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W4285471841 abstract "Article Figures and data Abstract Editor's evaluation Introduction Results and discussion Materials and methods Data availability References Decision letter Author response Article and author information Metrics Abstract Failures of neural tube closure are common and serious birth defects, yet we have a poor understanding of the interaction of genetics and cell biology during neural tube closure. Additionally, mutations that cause neural tube defects (NTDs) tend to affect anterior or posterior regions of the neural tube but rarely both, indicating a regional specificity to NTD genetics. To better understand the regional specificity of cell behaviors during neural tube closure, we analyzed the dynamic localization of actin and N-cadherin via high-resolution tissue-level time-lapse microscopy during Xenopus neural tube closure. To investigate the regionality of gene function, we generated mosaic mutations in shroom3, a key regulator or neural tube closure. This new analytical approach elucidates several differences between cell behaviors during cranial/anterior and spinal/posterior neural tube closure, provides mechanistic insight into the function of shroom3, and demonstrates the ability of tissue-level imaging and analysis to generate cell biological mechanistic insights into neural tube closure. Editor's evaluation This manuscript by Baldwin and colleagues on vertebrate neural tube closure will be of interest to developmental and cell biologists studying tissue morphogenesis as well as human geneticists focusing on neural tube defects. It is timely, as it introduces a new technology for large-scale imaging of cell behaviours in large embryos. Specifically, it uses advanced image analysis to quantitatively describe and correlate active cell behaviours and localization dynamics of key cytoskeletal and adhesion proteins driving a central step of neural tube closure. Data analysis is detailed and followed by careful conclusions. https://doi.org/10.7554/eLife.66704.sa0 Decision letter eLife's review process Introduction Congenital birth defects are the number one biological cause of death for children in the US, and neural tube defects (NTDs) represent the second most common class of human birth defect (Murphy et al., 2018; Wallingford et al., 2013). NTDs represent a highly heterogenous group of congenital defects in which failure of the neural folds to elevate or fuse results in a failure of the skull or spine to enclose the central nervous system (Wallingford et al., 2013). While genetic analyses in both humans and animal models have revealed dozens of genes necessary for normal neural tube closure, several key questions remain. One central unanswered question relates to the regional heterogeneity of both normal neural tube closure and pathological NTDs. For example, the collective cell movements of convergent extension dramatically elongate the hindbrain and spinal cord of vertebrates, but not the midbrain and forebrain (Nikolopoulou et al., 2017; Wallingford et al., 2013). Accordingly, disruption of genetic regulators of convergent extension such as the planar cell polarity (PCP) genes results in failure of neural tube closure in posterior regions of the neural ectoderm, but not anterior (Kibar et al., 2001; Wang et al., 2006). Conversely, the shroom3 gene is implicated in apical constriction, a distinct cell behavior that drives epithelial sheet bending, and disruption of shroom3 elicits highly penetrant defects in anterior neural tube closure, but only weakly penetrant defects in the posterior (Haigo et al., 2003; Hildebrand and Soriano, 1999). This regional deployment of apical constriction in the anterior and convergent extension in the posterior during neural tube closure is poorly understood. In addition, the underlying mechanisms of individual cell behaviors necessary for neural tube closure remain incompletely defined. While apical constriction is driven by actomyosin contraction, the precise site of actomyosin action during this process is unclear and constitutes a long-term problem in the field (Martin and Goldstein, 2014). For example, analysis of apical constriction during gastrulation in both Drosophila and Caenorhabditis elegans has shown integration of discrete junctional and medio-apical (‘medial’) populations of actomyosin (Coravos and Martin, 2016; Martin et al., 2009; Roh-Johnson et al., 2012). Recent studies in frog and chick embryos have also described similar pulsed medial actomyosin-based contractions occurring during neural tube closure (Brown and García-García, 2018; Christodoulou and Skourides, 2015; Suzuki et al., 2017), but how those contractions are controlled and how they contribute to tissue-wide cell shape changes during neural tube closure are not known. For example, Shroom3 is among the more well-defined regulators of apical constriction, being both necessary and sufficient to drive this cell shape change in a variety of cell types, including the closing neural tube (Haigo et al., 2003; Hildebrand, 2005; Plageman et al., 2010; Plageman et al., 2011b). Shroom3 is known to act via Rho kinase to drive apical actin assembly and myosin contraction (Das et al., 2014; Hildebrand, 2005; Nishimura and Takeichi, 2008; Plageman et al., 2011a). However, the relationships between Shroom3 and the medial and junctional populations of actin have not been explored. An additional outstanding question relates to the interplay of actomyosin contractility and cell adhesion during apical constriction. The classical cadherin Cdh2 (N-cadherin) is essential for apical constriction during neural tube closure in Xenopus (Morita et al., 2010; Nandadasa et al., 2009), and shroom3 displays robust genetic interactions with n-cadherin in multiple developmental processes, including neural tube closure (Plageman et al., 2011b). Moreover, a dominant-negative N-cadherin can disrupt the ability of ectopically expressed Shroom3 to induce apical constriction in MDCK cells (Lang et al., 2014). Nonetheless, it is unclear if or how Shroom3 controls the interplay of N-cadherin and actomyosin during apical constriction. This is an important gap in our knowledge, because despite the tacit assumption that cadherins interact with each other and control actomyosin at cell-cell junctions, N-cadherin displays multiple cell-autonomous activities (Rebman et al., 2016; Sabatini et al., 2011). Intriguingly, several papers now demonstrate that extra-junctional cadherins at free cell membranes can engage and regulate the actomyosin cortex (Ichikawa et al., 2020; Padmanabhan et al., 2017; Sako et al., 1998; Wu et al., 2015). Finally, though Shroom3 has been extensively studied in the context of cranial apical constriction, the gene is expressed throughout the neural plate (Haigo et al., 2003; Hildebrand and Soriano, 1999) and recent studies have also implicated Shroom family proteins in the control of convergent extension (McGreevy et al., 2015; Nishimura and Takeichi, 2008; Simões et al., 2014). Several studies indicate a genetic and cell biological interplay of Shroom3 and the PCP proteins (Durbin et al., 2020; McGreevy et al., 2015), and one study directly links PCP, apical constriction, and convergent extension (Nishimura et al., 2012). Conversely, some studies also suggest a role for PCP proteins in apical constriction (Ossipova et al., 2015). Together, these studies highlight the complexity of neural tube closure, which is compounded by the sheer scale of the tissue involved. The neural ectoderm is comprised of hundreds to thousands of cells (depending on organism) and stretches from the anterior to posterior poles of the developing embryo. However, the vast majority of dynamic studies of cell behavior in the neural tube closure, including our own, have focused on small numbers of cells due to constraints of both imaging and image analysis. Here, we used image-tiling time-lapse confocal microscopy to obtain over 750,000 individual measurements of cell behaviors associated with neural tube closure in Xenopus tropicalis. Using these data, we demonstrate that the cell biological basis of apical constriction differs substantially between the anterior and the posterior neural plate. The data further suggest that the crux of Shroom3 function lies not in actin assembly per se, but rather in the coupling of actin contraction to effective cell surface area reduction. Third, we demonstrate that the control of N-cadherin localization is a key feature of Shroom3 function during neural tube closure. Finally, we demonstrate that the incompletely penetrant posterior phenotypes related to shroom3 loss stem from dysregulation of both actin and N-cadherin localization. Overall, these findings (a) elucidate differences between cell behaviors during cranial/anterior and spinal/posterior neural tube closure, (b) provide new insights into the function of shroom3, an essential neural tube closure gene, and (c) demonstrate the power of large-scale imaging and analysis to generate both cell-level mechanistic insight and new hypotheses for exploring neural tube closure. Results and discussion High-content imaging of cell behavior and protein localization during vertebrate neural tube closure X. tropicalis affords several advantages for imaging neural tube closure, as its cells are large and easily accessible; its culture conditions for imaging are no more complex than synthetic pond water held at room temperature; and its broad molecular manipulability allows examination of diverse fluorescent markers. We developed methods for confocal microscopy and image tiling to collect high-magnification datasets spanning broad regions of the folding neural ectoderm from embryos injected at blastula stages with mRNAs encoding fluorescent reporters (Figure 1A). At the onset of neurulation (approximately Nieuwkoop and Faber, 1994; Nieuwkoop and Faber, 1994, stages 12.5–13), embryos were positioned to image either the anterior or the posterior regions of the neural ectoderm. We then established a pipeline by which cells captured in our movies were segmented using Tissue Analyzer, CSML, and EPySeg (Aigouy et al., 2020; Aigouy et al., 2016; Ota et al., 2018), yielding a map of both the apical cell surfaces and all individual junctions (Figure 1B). Finally, we built pipelines to process these data with Tissue Analyzer (Aigouy et al., 2010; Aigouy et al., 2016) and Fiji (Schindelin et al., 2012) to quantify both cell behaviors and the localization of fluorescent protein reporters across the neural plate and across neurulation. Figure 1 with 1 supplement see all Download asset Open asset Tissue-level imaging and analysis of contractile protein dynamics during neural tube closure in Xenopus. (A) Schematic of mRNA injections and subsequent imaged regions of the Xenopus tropicalis embryo. (B) Cell segmentation and tracking workflow. Binary segmentation, cell surface tracking, and cell junction tracking were all generated using Tissue Analyzer. (C) Example Xenopus cells with analyzed subcellular domains labeled. Orange label = medial, cyan labels = junctional/junctions. (D) Schematic and N values of whole cell measurements. (E) Schematic and N values of individual cell junction measurements. With these methods in place, we considered three interrelated problems in neural tube closure biology: First, the incidence and form of NTDs differ widely between the brain and spinal cord (Nikolopoulou et al., 2017; Wallingford et al., 2013), yet our understanding of the dynamic cell behaviors in the two regions remains limited. Second, a unified mechanism for apical constriction has emerged in recent years involving the coordinated action of two discrete populations of actomyosin positioned either at apical cell-cell junctions or the medial apical cell surface (Coravos and Martin, 2016; Martin and Goldstein, 2014; Martin et al., 2009; Roh-Johnson et al., 2012), but the extent to which this model, developed in Drosophila and C. elegans, applies to vertebrates is unknown. Third, N-cadherin is essential for apical constriction in Xenopus (Nandadasa et al., 2009), but its functional interplay with junctional and/or medial actin is unknown. Accordingly, we made movies focused on either the anterior or posterior neural plate during neurulation, imaging the fluorescent actin biosensor LifeAct-RFP (Riedl et al., 2008; Figure 1A, magenta) and N-cadherin-GFP (Figure 1A), and independently quantified the mean fluorescent intensity of junctional and medial populations for both reporters (Figure 1C–D). To account for noise in these measurements, we have smoothed the data within individual cell tracks by averaging the data over a 7-frame window (Figure 1—figure supplement 1A, B). In total, our dataset is comprised of ~250,000 observations of apical cell surfaces from over 3700 cells and ~580,000 observations from over 13,000 individual cell-cell junctions across nine embryos (Figure 1D, E). Images were collected at a rate of 1 frame/observation per minute over 1–2 hr, spanning roughly stages 13–18. Initial cell sizes and fluorescent varied among cells and embryos due to both natural variation and staging as well as variation introduced via mRNA microinjection. Because our primary interest is in the dynamics of apical constriction, we standardized many of the parameters in our analyses to account for variation in cell size and fluorescent intensity. This standardization involved mean-centering the data for each individual cell track to zero and then dividing the resulting mean-centered values by the standard deviation of each track, such that the standardized parameters are now measured in standard deviations rather than square microns or arbitrary units, allowing for simpler comparisons of overall changes in parameters between cells over time (Figure 1—figure supplement 1A, D). We first performed an initial test of the validity of our approach, examining our dataset for well-known trends expected for neural epithelial cells during neural tube closure, namely an overall decrease in apical area and an overall increase in apical actin intensity. The heat maps in Figure 2A, B reveal that cells in both anterior and posterior regions generally reduce their apical surface area and increase medial actin intensity, as expected. These overall trends are backed by examination of individual cells, as shown for specific representative cells in Figure 2C. Overall, this analysis suggests that our pipeline is generally effective for quantifying cell shape and actin intensity over time during neural tube closure. Figure 2 Download asset Open asset Tissue-level analysis of individual cell behaviors reveals dynamic heterogeneity. (A) Overall change (Δ) in apical surface area (standardized) across anterior (left) and posterior (right) control embryos. (B) Overall change in medial LifeAct/actin localization (standardized) across anterior (left) and posterior (right) control embryos. Circle and triangle in A and B denote a representative cell for each embryo. Scale bars = 100 µm. (C) Standardized apical surface area (black) and medial actin (red) over time in representative cells from anterior (left/circle) and posterior (right/triangle). s.d. = standard deviation. Distinct patterns of apical constriction behavior, actin assembly, and N-cadherin localization in anterior and posterior regions of the closing neural tube Our dataset revealed several interesting trends. First, while bulk measurements showed a decrease in apical area in both anterior and posterior regions over time (Figure 3A), we observed distinct region-specific distributions for these changes. For example, in the anterior, the vast majority of cells displayed significant apical constriction, and this constriction proceeded gradually across neurulation (Figure 3A and A’, left). In the posterior, however, a much smaller proportion of posterior cells constricted and a substantial number actually dilated (Figure 3A, right). Moreover, constriction of cells in the posterior was initiated very late in neurulation and proceeded very rapidly (Figure 3A’, right). Figure 3 with 1 supplement see all Download asset Open asset Cells in the anterior and posterior neural ectoderm both apically constrict but differ in their contractile protein dynamics. Tissue-level cell size and protein localization dynamics from control embryos in Figure 2. (X) Distribution of overall change (Δ) in displayed parameter (standardized) among cells from control embryos. Horizontal lines on density plots/violins indicate quartiles of distribution. Black circles are individual cells. Statistical comparisons performed by Kolmogorov-Smirnov (KS) test. (X’) 2D density plots of standardized variable versus time for all observations/cells in each control embryo in Figure 2. Green points are measurements from the representative cells denoted in Figure 2. s.d. = standard deviation. We further observed that both medial and junctional actin intensity generally increased over time in both regions, with their temporal progressions being reciprocal to the changes in apical area described above (Figure 3B, B’, C, and C’, right). Again, these distributions were significantly different between the anterior and posterior regions, with cells in the spine having a significantly more heterogeneous distribution of actin accumulation outcomes (Figure 3B, C, left). By far the most intriguing results related to the dynamics of N-cadherin localization, for which we observed two surprising patterns. First, in the anterior neural plate N-cadherin accumulated dramatically not only in the junctional region but also in the medial region (i.e. the free apical surface) (Figure 3D, D’, E and E’). Thus, N-cadherin localization closely parallels actin dynamics in the normal anterior neural plate. This result was surprising because classical cadherins such as N-cadherin are typically known for their action at cell-cell junctions. Nonetheless, immunostaining for endogenous N-cadherin in fixed embryos confirmed this medial accumulation in the apical surfaces of anterior neural ectoderm cells (Figure 3—figure supplement 1). Our dataset lacked the time resolution to determine precise patterns of N-cadherin movement during apical constriction, but in Z-projections of highly constricted cells, we observed N-cadherin signal not just coincident with, but also basal to, to the apical actin signal (Figure 4). This result is consistent with the emerging understanding of the cell-autonomous roles for cadherins in both actin organization and endocytosis (Ichikawa et al., 2020; Padmanabhan et al., 2017; Rebman et al., 2016; Sabatini et al., 2011; Sako et al., 1998; Wu et al., 2015). Figure 4 with 1 supplement see all Download asset Open asset N-cadherin localizes both at the apical surface and basally as well. (A) XY (top row) and XZ (bottom row) projections of N-cadherin-GFP and LifeAct-RFP in the anterior neural ectoderm of a Xenopus tropicalis embryo. (B) XY (top panel) and XZ (bottom panel) projections of NCD-2 (monoclonal α-N-cadherin antibody) in the anterior neural ectoderm of a X. tropicalis embryo. Dashed cyan lines marks the position of the XZ projection. The medial accumulation of both N-cadherin and actin led us to explore the localization of Shroom3 itself. No antibodies are available for Xenopus Shroom3, and gain-of-function effects during early development preclude analysis of tagged wild-type Shroom3 during neural tube closure. That said, ectopic Shroom3 clearly decorates both junctional and medial regions in diverse epithelial cells (Haigo et al., 2003; Kowalczyk et al., 2021; Lee et al., 2009). To gain insight into Shroom3 localization during neural tube closure, we imaged the localization of the a GFP-tagged Shroom3 construct lacking the c-terminal Rok-binding domain, similar to a construct previously used to explore Shroom dynamics in Drosophila (Farrell et al., 2017; Simões et al., 2014). In movies of the folding neural plate, the construct localized in a pattern essentially identical to actin, accumulating in both junctional and medial regions of the anterior neural plate (Figure 4—figure supplement 1). Finally, we observed a strikingly different trend in the posterior neural plate, where N-cadherin dynamics did not closely parallel actin dynamics. In fact, neither junctional nor medial N-cadherin displayed significant accumulation in the posterior neural plate during the period of observation (Figure 3D, E), despite robust actin accumulation in this region (Figure 3B, C). Together, these data provide a comprehensive, quantitative description of apical constriction, actin dynamics and N-cadherin localization in the anterior and the posterior neural plate during Xenopus neural tube closure. The data further suggest that the mechanisms linking actin and N-cadherin to apical surface area differ in the two regions. Mosaic mutation of Shroom3 reveals distinct anterior and posterior phenotypes in the neural ectoderm The differences in cell behaviors we observed between anterior and posterior neural ectodermal regions reflect the region-specific nature of NTDs in both humans and animal models. To explore the relationships in more detail, we next turned to loss-of-function manipulation of shroom3, which is implicated in human NTDs and has variably penetrant effects on anterior and posterior neural tube closure (Deshwar et al., 2020; Haigo et al., 2003; Hildebrand and Soriano, 1999; Lemay et al., 2015). F0 mutagenesis using CRISPR has recently emerged as a powerful tool in Xenopus and zebrafish, and mosaic crispants generated by targeted injections allow simultaneous observation of wild-type and crispant cellular phenotypes so that observations are automatically staged and synchronized (Aslan et al., 2017; Kakebeen et al., 2020; Kroll et al., 2021; Szenker-Ravi et al., 2018; Willsey et al., 2020). We therefore designed sgRNAs that effectively targeted the coding region of shroom3, approximately 28 amino acids from the 5’ end of the transcript, such that any indels generated by CRISPR targeting are likely to disrupt all functional domains of the Shroom3 protein (Figure 5—figure supplement 1A). Using injection into the two dorsal-animal blastomeres at the 8-cell stage to target the neural plate, we demonstrated that our sgRNAs elicited both mutation of the shroom3 locus as well as the anterior neural tube closure defects expected based on results from knockdown in Xenopus using MOs (Haigo et al., 2003), as well as the results in mouse genetic mutants (Hildebrand and Soriano, 1999). As a critical negative control, injections of sgRNA without Cas9 protein had no effect (Figure 5—figure supplement 1C, D). We next performed more targeted injections to generate mosaic embryos. To do so, we labeled the neural plate by injection of fluorescent reporters into both dorsal blastomeres at the 4-cell stage, and then injected a mixture of shroom3-targeted sgRNA, Cas9 protein, and membrane-BFP mRNA into one dorsal blastomere of 8-cell stage embryos (Figure 5A and see Figure 5—figure supplement 1). We then identified shroom3 crispant cells via membrane-BFP localization (Figure 5B). Because cell junction behavior may be altered at mosaic cell-cell interfaces (i.e. junctions between a control and a crispant cell), we excluded this relatively small number of cells from our analysis. Importantly, this mosaic F0 CRISPR-based approach also generally recapitulated the known phenotype of Shroom3 loss, as we observed gross failure of anterior neural tube closure. Figure 5 with 3 supplements see all Download asset Open asset Disruption of shroom3 via mosaic F0 CRISPR mosaic causes differential apical constriction phenotypes between regions of the neural ectoderm. (A) Schematic of mosaic F0 CRISPR/Cas9 injections in Xenopus tropicalis embryos. (B) Workflow of identification and analysis of mosaic F0 crispants. (C) Top row, distribution of initial area (square microns) of tracked cells from anterior (left) and posterior (right) embryos. Lower row, distribution of final area (square microns) of tracked cells. (D) Distribution of overall change (Δ) in apical area (standardized) from all cells/embryos. In C and D, horizontal lines on density plots/violins indicate quartiles of distribution. Black circles are individual cells. Statistical comparisons performed by Kolmogorov-Smirnov (KS) test. Cells situated along the mosaic interface were excluded from these analyses. s.d. = standard deviation. At the level of cell behaviors, we observed a surprising difference in anterior and posterior phenotypes. In the anterior region, shroom3 crispant cells displayed significantly enlarged apical surfaces at the onset of our imaging (~stage 13), and this phenotype grew more severe over time (Figure 5C, left); the majority of cells not only failed to constrict but instead dilated (Figure 5D, left). In the posterior, however, the majority of shroom3 crispant cells still strongly constricted, though collectively they displayed a mildly significant defect in apical constriction (Figure 5C, D, right). Thus, the magnitude of apical constriction defects in shroom3 crispant cells reflects the penetrance of NTDs in the anterior and posterior regions (Haigo et al., 2003; Hildebrand and Soriano, 1999). Loss of Shroom3 uncouples actin dynamics from N-cadherin localization in the anterior neural ectoderm Loss of Shroom3 disrupts apical actin assembly in the neural plate (Haigo et al., 2003; McGreevy et al., 2015), but the precise nature of this defect and whether or how it relates to junctional and/or medial actin is unknown. Likewise, N-cadherin is implicated in Shroom3 function and apical constriction (Lang et al., 2014; Nandadasa et al., 2009; Plageman et al., 2011b), but how this relates to actin dynamics is poorly defined. We therefore examined the relationship between apical constriction, actin dynamics, and N-cadherin dynamics, focusing first on the anterior neural plate. We found that wild-type cells tended to increase both medial and junctional actin localization over time, as expected (Figure 6A–C, left). Consistent with the known role in apical actin accumulation (Haigo et al., 2003; Hildebrand, 2005), shroom3 crispant cells displayed significantly reduced accumulation of both medial and junctional actin (Figure 6B, C, blue violins). However, this effect was surprisingly mild, and in fact, the majority of shroom3 crispant cells actually increased both junctional and medial actin over the period of imaging (Figure 6B, C, blue violins). Figure 6 Download asset Open asset Medial actin accumulation drives apical constriction while loss of shroom3 disrupts actin accumulation and constriction in the anterior neural ectoderm. (A) Representative images of LifeAct/actin localization in control cells (left) and shroom3 crispant cells (right) from the anterior region of the neural ectoderm. Scale bar = 15 µm. (B) Distribution of overall change (Δ) in medial LifeAct/actin (standardized) from anterior cells. (C) Distribution of overall change (Δ) in junctional LifeAct/actin (standardized) from anterior cells. In B and C, horizontal lines on density plots/violins indicate quartiles of distribution, black circles are individual cells, and statistical comparisons performed by Kolmogorov-Smirnov (KS) test. (D and E) 2D distribution of changes in apical area and medial (D) or junctional (E) LifeAct/actin (both standardized). Percentages in white indicate the percentage of total cells in each quadrant. Statistical comparisons performed by Peacock test, a 2D implementation of the KS test. (F and G) 2D density plots of all observations of apical area versus medial (F) or junctional (G) LifeAct/actin for all cells within each group. Red lines indicate best-fit line through the observations. Statistics (r and p) are calculated for Pearson’s correlation. Cells situated along the mosaic interface were excluded from these analyses. s.d. = standard deviation. In contrast to this surprisingly modest change in actin intensity (Figure 6B, C), bulk measurements revealed that shroom3 crispant cells displayed a profound failure to accumulate both junctional and medial N-cadherin, and in fact the majority of cells actually reduced N-cadherin levels (Figure 6A, D, and E, blue violins). Moreover, this effect was far more pronounced for the medial population of N-cadherin (Figure 6D). Thus, loss of Shroom3 elicits a substantially more severe effect on the dynamics of N-cadherin than of actin, apparently uncoupling the two. Shroom3 links actin and N-cadherin dynamics to effective apical constriction in the anterior neural ectoderm To explore these surprising results in more detail, we directly compared changes in apical area with changes in actin and N-cadherin intensity for each cell individually. In 838 control cells, we observed that the vast majority displayed a strong reduction in apical area and a strong increase in both junctional and medial actin intensity (Figure 7A, B). As noted in the bulk statistics above, the majority of 147 shroom3 crispant cells displayed increased actin intensity; however, these crispant cells displayed a bimodal distribution of changes in apical area, with some cells constricting and other cells dilating, yet even cells that increased their apical area after loss of Shroom3 nonetheless accumulated medial and junctional actin (Figure 7A, B). Thus, loss of shroom3 does not lead to loss of apical actin in the anterior neural plate but rather to a reduced accumulation of apical actin. Figure 7 Download asset Open asset Medial N-cadherin accumulation is severely disrupted in anterior shroom3 crispant cells that fail to apically constrict. (A) Representa" @default.
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W4285471841 title "Decision letter: Global analysis of cell behavior and protein dynamics reveals region-specific roles for Shroom3 and N-cadherin during neural tube closure" @default.
W4285471841 doi "https://doi.org/10.7554/elife.66704.sa1" @default.
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