC-3⬘.
The following plasmids were used for probe synthesis: xMID1 and xMID2
(constructed for this study); Sox2 (XL039o24, XDB3); NCAM (Kintner and
Melton, 1987); N-cadherin (XL289n05ex, XDB3); Epidermal keratin I
(XL056e18, XDB3); Shh (Yakushiji et al., 2007); Ptc2 (Yakushiji et al.,
2007); Gli1 (Takabatake et al., 2000); Gli3 (Takabatake et al., 2000);
HNF3 (FoxA2a) (XL016l12, XDB3); Pintallavis (FoxA4a) (XL047n03,
XDB3); N-tubulin (Takabatake et al., 2002); Shroom3 (Haigo et al., 2003);
and Pax3 (XL014p10, XDB3).
Western blotting and immunoprecipitation
For western blotting to test the specificity of xMID-Mo, 20 embryos at
stage 14 were lysed in 400 l lysis buffer [50 mM Tris-HCl (pH 7.5), 150
mM NaCl, 5 mM EDTA, 0.5% NP40, 50 mM NaF, protease inhibitors].
For immunoprecipitation of EGFP-tubulin, 30 embryos at the late neurula
stage were lysed in 600 l lysis buffer. Immunoprecipitation was
performed as described (Ohkawara et al., 2003). Antibodies to GFP (598,
MBL) and acetylated tubulin (TT6793, Sigma) were used.
Immunohistochemistry
Embryos were fixed in Dent’s Fixative (for -catenin, C-cadherin, ZO-1),
low-FGT Fixative (for -tubulin and cytoplasmic acetylated tubulin)
(Becker and Gard, 2006) or MEMFA. Published procedures were used to
prepare (Fagotto and Gumbiner, 1994) and stain (Suzuki et al., 2007) fish
gelatin cryosections, or thick sections (Becker and Gard, 2006), with minor
modifications. Antibodies to the following proteins were used: Sox2
(AB5603, Chemicon); phospho-histone H3 (06-570, Upstate); active
caspase 3 (559565, BD Pharmingen); laminin (L9393, Sigma); Xen1
(DSHB); MZ15 (DSHB); -catenin (C7082, Sigma); C-cadherin (6B6,
DSHB); ZO-1 (AB01003, Sanko Junyaku); -tubulin (T9026, Sigma);
acetylated tubulin (T6793, Sigma); Flag (F3165 and F7425, Sigma); GFP
(598, MBL); and RFP (PM005, MBL). The secondary antibodies were
anti-mouse HRP (170-6516, Bio-Rad); anti-mouse Alexa 488 (A11017,
Molecular probes); anti-mouse Alexa 555 (A21425, Molecular probes);
anti-rabbit Alexa 488 (A11070, Molecular probes); anti-rabbit Alexa 555
(A21430, Molecular probes); and anti-rabbit Cy5 (111-176-047, Jackson).
When necessary, sections were counterstained with Alexa 546-phalloidin
(A22283, Molecular probes) or TO-PRO-3 (T3605, Molecular probes) to
label F-actin or nuclei, respectively.
Imaging and image analysis
For rescue experiments, we co-injected 0.25% Rhodamine-Dextran (D1817,
Molecular Probes) with the Mo, selected embryos with appropriate
fluorescence in the neural plate, and fixed them in MEMFA when the control
embryos reached the neural-fold stage. For quantitative analysis, the distance
DEVELOPMENT
laryngotracheoesophageal abnormalities, imperforate anus, cardiac
defects and brain abnormalities (Fontanella et al., 2008; So et al.,
2005).
MID1 and MID2 encode conserved proteins associated with
microtubules belonging to the RBCC/TRIM (N-terminal RING
finger-B box-coiled coil/tripartite motif) superfamily (Buchner
et al., 1999; Cainarca et al., 1999; Schweiger et al., 1999; Short
and Cox, 2006). MID1 and MID2 are known to be expressed
during development in human, mouse and chick (Buchner et al.,
1999; Dal Zotto et al., 1998; Granata et al., 2005; Pinson et al.,
2004; Quaderi et al., 1997; Richman et al., 2002). However,
although biochemical and in vitro cell biological studies have
yielded some information, the physiological and developmental
functions of the MID proteins are still unclear, as is the
pathological role of the MID1 mutant in OS. We report here that
Xenopus MID1 and MID2 (xMID1 and xMID2) are essential for
neural tube closure through their stabilization of microtubules,
which is required for cell elongation and apical constriction. We
propose that microtubule regulation by the MIDs is crucial for
a variety of epithelial remodeling processes during the
development of many vertebrate species.
Development 137 (14)
Microtubule and xMIDs in neurulation
RESEARCH ARTICLE 2331
between the neural folds marked by pigmented margins in the anterior spinal
cord anlage was measured. To quantify the cellular characteristics, images of
stained sections were obtained with a Zeiss LSM510 META confocal
microscope equipped with a 63⫻, NA 1.4, oil-immersion or a 40⫻, NA 1.2,
water-immersion objective lens. The cell height, apical width, basal width,
perimeter of the apical side, and pixel intensities were all determined using
ImageJ (NIH) software (see Fig. S4E in the supplementary material). To
quantify the cell morphologies, we selected cells with easily detected tracer
fluorescence and a visible nucleus. The apical width and perimeter were
defined as the distance and outline length between apical cell-cell junctions,
respectively. The cell height was defined as the maximal length along the
axis perpendicular to the apical width. Similarly, the basal width was defined
as the distance between the basal cell-cell junction with the neighboring cell
and the line perpendicular to the apical width in contact with the opposite
basal junction. Cell width was defined as the larger of the apical and basal
widths (see Fig. S4E in the supplementary material).
To quantify molecular markers, labeled areas in the apical cell junctions
(ZO-1, C-cadherin, -catenin) or the basal lamina (laminin) were
manually selected to exclude background staining. For apical cell
junctions, we selected the region with higher fluorescence intensity than
detected more basally, and excluded almost all of the cell membrane with
baseline-level staining and the cytoplasm, because we could not clearly
distinguish between low-level specific signals and non-specific
background labeling. Then, the total pixel intensity in each selected area
was measured. In the case of laminin, we drew a thin (one-pixel-wide) line
along the basal end of the cell, at the level of the basal lamina, where the
laminin fluorescence was brightest, and measured the fluorescence
intensity on the line. Since the basal width varied greatly among cells, we
normalized the data by dividing the total intensity by the length of the line.
Data were analyzed by Student’s t-test, and are presented as the mean ±
s.e.m.
RESULTS
Identification of the Xenopus homologs of MID1
and MID2
To identify candidate molecules for regulating neural tube closure,
we focused on MIDs that were implicated in epithelial
morphogenesis through microtubule regulation. We isolated two
cDNAs that encode proteins of 668 and 687 amino acids and
exhibit 92% identity to human MID1 and 83% to human MID2,
respectively (see Fig. S1A in the supplementary material). A
phylogenetic analysis confirmed that the two genes were closest to
human MID1 and MID2 (see Fig. S1B in supplementary material)
and we therefore named them xMID1 and xMID2.
By reverse transcription PCR (RT-PCR), we found that both
xMIDs were expressed throughout embryogenesis (Fig. 1A). In
addition, both genes were expressed in the neural plate of the early
neurula (Fig. 1B). Next, we analyzed the spatial expression of the
xMIDs in detail by in situ hybridization. Before neurulation, neither
gene was detectable (Fig. 1C; data not shown). During early to midneurulation, xMID1 was upregulated uniformly in the embryo (Fig.
1D), and by late neurulation its transcripts were detected in the
epithelial organs, including the neural tube, optic and otic vesicles,
cement gland and newly epithelialized somites (Fig. 1E,F). At the
tailbud stages, additional tissues expressed xMID1, including the
midbrain, hindbrain, pronephros, pharyngeal pouch, heart tube and
scattered epidermal cells (Fig. 1H,I). By contrast, expression of
xMID2 was undetectable at neurula stages (data not shown),
whereas weak expression was observed in the pineal gland, otic
vesicle and heart tube at the tailbud stages (Fig. 1J).
DEVELOPMENT
Fig. 1. Expression patterns of xMID1 and xMID2 through early embryogenesis. (A,B)RT-PCR analysis. (A)xMID1 and xMID2 mRNAs are
expressed maternally and zygotically. Ornithine decarboxylase (ODC) was used as an internal control. The number above each lane is the embryonic
stage. –RT, control experiment without reverse transcriptase. (B)Ectodermal expression of xMIDs in the early neurula (stage 14) Xenopus embryo. In
the neural plate, xMID1 and xMID2 were expressed at similar levels to in ventral epidermis (Epidermis) and whole embryo (W.E.). NCAM and Sox2
are pan-neural markers. Epidermal keratin I (Epi. keratin) is an epidermal marker. (C-I)Expression pattern of xMID1 in embryos at the stages
indicated (lower right in each panel). (C)Dorso-posterior view, dorsal to the top. (D,E)Dorsal view, anterior to the top. (F)Anterior view, dorsal to
the top. (G)Dorsal view, labeled with sense probe (S), anterior to the top. (H,I)Lateral view, anterior to the left, dorsal to the top. (J,K)xMID2
expression. Lateral view, anterior to the left, dorsal to the top. (K)Sense probe (S). cg, cement gland; hb, hindbrain; ht, heart tube; mb, midbrain;
nt, neural tube; op, optic vesicle; ot, otic vesicle; pg, pineal gland; pn, pronephros; pp, pharyngeal pouch; pr, proctodeum; so, somite.
2332 RESEARCH ARTICLE
Development 137 (14)
Fig. 2. Depletion of xMIDs causes neural tube defects. (A)The
complementary sequence of the xMID-Mo compared with xMID1 and
xMID2. The ATG codons for the first methionine are underlined.
(B)Western blot analysis of the N-terminal domain of the xMID
proteins, tagged with Venus at the C-terminus (xMID1-Vns, xMID2-Vns,
250 pg), probed with an anti-GFP antibody. The xMID-Mo (17 ng) did
not block translation of the mRNA for Venus (Vns, 100 pg) or the
mRNA for xMID1-Venus with six silent mutations in the Mo recognition
site (mut-xMID1-Vns, 250 pg). (C,D)Dorsal views of the unilaterally
injected morphants; anterior is to the top. Control-Mo (C) or xMID-Mo
(D) was injected into the right dorsal blastomere at the four-cell stage.
Dashed lines indicate the boundaries between the neural and nonneural ectoderm. The black bracket marked by an asterisk indicates the
distance between the neural folds in a rescue experiment. (E)Dorsal
view of bilaterally injected xMID morphants at stage 20; anterior is to
the top. (E⬘)Higher magnification view of the area marked by the
bracket in E. (F,G)Transverse sections through the neural plate of
Xenopus embryos with unilateral injection of control-Mo (F) or xMIDMo (G). Dashed lines indicate the outlines of neural tissues, and
brackets show the distance between the neural folds in a rescue
experiment. (H)Average distance between the neural folds of
unilaterally injected xMID morphants were dose-dependently reduced
by the Mo-insensitive xMID1 (50, 100, 200, 500 pg), xMID2 (50, 100,
200, 500 pg), or xMID1+2 (25, 50, 100, 250 pg each) mRNAs. The
number of embryos examined is indicated above each bar.
Knockdown of xMIDs causes neural tube defects
To deplete the endogenous xMID proteins and elucidate their in vivo
role, we designed a specific antisense morpholino oligonucleotide
(xMID-Mo) that efficiently blocked the translation of not only 5⬘
UTR-xMID1-Venus, but also 5⬘ UTR-xMID2-Venus, owing to the
similarity of the Mo recognition site (Fig. 2A,B). The xMID-Mo did
not reduce the protein level of a 5⬘ UTR-xMID1-Venus with six silent
mutations in the target sequence (mut-xMID1-Vns).
The xMID-Mo, injected into one dorsal blastomere of four-cell
embryos, caused a marked delay in neural tube closure (Fig. 2D,G),
whereas the control-Mo had no effect on embryonic development
(Fig. 2C,F). In the xMID morphants, the distance between neural
folds was significantly greater (by ~3- to 5-fold) than in control
sibling embryos (Fig. 2H). Furthermore, the bilateral injection of the
xMID-Mo caused severe defects in which the neural tube remained
open even at the late neurula stage (Fig. 2E). In addition, xMID
morphant cells were consistently found as a dissociated clump of
Knockdown of xMIDs does not affect gastrulation
movements, cell viability, neural development or
primary ciliogenesis
To investigate whether the effects of xMID-Mo were specific to
epithelial remodeling, we examined gastrulation movement, cell
viability and neural specification and patterning. xMID-Mo did not
affect gastrulation movement, activin-induced animal cap
elongation or cell proliferation and viability as revealed by staining
for phospho-histone H3 and the apoptosis marker active caspase 3
(see Fig. S2 in the supplementary material).
Since MID1 and MID2 repress Shh in Hensen’s node in chicken
(Granata and Quaderi, 2003; Granata et al., 2005), and a deficiency
in Shh activity inhibits neural plate bending (Ybot-Gonzalez et al.,
2002), we examined neural specification, dorsoventral patterning
and the Shh pathway in the xMID morphants. We detected no
apparent change in the markers tested, except that the delayed
neural tube closure resulted in expression domains that were wider
than normal at these stages (see Fig. S2E and Fig. S3 in the
supplementary material). From these results, we concluded that the
loss of xMID function did not affect gastrulation, cell viability or
neural specification and patterning.
Recent studies have shown that mutations in ciliary genes that
result in agenesis of the primary cilium, a microtubule-based
organelle, cause NTDs, indicating some linkage between primary
cilium formation and neural tube closure (Bisgrove and Yost, 2006).
Hence, we analyzed the genesis of the primary cilia in the neural
tube. Knockdown of the xMIDs did not obviously affect the length
DEVELOPMENT
cells at the anterior neural plate [96% (n73 embryos) compared
with 0.02% (n95 embryos) in control morphants] (Fig. 2E⬘). Coinjection of the xMID-Mo with the mRNAs of rescue constructs with
silent mutations in the Mo recognition site of xMID1, xMID2, or
xMID1+2 dose-dependently reversed the Mo-induced distance
between the neural folds (Fig. 2H). Therefore, the knockdown of the
xMIDs by the xMID-Mo specifically caused the NTDs.
Microtubule and xMIDs in neurulation
RESEARCH ARTICLE 2333
Fig. 3. xMIDs regulate epithelial
morphology and organization.
(A-A⬙) Transverse section through the neural
plate of unilaterally injected xMID morphants
stained with phalloidin. (A)Apical actin
assembly in the morphant cells (white
arrowheads) was attenuated compared with
that on the control side (open arrowheads).
(A⬘)EGFP mRNA (50 pg) was co-injected as a
tracer. Dashed line indicates Mo-injected cells.
(A⬙)Schematic illustration of A,A⬘ showing the
non-Mo-containing cells outlined in blue and
morphant cells in pink. (B-G)Quantification of
cell morphological features in the superficial
layer of the neural plate. Control-Mo (Cont.Mo), n53 cells (8 embryos); xMID-Mo, n59
cells (7 embryos); xMID-Mo + xMID1, n56
cells (13 embryos); xMID-Mo + xMID2, n54
cells (9 embryos); xMID-Mo + xMID1&2, n42
cells (10 embryos). *P<0.05, **P<0.01,
***P<0.001, compared with xMID-Mo.
Knockdown of xMIDs induces aberrant cell
morphology in the neural plate
Next, we analyzed the epithelial cell morphology in the xMID
morphants by phalloidin staining. On the control side, the
neuroepithelial cells showed normal apicobasal elongation and
apical constriction (Fig. 3A,A⬙, blue outlines). In striking contrast,
the xMID morphant cells did not elongate, but remained rounded,
and their apical constriction was perturbed (Fig. 3A-A⬙, pink
outlines; see also Fig. S4A-D in the supplementary material).
Consistent with this, the cortical actin, the assembly of which is a
prominent feature of apical constriction, was clearly attenuated
(Fig. 3A, arrowheads).
To quantify the morphological defects in the xMID morphants
(see Fig. S4E in the supplementary material), we analyzed the
neuroepithelial cells in the superficial and deep layers separately
(Schroeder, 1970). In the xMID-depleted superficial layer, the cell
height and apical width were significantly decreased and increased,
respectively, compared with the control (Fig. 3B-D). Consequently,
the ratios of cell height to width and apical width to basal width
were altered (Fig. 3E,F). We observed similar defects in the deep
layer, except that the basal width of the deep morphant cells was
significantly increased (see Fig. S4F-J in the supplementary
material). In addition, by assessing the ratio of the apical perimeter
to the cell width, we found that the normally flat apical surface of
the cell tended to protrude in the xMID morphants (Fig. 3G). Thus,
the knockdown of the xMIDs caused cellular defects in both the
superficial and deep layers of the neural plate.
Consistent with the rescue data described above (Fig. 2H), the
xMID1 and xMID2 mRNAs partially, but convincingly, rescued the
apical width (Fig. 3C,F) and the ratio of the apical perimeter to the
cell width (Fig. 3G). Although co-expression of the xMIDs did not
rescue the cell height, it significantly decreased the basal width
compared with that of control cells (Fig. 3B,D), which resulted in
an increased height-to-width ratio (Fig. 3E). Therefore, the
exogenous xMIDs rescued the cell morphology from a rounded to
a columnar shape. These data strongly suggest that the NTDs of the
xMID morphants were due to defects in cellular morphogenesis in
the neural plate.
Defects in cell-cell and cell-extracellular matrix
contacts in the neural plate
To dissect the cellular phenotype of the xMID morphants, we
examined the localization of proteins involved in cell-cell and cellextracellular matrix (ECM) contacts. In the control cells, ZO-1
(also known as TJP1), a tight junction marker (Itoh et al., 1993),
and C-cadherin and -catenin, the major components of the
cadherin complex (Brieher and Gumbiner, 1994; Levine et al.,
1994), were concentrated at the apical junction (Fig. 4A,C,E). By
contrast, at the apical junction in the xMID morphant cells, the ZO1 signal was obscure (Fig. 4B,I) and the levels of C-cadherin and
-catenin were severely reduced (Fig. 4D,F,J,K), although the
transcription and translation of these molecules were unaffected
(data not shown). We also examined laminin, a major component
of the basal lamina (Miner et al., 1998), and found that its
localization in the basal lamina was attenuated in the xMID
morphants (Fig. 4G,H,H⬘,L). Thus, the knockdown of xMIDs
resulted in the aberrant organization of cell-adhesive machineries
and the polarized distribution of the ECM.
Defective microtubule organization and
stabilization in xMID morphants
We next analyzed the subcellular localization of EGFP-tagged
xMID1. Interestingly, EGFP-xMID1 colocalized with bundles of
non-centrosomal microtubules stained with anti--tubulin
antibody (Fig. 5A), which are readily observed in apicobasally
elongated epithelial cells (Bacallao et al., 1989; Bartolini and
Gundersen, 2006; Lee et al., 2007). In the control columnar
epithelial cells, the apicobasal arrays of microtubules were also
readily apparent (100%, n11 cells, three embryos) (Fig. 5B). By
contrast, in the xMID morphant cells, the arrays of microtubules
DEVELOPMENT
or number of primary cilia (see Fig. S2F in the supplementary
material). Therefore, the NTDs of the xMID morphants are not
attributable to a defect in primary cilium formation.
2334 RESEARCH ARTICLE
Development 137 (14)
Fig. 4. xMIDs regulate the localization of adhesive molecules. (A-H⬘) Transverse sections through the neural plate of unilaterally control-Moinjected (Cont.-Mo) (A,C,E,G) and xMID-Mo-injected (B,D,F,H) Xenopus embryos at stage 15.5, stained with antibodies against ZO-1 (A,B), Ccadherin (C-cad.) (C,D), -catenin (-cat.) (E,F), or laminin (G,H). Flag--globin mRNA (250 pg) was co-injected as a tracer, and stained with an antiFlag antibody (magenta). (A-F)Dashed lines indicate Flag-positive Mo-injected cells. (H)Arrowheads indicate the attenuation of laminin
accumulation basal to the xMID-Mo-injected cells. (H⬘)Higher-magnification view of the boxed area in H. Scale bars: 50m. (I-L)Quantification of
marker intensities in the morphants at stage 15.5. For ZO-1 intensity: control-Mo, n27 sites (3 embryos); xMID-Mo, n44 sites (6 embryos). For Ccadherin: control-Mo, n16 sites (4 embryos); xMID-Mo, n20 sites (5 embryos). For -catenin: control-Mo, n7 sites (3 embryos); xMID-Mo, n15
sites (4 embryos). For laminin: control-Mo, n19 sites (4 embryos); xMID-Mo, n19 sites (3 embryos). *P<0.05, **P<0.001. (M,N)Schematic
illustrations showing the control (M) and xMID (N) morphants. Rectangles indicate the regions analyzed in this study. so, somite; nt, notochord.
tau-injected xMID morphant cells (see Fig. S5E,F in the
supplementary material). These findings suggest that xMIDs are
required for the stabilization of microtubules.
xMIDs functionally interact with Mig12 in neural
tube closure
Mig12 (also known as G12-like and MID1IP1), which encodes a
MID1-interacting molecule, is expressed in the ventral midline of
the neural plate (Berti et al., 2004; Conway, 1995; Hayes et al.,
2007). The Mo-mediated knockdown of Mig12 causes NTDs (Fig.
6A-C) (Hayes et al., 2007) and defects in epidermal ciliogenesis
(Hayes et al., 2007). To investigate the functional interaction of
xMIDs and Mig12, we performed individual injections or coinjections of xMID-Mo and Mig12-Mo. In morphants that received
either Mo alone at a low dose, only a slight delay in neural tube
closure was induced (Fig. 6D-F,H). By contrast, the co-injection of
xMID-Mo and Mig12-Mo at the same low dose induced severe
NTDs (Fig. 6G,H), suggesting that xMIDs and Mig12 interact
functionally. We then performed rescue experiments of xMID
morphants with Mig12 and vice versa. The NTDs of the Mig12
morphants were rescued by xMID1 mRNA in a dose-dependent
manner (Fig. 6I). However, the NTDs in the xMID morphants were
not rescued by Mig12 mRNA (Fig. 6J), indicating that Mig12
requires the xMIDs to function in neural tube closure.
DEVELOPMENT
were not polarized, and the cells were more rounded (85%, n13
cells, four embryos) (Fig. 5D). To assess the stability of the
polymerized microtubules, we analyzed their acetylation status
(Creppe et al., 2009; Verhey and Gaertig, 2007). In the control
cells, filamentous and continuous staining was detected,
particularly in the apical region (86%, n7 cells, two embryos)
(Fig. 5C). By contrast, in the rounded xMID morphant cells the
acetylated -tubulin staining was punctate (92%, n13 cells, four
embryos) (Fig. 5E). Furthermore, the acetylation of the
overexpressed EGFP-tubulin was markedly decreased in the
xMID morphants (40±4.5%, P<0.01, n3) (Fig. 5F,G). Thus, the
microtubules of the xMID morphant cells were disorganized and
destabilized.
We next examined the relationship between the microtubule
destabilization and the NTDs in the morphant embryos by coinjecting the xMID-Mo with the mRNA for an unrelated
microtubule-stabilizing factor, tau (also known as MAPT), which
is a classical microtubule-associated protein (Kanai et al., 1992;
Lu and Kosik, 2001). The forced expression of tau not only
rescued the disrupted neural cell morphologies (see Fig.
S5C,D,G-L in the supplementary material), but also partially
rescued the NTDs of the xMID morphants (see Fig. S5A,B in
the supplementary material). Furthermore, the apicobasal
polarization of the microtubule arrays was restored in the
Microtubule and xMIDs in neurulation
RESEARCH ARTICLE 2335
Fig. 5. xMIDs associate with and regulate microtubules.
(A)Transverse section through the neural plate of a Xenopus embryo
injected with EGFP-xMID1 mRNA (50 pg) at stage 15.5, and stained with
anti-GFP and anti--tubulin antibodies. Scale bar: 20m. (B-E)Transverse
sections through the neural plate of embryos unilaterally injected with
control-Mo (B,C) or xMID-Mo (D,E), and stained for -tubulin (B,D) or
acetylated tubulin (Ac.-Tubulin) (C,E) antibodies. Flag--globin mRNA
(250 pg) was co-injected as a tracer, and stained with an anti-Flag
antibody (middle panels). Bottom panels show traced drawings of cells
stained by -tubulin and anti-acetylated tubulin antibodies. (F)Western
blots of immunoprecipitates (IP) or lysates from embryos expressing EGFPtubulin (EGFP-tub.) mRNA (1 ng), detected with anti-GFP and antiacetylated tubulin (Acetylated-tub.) antibodies. Immunoprecipitation was
performed using the anti-GFP antibody. (G)Quantification of
immunoprecipitation assay, showing the signal intensities of acetylated
tubulin from three independent experiments, normalized to those of
EGFP-tubulin. Error bars indicate s.e.m. *P<0.01.
The neuroepithelial cells of the Mig12 morphants and
xMID/Mig12 double morphants resembled those of the xMID
morphants (Fig. 6K-M). Consistent with this, the assembly of
apicobasally polarized microtubules was decreased in the double
xMIDs contribute to the development of other
epithelial organs
To further investigate the role of xMIDs in epithelial morphogenesis,
we performed targeted Mo injections into the presumptive head
region of embryos and found that xMID-Mo caused developmental
defects of the eye (data not shown). Characterization using a neural
marker, Xen1, and a notochord marker, MZ15, revealed hypoplasia
of the anterior central nervous system in the xMID morphants,
although the notochord formed normally (Fig. 7A,B, insets; data not
shown). We also analyzed neuroepithelial cell morphologies and
laminin localization in the basal lamina at the tailbud stage. The
normally multi-layered structures of the brain and optic vesicles (Fig.
7A,C) were disorganized, and the neuroepithelial cells had not
elongated (Fig. 7B,D). Furthermore, neuroepithelial cells had
dissociated from the apical surface of the epithelial sheet, which
lacked actin filaments (Fig. 7L; data not shown). The neuroepithelial
cells found in the ventricle were positive for active caspase 3 (Fig.
7K,L, dashed line), suggesting anoikis, a form of apoptosis caused
by the loss of cell adhesion (Frisch and Screaton, 2001). Moreover,
a continuous basal ECM failed to form, as indicated by the noncontinuous and attenuated laminin staining (Fig. 7B,D, arrowheads).
Thus, the NTDs that develop in xMID morphants ultimately cause
the catastrophic collapse of the central nervous system.
Similar defects in cell morphogenesis were found in the cement
gland (Fig. 7E,F), where xMID1 is strongly expressed (Fig. 1F,H,I).
Furthermore, in the pronephros, the epithelial cells failed to adopt a
columnar shape or exhibit apical actin assembly, and no tubular
structure was formed (Fig. 7G,H). The area in which the pronephros
normally forms was filled with disorganized cell aggregates (Fig.
7H). In the foregut, derivatives of which are affected in OS patients
(Fontanella et al., 2008; So et al., 2005), continuous apical actin
failed to assemble in the endodermal cells, and the cells protruded
apically, which caused a deformity of the luminal structure (Fig.
7I,J). Taken together, our findings indicate that xMIDs play a
fundamental role in the remodeling of multiple epithelial tissues.
DISCUSSION
Role of microtubule regulation by xMIDs in cell
shape changes and maintenance during neural
tube closure
Here, we demonstrated that the xMIDs are required for normal
neural tube closure, a multi-step event that involves neural
specification, cell proliferation and morphogenetic movements
DEVELOPMENT
morphants (Fig. 6K-M, bottom). These results highlight the
functional relationship between the xMIDs and Mig12 in regulating
microtubule organization and cellular morphogenesis.
In vitro, when co-expressed with MID1, Mig12 colocalizes with
microtubules and stabilizes them (Berti et al., 2004). To investigate
the subcellular localization of Mig12, we expressed Mig12-GFP
with xMID1 in animal caps and the neural plate (see Fig. S6 in the
supplementary material). When expressed alone, Mig12 was
distributed throughout the cells of the animal cap and neural plate
(see Fig. S6A,C,D in the supplementary material). However, when
co-expressed with xMID1, Mig12-GFP colocalized with the
microtubules in the animal cap cells, although such colocalization
was not evident in neuroepithelial cells (see Fig. S6B,E in the
supplementary material). Thus, the functional interaction of the
xMIDs and Mig12 appears to be highly dynamic and contextdependent, and in the neuroepithelial cells controlled physical and
functional interactions allow highly organized microtubule
remodeling.
2336 RESEARCH ARTICLE
Development 137 (14)
(Copp et al., 2003). In particular, a collective cell movement, which
is based on the morphogenesis of cells in the neural plate, serves
as the major driving force for its invagination (Colas and
Schoenwolf, 2001; Pilot and Lecuit, 2005; Quintin et al., 2008). In
xMID morphants, the neuroepithelial cells remained rounded, and
the localization of adhesive molecules was perturbed, indicating
that the epithelial organization was not maintained. The
rearrangement and assembly of microtubules was also impaired in
the xMID morphants. The prevention of NTDs in xMID morphants
by the expression of another microtubule-associated protein
suggests that the primary function of xMIDs is to stabilize
microtubules. From these data, we propose that the xMIDs regulate
cellular morphogenesis and epithelial organization during neural
tube closure through the assembly and stabilization of
microtubules. Since the knockdown of the xMIDs did not cause
any obvious defects in classical microtubule function in mitosis or
primary cilium formation, the impact of xMID knockdown on
microtubules might be limited, affecting only their rearrangement
and assembly along the apicobasal axis.
Since the cellular and molecular mechanisms of neural tube
closure in amphibians are closely related to those in amniotes
(Davidson and Keller, 1999), MIDs are probably required for
neural tube closure in amniotes, including humans. However,
NTDs, such as anencephaly and spina bifida, have not been
reported in OS patients, although the expression of human MID1
in the developing neural tube has been reported (Pinson et al.,
2004), and abnormalities of the brain, including agenesis or
hypoplasia of the cerebellar vermis and corpus callosum, are seen
in OS (Fontanella et al., 2008; So et al., 2005). The overlapping
expression of MID1 and MID2 in developing neural tissues
(Buchner et al., 1999; Granata et al., 2005; Dal Zotto et al., 1998)
and the finding that MID1 and MID2 have redundant activities in
DEVELOPMENT
Fig. 6. Mig12 functions with xMIDs in
neural tube closure. (A,B)Dorsal views of
Xenopus embryos given bilateral injections of
control-Mo (A) or Mig12-Mo (B); anterior is to
the top. (C)Average distance between the
neural folds of unilaterally injected Mig12
morphants was reduced by the Mo-insensitive
Mig12 mRNA (25, 50 pg). (D-H)Synergistic
effects of Mig12-Mo and xMID-Mo on neural
tube closure. Dorsal views, anterior to the
top. Injection of low doses (8.4 ng) of Mig12Mo (E) or xMID-Mo (F) caused slight delays in
neural tube closure. Co-injection of Mig12Mo and xMID-Mo (8.4 ng each) induced
severe NTDs (G,H). (I)Average distance
between the neural folds of unilaterally
injected Mig-12 morphants was reduced
dose-dependently by xMID1 mRNA (50, 100,
250, 500, 1000 pg). (J)Average distance
between the neural folds of unilaterally
injected xMID morphants was not reduced by
Mig12 mRNA (25, 50, 100, 250, 500 pg). The
number of embryos examined in C,H,I,J is
indicated above each bar. (K-M)Transverse
sections through the neural plate of embryos
given unilateral injections of control-Mo (K,
8.4 ng), Mig12-Mo (L, 17 ng), or Mig12-Mo
and xMID-Mo (M, 8.4 ng each), stained with
-tubulin (top) and anti-GFP (middle)
antibodies. mRNA (125 pg) encoding
membrane-bound EGFP was co-injected as a
tracer. Bottom panels show traced drawings
of cells stained by anti--tubulin.
Microtubule and xMIDs in neurulation
RESEARCH ARTICLE 2337
chick left-right determination (Granata et al., 2005), suggest that
MID1 and MID2 have redundant functions in neural tube closure
such that their role in this process is not unveiled by the
knockdown or mutation of only one of them.
xMID-Mig12 collaboration is required for the
rearrangement and stabilization of microtubules
in vivo
Mig12 was identified as a gene encoding a 152 amino acid protein
that is expressed in gastrula-stage zebrafish (Conway, 1995). In
Cos7 cells, Mig12 colocalizes with MID1 and stabilizes
microtubules, suggesting that Mig12 might function cooperatively
with the xMIDs. In this study, we showed that Mig12 cooperates
functionally with the xMIDs in regulating microtubule organization
during neural tube closure. However, all our data support the idea
that the xMIDs function as the dominant regulators of neural tube
closure. The strongest evidence for this is that the NTDs of Mig12
morphants were rescued by xMID expression, whereas those of
xMID morphants were not rescued by Mig12. Furthermore, the
cytoplasmic localization of Mig12-GFP was not changed by the
gain or loss of xMID function. These data all suggest that, at least
in the neuroepithelial cells, the xMIDs are the main players in the
xMID-Mig12 complex, and Mig12 might be recruited in a limited
amount to finely modulate the xMID activities. Furthermore, the
functional interaction of these proteins might be dynamic and
tightly regulated in time and space to avoid overstabilization of
the microtubules, which might lead to defects in cellular
morphogenesis. Since a regulatory subunit of protein phosphatase
2A (PP2A) binds MID1 and MID2 (Liu et al., 2001; Short et al.,
2002; Trockenbacher et al., 2001), it is possible that the PP2A
complex is involved in this mechanism.
Molecular link between microtubules and apical
constriction
The molecular mechanisms governing cellular morphogenesis in
epithelia are well documented, especially with regard to apical
constriction in Drosophila (Dawes-Hoang et al., 2005; Kolsch et
al., 2007; Nikolaidou and Barrett, 2004; Pilot and Lecuit, 2005;
DEVELOPMENT
Fig. 7. xMIDs function in the developing brain
and other epithelial organogeneses.
(A-J)Transverse sections through the brain (A,B), optic
vesicle (C,D), cement gland (E,F), pronephros (G,H)
and foregut (I,J) of Xenopus embryos injected with
control-Mo (A,C,E,G,I) or xMID-Mo (B,D,F,H,J), and
stained with phalloidin (green), anti-laminin antibody
(magenta), or anti-Flag antibody (gray). (A,B)Insets
show stage 40 embryos stained for the pan-neural
marker Xen1. Lateral views: anterior to the left, dorsal
to the top. Arrowheads indicate head defects in the
xMID morphant. (B,D)Arrowheads indicate noncontinuous and attenuated laminin staining in the
basal lamina. (E,F)Brackets indicate the lengths of
Flag-positive cells. (G)Asterisks indicate tubular
structures of the pronephros. (I,J)Asterisks indicate
luminal structures of the foregut. Arrowheads indicate
the lack of apical actin assembly in the xMID
morphant cells. (K,L)Transverse sections through the
brain of control-Mo-injected (K) and xMID-Moinjected (L) embryos, stained with anti-active caspase
3 (left) and anti-Flag (right) antibodies. Dashed line
delimits the ventricle.
Quintin et al., 2008). In vertebrates, Shroom3-mediated activation
of ROCKs and myosin II plays a crucial role in driving apical
constriction (Haigo et al., 2003; Hildebrand, 2005; Nishimura and
Takeichi, 2008; Rolo et al., 2009). Our study of the xMIDs raises
the important question of how microtubules control apical
constriction in neuroepithelial cells. In Drosophila, RhoGEF2
associates with microtubule plus ends in an EB1-dependent manner
(Rogers et al., 2004), and inhibition of microtubule polymerization
prevents apical actin assembly and Myosin light chain
phosphorylation, thus blocking apical constriction (Corrigall et al.,
2007). In addition, the enhancement of cadherin-based cell
adhesion is dependent on microtubules in human epithelial cells
(Meng et al., 2008). These data suggest that the molecular link
between microtubules and apical constriction is mediated by the
transport of key regulators of actin polymerization, myosin II
activation, and/or cell-cell contacts. It will therefore be intriguing
to examine whether intracellular transport is affected in xMID
morphants.
Insights into the molecular and pathological
mechanisms of Opitz G/BBB syndrome
We demonstrated that xMID1 is required for the morphogenesis of
epithelial organs, such as the cement gland, pronephros and
foregut. Furthermore, in the xMID morphants, the epithelial cell
morphology and organization were severely affected, and the
distribution of laminin in the basal lamina was compromised.
These data indicate that the morphogenetic defects in xMID
morphants are due to the loss of epithelial integrity.
In OS patients, various developmental abnormalities, including
craniofacial, urogenital, gastrointestinal and cardiovascular defects
are observed, although the pathological mechanisms have not been
identified (Fontanella et al., 2008; So et al., 2005). Importantly, a
recent analysis of the MID1 expression pattern in the human
embryo revealed it to be expressed in various epithelial tissues,
including the central nervous system, kidney primordia, and the
pharyngeal, respiratory and gastrointestinal epithelia (Pinson et al.,
2004). In addition, MID1 is expressed in the anal folds and genital
tubercle (Pinson et al., 2004). These expression patterns indicate a
strong correlation between epithelial MID1 expression and the
development of organs affected by OS. Although no extensive
histological characterizations of tissues from OS patients have been
reported, there are notable similarities in the pathological features
of OS patients and the epithelial defects of xMID morphants. In
addition, Mid1 shows similar epithelial expression patterns in
mouse (Dal Zotto et al., 1998) and chick (Richman et al., 2002).
We propose that common mechanisms underlie the normal
morphogenesis of the organs affected in OS patients and in the
Xenopus embryos in this study. Taken altogether, our findings
demonstrate the general importance of microtubule regulation by
MID1 and MID2 in cell shape change and maintenance in
epithelial morphogenesis during vertebrate embryogenesis.
Acknowledgements
We thank T. C. Cox, K. S. Kosik, K. Nakayama, T. Okubo, T. Takabatake, K.
Tamura and J. B. Wallingford for plasmids and reagents; and members of the
N.U. laboratory and S. Nonaka laboratory for valuable discussions, comments
and technical assistance. This work was supported by KAKENHI (07J05064,
21870043 to M.S.; 17207015, 21370102 to N.U.) from the Japan Society for
the Promotion of Science (JSPS). M.S. and Y.H. were supported by JSPS
Research Fellowships for Young Scientists.
Competing interests statement
The authors declare no competing financial interests.
Development 137 (14)
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/lookup/suppl/doi:10.1242/dev.048769/-/DC1
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DEVELOPMENT
Microtubule and xMIDs in neurulation