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Cell volume changes contribute to epithelial morphogenesis in zebrafish Kupffer’s vesicle

  1. Agnik Dasgupta
  2. Matthias Merkel
  3. Madeline J Clark
  4. Andrew E Jacob
  5. Jonathan Edward Dawson
  6. M Lisa Manning  Is a corresponding author
  7. Jeffrey D Amack  Is a corresponding author
  1. State University of New York, Upstate Medical University, United States
  2. Syracuse University, United States
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Cite this article as: eLife 2018;7:e30963 doi: 10.7554/eLife.30963

Abstract

How epithelial cell behaviors are coordinately regulated to sculpt tissue architecture is a fundamental question in biology. Kupffer’s vesicle (KV), a transient organ with a fluid-filled lumen, provides a simple system to investigate the interplay between intrinsic cellular mechanisms and external forces during epithelial morphogenesis. Using 3-dimensional (3D) analyses of single cells we identify asymmetric cell volume changes along the anteroposterior axis of KV that coincide with asymmetric cell shape changes. Blocking ion flux prevents these cell volume changes and cell shape changes. Vertex simulations suggest cell shape changes do not depend on lumen expansion. Consistent with this prediction, asymmetric changes in KV cell volume and shape occur normally when KV lumen growth fails due to leaky cell adhesions. These results indicate ion flux mediates cell volume changes that contribute to asymmetric cell shape changes in KV, and that these changes in epithelial morphology are separable from lumen-generated forces.

https://doi.org/10.7554/eLife.30963.001

Introduction

In an embryo, epithelial cells undergo tightly regulated shape changes to drive tissue remodeling and organ development. Changes in epithelial cell morphology can be mediated by intrinsic mechanisms, such as rearrangements of the actomyosin cytoskeleton that often occur in response to biochemical signaling cascades (Fuss et al., 2004; Escudero et al., 2007). Extrinsic biophysical forces can also influence epithelial morphogenesis (Navis and Nelson, 2016; Navis and Bagnat, 2015). For example, the mechanical properties of neighboring cells can help shape how an epithelium develops (Luu et al., 2011; Sedzinski et al., 2016). Another source of extrinsic force found in many organs is a fluid-filled lumen. Forces generated by increased fluid pressure during lumen expansion can have an impact on individual cell shapes and the overarching epithelial architecture (Bagnat et al., 2010). Conversely, movements of fluids from epithelial cells to the lumen have been proposed to regulate both lumen growth and thinning of the epithelium (Hoijman et al., 2015). Thus, exactly how intrinsic molecular mechanisms and extrinsic mechanical forces interact to regulate epithelial cell shape changes during organogenesis remains an open and intriguing question.

In this study, we used the zebrafish Kupffer’s vesicle (KV) as a model organ to investigate mechanisms that regulate shape changes of single cells during epithelial morphogenesis. KV, which functions as a ‘left-right organizer’ to determine left-right asymmetry of the zebrafish embryo (Essner et al., 2005; Kramer-Zucker et al., 2005), is a transient organ comprised of a single layer of ~50 epithelial cells that surround a fluid-filled lumen. Each KV cell extends a motile monocilium into the lumen to generate asymmetric fluid flows that direct left-right patterning signals. Fate mapping studies have identified precursor cells, called dorsal forerunner cells (DFCs), which differentiate into epithelial KV cells at the end of gastrulation stages of development (Melby et al., 1996; Cooper and D'Amico, 1996). These cells then form the KV organ in the tailbud at the embryonic midline (marked by notochord) during early somitogenesis stages (Figure 1A). Similar to other organs that develop a fluid-filled lumen—such as the gut tube (Alvers et al., 2014) or pancreas (Villasenor et al., 2010)—KV cells form a rosette-like structure to give rise to a nascent lumen that expands over time (Amack et al., 2007; Oteíza et al., 2008). Previous 2-dimensional (2D) analyses of KV cells revealed that during expansion of the KV lumen, KV cells at the middle focal plane undergo asymmetric cell shape changes along the anteroposterior (AP) axis that sculpt the architecture of the mature KV organ (Wang et al., 2012). A transgenic strain developed in this study, Tg(sox17:GFP-CAAX), provides bright labeling of KV cell membranes with green fluorescent protein (GFP) that shows the AP asymmetric architecture of the whole organ (Video 1) and the 2D shapes of epithelial KV cells (Figure 1A). KV cells have similar shapes at early stages of development, whereas at later stages the cells in the anterior half of KV (KV-ant cells) develop columnar morphologies that allow tight packing of these cells and posterior KV cells (KV-post cells) become wide and thin (Wang et al., 2012) (Figure 1B). This morphogenetic process, which we refer to as ‘KV remodeling’, results in an AP asymmetric distribution of cilia that is necessary to drive fluid flows for left-right patterning (Wang et al., 2012). Thus, KV is a simple and accessible organ that is ideal for probing the relationship between intrinsic and extrinsic mechanisms that drive epithelial morphogenesis.

Mosaic labeling of KV cells.

(A) A dorsal view of the tailbud in a live zebrafish embryo at the 8-somite stage (8 ss) of development. Kupffer’s vesicle (KV) is positioned at the end of the notochord. The inset shows GFP-labeled KV cells surrounding the fluid-filled KV lumen in a Tg(sox17:GFP-CAAX) transgenic embryo at 8 ss. This is the middle plane of the KV. Scale = 10 μm. (B) Schematic of cell shape changes during KV remodeling. KV-ant cells (blue) and KV-post cells (red) have similar shapes at 2 ss, but then undergo regional cell shape changes such that KV-ant cells are elongated and KV-post cells are wide and thin by 8 ss. These cell shape changes result in asymmetric positioning of motile cilia that generate fluid flows for left-right patterning. (C) Structure of the ubi:zebrabow and sox17:CreERT2 transgenes and the possible recombination outcomes of the ‘zebrabow’ transgene by Cre recombinase activity in KV cell lineages. (D) Time course of mosaic labeling of KV cells. Brief treatment of double transgenic Tg(sox17:CreERT2); Tg(ubi:Zebrabow) embryos with 4-OHT from the dome stage to the shield stage generates low levels of Cre activity that changes expression of default RFP to expression of YFP in a subset of KV cells. (E) Mosaic labeled YFP+ KV cells at the middle plane of KV at 8 ss. (F) 3D reconstructed KV cells (green) and KV lumen (magenta) at 8 ss. Scale = 10 μm. (G–H) Morphometric parameters of 3D rendered KV-ant (G) and KV-post (H) cells: length = axis spanning from apical to basal side of the cell, height = axis spanning from dorsal to ventral side of the cell, and width = axis connecting lateral sides of the cell. Scale = 5 μm.

https://doi.org/10.7554/eLife.30963.002

Previous studies of KV have successfully contributed to our understanding of how epithelial cell shapes are regulated during embryogenesis, but the mechanisms that control KV cell shape changes are not fully understood. Experimental results and mathematical simulations from our group indicate that actomyosin contractility and differential interfacial tensions between KV cells mediate asymmetric cell shape changes (Wang et al., 2012). Additional studies identified an AP asymmetric deposition of extracellular matrix (ECM) implicated in restricting anterior KV cell shape during KV lumen expansion (Compagnon et al., 2014). We reasoned that these mechanisms likely work in concert with yet additional mechanisms to fully instruct epithelial morphogenesis during KV organ formation. Here, we developed methods to analyze single KV cells in 3-dimensions (3D) and created novel mathematical vertex models of KV development to identify mechanisms that contribute to AP asymmetric epithelial cell shape changes in KV. 3D analyses revealed that KV-ant cells increase their volume and KV-post cells decrease their volume during KV morphogenesis. These asymmetric cell volume changes occur at the same time as asymmetric cell shape changes. At the molecular level, KV cell volume and shape changes are mediated by ion channel activity that regulates ion flux and fluid transport. We next tested whether extrinsic biophysical forces had an impact on these cell morphology changes. Mathematical models indicate that mechanical properties of external tissues surrounding the KV can impact cell shape changes in the KV. Models predicted that when external tissues are solid-like, asymmetric cell volume changes in KV cells contribute to cell shape changes even in the absence of lumen expansion. Consistent with mathematical model predictions, experimental perturbations of lumen expansion indicated that changes in KV cell volume and shape can occur independent of forces associated with lumen growth. Together, our results suggest ion channel mediated fluid flux serves as an intrinsic mechanism to regulate epithelial cell morphodynamics that create asymmetry in the KV organ. These findings shed new light on the interplay between lumenogenesis and epithelial morphogenesis and provide an example of cell morphology changes that can be uncoupled from mechanical forces exerted during lumen expansion.

Video 1
KV organ architecture in 3D in a live Tg(sox17:GFP-CAAX) embryo at 8 ss.

The membrane-localized GFP marks all cells in KV. KV is rotating along its anteroposterior (AP) axis. Scale = 20 μm.

https://doi.org/10.7554/eLife.30963.003

Results

Mosaic labeling enables 3D analysis of single KV cells

To investigate 3D behaviors of KV cells we first generated stable Tg(sox17:GFP-CAAX) transgenic zebrafish using a sox17 promoter (Sakaguchi et al., 2006) to express membrane localized GFP (GFP-CAAX) in KV cells. This transgene marks all cells in the KV and is useful for delineating 2D cell morphology (Figure 1A). However, due to difficulties in determining exact cell-cell boundaries in the KV (Video 1), this strain is not ideal for visualizing individual KV cells in 3D. Therefore, we next developed a Cre-loxP based mosaic cell labeling method to visualize single KV cells. For this approach, we generated transgenic Tg(sox17:CreERT2) zebrafish that expresses Cre recombinase in the KV cell lineage that has inducible activity through the addition of 4-hydroxytamoxifen (4-OHT) (Feil et al., 1997). Transgenic Tg(sox17:CreERT2) fish were crossed with a previously described Tg(ubi:Zebrabow) (‘zebrabow’) strain (Pan et al., 2013) that can be used to generate differential fluorescent labeling of cells based on stochastic Cre-mediated recombination of the zebrabow transgene (Figure 1C). Double transgenic Tg(sox17:CreERT2); Tg(ubi:Zebrabow) embryos were briefly treated with 4-OHT from the dome stage (4 hr post-fertilization or hpf) to the shield stage (6 hpf) (Figure 1D) to induce low levels of Cre activity that resulted in a switch from the default RFP (red fluorescent protein) expression to YFP (yellow fluorescent protein) expression in a subset of KV cells (Figure 1E). This strategy reliably created mosaic labeled KVs containing a few YFP+ KV cells with boundaries that are easily distinguished from surrounding RFP+ cells (Video 2). Images of mosaic labeled KVs in living Tg(sox17:CreERT2); Tg(ubi:Zebrabow) embryos were volume-rendered using Imaris (Bitplane) software to generate 3D reconstructions of the lumen and KV cells (Figure 1F). To assess the 3D morphology of individual KV cells residing in either the anterior or posterior region of the KV organ, we used the lumen surface as a reference point to ensure that all 3D datasets were analyzed from the same perspective. To make morphometric measurements, we defined three axes of KV cells: (1) cellular ‘height (H)’ is the length of the axis connecting dorsal and ventral surfaces of the cell, (2) cellular ‘length (L)’ is the length of the axis connecting apical (associated with the lumen) and basal surfaces of the cell, and (3) cellular ‘width (W)’ is the length of the axis connecting two lateral sides of the cell (Figure 1G,H). The combination of mosaic labeling, live imaging and 3D cell morphometrics provides a new approach to investigate the cellular and mechanical mechanisms underlying KV epithelial morphogenesis at single cell resolution.

Video 2
3D projection of KV in a live mosaic labeled Tg(sox17:CreERT2); Tg(ubi:Zebrabow) embryo at 8 ss.

Stochastic Cre-mediated recombination labels only few cells with YFP expression with clear boundaries that are easily distinguishable from non-recombined RFP+ cells. The KV organ is rotating along its anteroposterior (AP) axis. Scale = 20 μm.

https://doi.org/10.7554/eLife.30963.004

To assess the dynamics of KV cells in 3D, we first performed time-lapse imaging of mosaic labeled KVs in live Tg(sox17:CreERT2); Tg(ubi:Zebrabow) embryos from the 2-somite stage (2 ss) when the lumen first forms to the 8 somite stage (8 ss) when the lumen is fully expanded (Amack et al., 2007; Wang et al., 2012; Gokey et al., 2016). These stages span the process of KV remodeling when KV cells change their shapes: 2 ss is pre-remodeling and 8 ss is post-remodeling (Wang et al., 2012). Time-lapse images from these stages indicated KV cells are highly dynamic during KV morphogenesis (Video 3). To quantify KV cell dynamics, we took several precautions in subsequent experiments to avoid artifacts. First, to avoid potential photobleaching from time-lapse imaging, live embryos were imaged only once at one stage of development—not continuously or at multiple stages. Second, to avoid differences in fluorescence signal due to differences in imaging depth, all KVs were visualized laterally (YZ orientation) and only mosaically labeled cells from the middle plane of the KV organ that is perpendicular to the dorsoventral axis were selected for analysis (Figure 2A). We define the middle plane as the plane with the largest lumen diameter when viewed dorsally (Figure 1A). Finally, we determined that the Cre activity was not spatially biased, but rather randomly labeled cells throughout the KV. By analyzing enough embryos, we sampled KV cells from all positions along the middle plane of KV at different stages of development (Figure 2—figure supplement 1).

Figure 2 with 3 supplements see all
3D morphometric analysis of single cells reveals asymmetric cell volume changes during asymmetric KV cell shape changes.

(A) A lateral view of a mosaic labeled KV in a Tg(sox17:CreERT2); Tg(ubi:Zebrabow) at 2 ss. The embryo diagram represents the orientation of the image. The notochord and KV are outlined in blue. Yellow lines mark the KV lumen. Examples of 3D reconstructed KV-ant and KV-post cells along the middle plane of KV are shown. Scale = 10 μm. (B) Representative snapshots of 3D rendered KV-ant and KV-post cells at different stages of KV development between 2 ss and 8 ss. The parameters including height (H), length (L) and width (W) were used to quantify cell morphology. Yellow lines indicate the KV luminal surface. Scale = 10 μm. (C–E) Quantification of height (C), length (D) and width (E) of individual KV-ant and KV-post cells during development. (F) A length-to-width ratio (LWR) was used to describe KV cell shapes. KV-ant and KV-post cells change shape between 4 ss and 6 ss. (G) Volume measurements of individual KV cells at different stages of development. Similar to cell shapes, KV-ant and KV-post cells change volume between 4 ss and 6 ss. All measurements presented in C-G were made on the same group of reconstructed cells. The number of KV-ant and KV-post cells analyzed is indicated in the graph in C. N = number of embryos analyzed at each stage. Graphs show the mean + SD. Results were pooled from three independent experiments. *p<0.01 and ns = not significant (p>5% with Welch’s T-Test).

https://doi.org/10.7554/eLife.30963.005
Video 3
Time-lapse imaging of KV cells in a live mosaic labeled Tg(sox17:CreERT2); Tg(ubi:Zebrabow) embryo treated with 4-OHT from dome stage to shield stage.

Images were collected every 5 min from 2 ss to 8 ss. The movie spans 105 min of development. Single 3D rendered KV-ant (blue) and KV-post (red) cells are followed during KV morphogenesis. Scale = 20 μm.

https://doi.org/10.7554/eLife.30963.009

To quantify the morphometric properties of KV-ant and KV-post cells during KV morphogenesis, live embryos with mosaic labeled KVs were imaged at a specific stage of development (2 ss, 4 ss, 6 ss and 8 ss) and then individual cells at the middle plane of the organ were volume-rendered (Figure 2B). We measured the height, length and width of KV cells from several wild-type embryos to determine the average parameters for KV-ant and KV-post cells. The results showed cell-to-cell variability, but pooling measurements from multiple cells at a specific developmental stage identified trends and statistically significant differences during KV morphogenesis. First, cell height of both KV-ant cells and KV-post cells increased from 2 ss to 8 ss (Figure 2C), reflecting the expansion of the apical surface of cells (see dotted line in Figure 2B) to accommodate lumen growth. There were no significant differences in cell height between KV-ant cells and KV-post cells during these stages (Figure 2C). Second, cell length decreased in both KV-ant cells and KV-post cells during KV morphogenesis, but a sharp reduction was observed from 4 ss to 6 ss in KV-post cells (Figure 2D). This resulted in the cell length of KV-post cells to become significantly different from KV-ant cells after 4 ss. Importantly, it is between 4 ss to 6 ss when cell shape changes associated with KV remodeling were previously observed (Compagnon et al., 2014; Wang et al., 2012). Third, analysis of cell width indicated KV-ant cells remained relatively constant during KV morphogenesis, whereas KV-post cells showed a distinctive increase in width between 4 ss and 6 ss that resulted in significant differences between KV-ant cells and KV-post cells after 4 ss (Figure 2E). In order to make comparisons with previous 2D studies, we calculated length-to-width ratios (LWR) to describe cell morphology. We found that both KV-ant and KV-post cells had similar morphologies at 2 ss and 4 ss and that these morphologies changed after 4 ss such that KV-ant cells had a significantly larger LWR than KV-post cells (Figure 2F). This analysis of individual KV-ant and KV-post cell shapes is consistent with asymmetric cell shape changes associated with KV remodeling that were previously identified using 2D measurements of KV cell LWRs (Wang et al., 2012) and provides for the first time a 3D quantification of epithelial cell morphodynamics that occur during KV development.

3D analysis of single cells reveals asymmetric cell volume changes during KV morphogenesis

We next used mosaic labeling to address whether the volume of individual KV cells changes during development. Since the cells are not regularly shaped, their volume is not simply given by the product of the length, width and height. Measuring the volume of 3D reconstructed KV cells revealed striking dynamics in cell size that mirrored changes in cell shape. At 2 ss and 4 ss, KV-ant and KV-post cells showed similar cell volumes (Figure 2G). However, between 4 ss to 6 ss—when KV cells change shape—the volume of KV-ant cells increased and the volume of KV-post cells decreased (Figure 2G). These dynamics were visualized in real time by tracking individual KV-ant and KV-post cells continuously for a brief time window (55 min) via time-lapse imaging during the critical period between 4 ss and 6 ss in live mosaic labeled embryos (Figure 2—figure supplement 2; Video 4). Our results indicate cell volume becomes significantly different between KV-ant and KV-post cells after 4 ss (Figure 2G). Thus, differential cell volume changes along the AP axis occur during the same stage as cell shape changes. Overall, between 2 ss and 8 ss, KV-ant cells increased volume from 2106 ± 1014 μm3 to 2547 ± 693 μm3 (shown are mean ±one standard deviation) and KV-post cell decreased volume from 2180 ± 1034 μm3 to 1564 ± 539 μm3 (Figure 2G). The considerable standard deviations here likely reflect the high degree of variability in KV size among wild-type embryos (Gokey et al., 2016). It is important to note that while the cell sizes are variable, the direction of size changes during development is constant. KV-ant cells always increase in volume and KV-post cells always decrease in volume. We refer to these regional changes in KV cell size along the AP body axis as asymmetric cell volume changes, which provide new insight into the mechanics of epithelial morphogenesis in KV.

Video 4
Time-lapse imaging of KV cells in a live mosaic labeled Tg(sox17:CreERT2); Tg(ubi:Zebrabow) embryo treated with 4-OHT from dome stage to shield stage.

Images were collected every 5 min from 4 ss to 6 ss. The movie spans 55 min of development. Single 3D rendered KV-ant (blue) and KV-post (red) cells are followed during KV morphogenesis. Scale = 10 μm.

https://doi.org/10.7554/eLife.30963.010

To better understand volume changes during KV development, we measured the volume of the entire KV organ at different stages. To make these measurements in live embryos, Tg(sox17:GFP-CAAX) transgenic fish were crossed with Tg(actb2:myl12.1-MKATE2) zebrafish that have enriched apical membrane expression of the fluorescent mKate2 protein fused to Myl12.1 (myosin light chain 12 genome duplicate 1) in KV cells. Double transgenic Tg(actb2:myl12.1mKATE2; Tg(sox17:GFP-CAAX) embryos were used to visualize both KV lumen and KV cells in living embryos (Figure 2—figure supplement 3A). The KV lumen was reconstructed in 3D using Tg(actb2:myl12.1-MKATE2) expression and then divided into two equal halves (hence equal volumes) along the AP axis (Figure 2—figure supplement 3B,B’). Next, the total KV cellular component of KV-ant and KV-post cells was reconstructed using Tg(sox17:GFP-CAAX) expression (Figure 2—figure supplement 3B,B’). Volume measurements indicated that both ‘total KV-ant cellular volume’ and ‘total KV-post cellular volume’ were similar at 2 ss, but then were significantly different at 8 ss (Figure 2—figure supplement 3C). Overall, ‘total KV-ant cellular volume’ increased from 1.4 × 105 μm3 to 1.7 × 105 μm3 and ‘total KV-post cell volume’ decreased from 1.2 × 105 μm3 to 0.98 × 105 μm3 between 2 ss and 8 ss stages (Figure 2—figure supplement 3C). Thus, consistent with volume changes in single cells at the middle plane of KV, we observed asymmetric volume changes along the anterior and posterior axis of KV when the entire cellular component of the organ was analyzed. These results suggested that asymmetric cell volume changes might contribute to lumen growth and/or cell shape changes during the concurrent processes of lumen expansion and epithelial morphogenesis in KV.

Inhibiting ion flux disrupts asymmetric cell volume changes, lumen expansion and cell shape changes in KV

We next sought to identify a mechanism that mediates KV cell volume changes between 2 ss and 8 ss. At the molecular level, cellular volume can be controlled through coordinated flux of Na+, K+ and Cl- ions via specific ion channels and pumps that results in osmotically driven water transport (Wilson et al., 2007; Barrett and Keely, 2000; Frizzell and Hanrahan, 2012; Frizzell, 1995; Damkier et al., 2013; Spring and Siebens, 1988; Hoijman et al., 2015; Saias et al., 2015). To test whether cell volume changes in KV cells are mediated by ion flux, we inhibited either the sodium-potassium pump (Na+/K+-ATPase) or the cystic fibrosis transmembrane conductance regulator (Cftr). Both have previously been shown to play a role in KV lumen expansion (Navis et al., 2013; Compagnon et al., 2014). We first treated mosaic-labeled (e.g. 4-OHT treated) Tg(sox17:CreERT2); Tg(ubi:Zebrabow) embryos with the small molecule ouabain to inhibit the Na+/K+-ATPase pump as previously described (Compagnon et al., 2014). The Na+/K+-ATPase has a central role in generating an electrochemical gradient across the plasma membrane that drives transport of water and solutes (Wilson et al., 2007; Damkier et al., 2013). Ouabain treatments are known to disrupt ion flux, resulting in an increase in intracellular sodium and calcium concentrations. To block Na+/K+-ATPase during the critical stages when cells change volume, we treated mosaic-labeled embryos with ouabain starting at 4 ss (11 hpf) and then imaged KVs at 6 ss (12 hpf). However, these brief treatments did not reduce KV lumen expansion (Figure 3—figure supplement 1A–C) as expected from previous work (Compagnon et al., 2014). In contrast, embryos treated with ouabain from the bud stage (10 hpf) to 8 ss (13 hpf) did show reduced KV lumen expansion (Figure 3B,E) relative to vehicle (DMSO; dimethyl sulfoxide) treated controls (Figure 3A,E). This indicated pharmacological treatments from the bud stage are more effective at blocking the lumen expansion process. Similar to wild-type embryos, mosaic labeled KV cells in control embryos treated with DMSO from bud stage to 8 ss underwent normal AP asymmetric changes in cell volume and shape (Figure 3A). KV-ant cells increased volume from 2019 ± 668 μm3 to 2304 ± 618 μm3 and KV-post cells decreased volume from 1811 ± 422 μm3 to 1479 ± 323 μm3 between 2 ss and 8 ss (Figure 3A). In contrast, AP asymmetric cell volume changes were not observed in embryos treated with ouabain between bud stage and 8 ss. In ouabain treated embryos both KV-ant and KV-post cells increased volume from 1920 ± 548 μm3 to 2328 ± 1050 μm3 and from 1819 ± 514 μm3 to 2407 ± 493 μm3 respectively (Figure 3B). These results support a model in which ion flux mediates asymmetric cell volume changes during KV morphogenesis.

Figure 3 with 3 supplements see all
Ion channel activity mediates asymmetric KV cell volume changes, KV lumen expansion and KV cell shape changes.

(A) 3D reconstructed KV-ant and KV-post cells in mosaic labeled Tg(sox17:CreERT2); Tg(ubi:Zebrabow) control embryos (treated with vehicle DMSO) showed asymmetric cell volume changes and asymmetric cell shape changes (length-to-width ratio) between 2 ss and 8 ss. (B) Inhibiting the Na+/K+-ATPase with ouabain treatments reduced KV lumen expansion and disrupted asymmetric cell volume changes. KV cells in ouabain treated embryos did not undergo asymmetric shape changes. (C–D) Interfering with Cftr function using the small molecule inhibitor CFTRinh-172 (C) or cftr MO (D) also blocked KV lumenogenesis and disrupted asymmetric cell volume changes and shape changes of KV cells. (E) Quantification of 3D reconstructed KV lumen volumes (insets depict lumen in YZ axis) in control and treated live embryos at 8 ss. For all quantitative analyses, the mean + SD is shown. The number of KV-ant and KV-post cells analyzed is indicated in the graphs in A-D. N = number of embryos analyzed. Results were pooled from two independent trials. Scale = 20 μm, *p<0.01 and ns = not significant (p>5% with Welch’s T-Test).

https://doi.org/10.7554/eLife.30963.011

As a second approach to test the role of ion flux in regulating KV cell volumes, we interfered with Cftr activity. Cftr is an apically localized chloride channel that moves Cl- ions out of the cell and establishes electrochemical gradients, which drive water into the lumen through osmosis (Navis and Bagnat, 2015). Cftr activity can also modulate several other ion-channels and transporters (Vennekens et al., 1999), making it a key driver of ion flux and fluid secretion (Braunstein et al., 2004; Valverde et al., 1995). Mosaic-labeled embryos were treated with the pharmacological compound CFTRinh-172 to inhibit Cftr activity (Roxo-Rosa et al., 2015) or a previously characterized antisense cftr morpholino oligonucleotide (cftr MO) (Gokey et al., 2016) to reduce Cftr protein expression. Treating embryos with 30 μM CFTRinh-172 from bud stage to 8 ss reduced KV lumen expansion (Figure 3C,E) as expected (Roxo-Rosa et al., 2015). To test whether Cftr has a role in modulating KV cell size, we performed a 3D analysis of mosaic labeled single cells. Similar to ouabain treatments, CFTRinh-172 treatments eliminated asymmetric volume changes in KV cells. In contrast to controls (Figure 3A), KV-post cells in CFTRinh-172 treated embryos did not lose volume, but rather increased in volume from 1878 ± 361 μm3 to 2123 ± 632 μm3. KV-ant cells increased in volume from 1513 ± 289 μm3 to 1944 ± 483 μm3 (Figure 3C). Reducing Cftr expression by injecting embryos with cftr MO had effects that were similar to CFTRinh-172 treatments: KV lumen failed to expand and asymmetric cell volume changes were disrupted (Figure 3D,E). In cftr MO treated embryos both KV-ant and KV-post cells increased their volume from 1872 ± 726 μm3 to 2252 ± 842 μm3 and from 2082 ± 637 μm3 to 2367 ± 770 μm3 respectively (Figure 3D). This suggests that in both CFTRinh-172 and cftr MO treated embryos KV-post cells fail to undergo volume loss due to inhibition of fluid efflux from these cells. Taken together (for statistical power analysis, see Figure 3—source data 1), these results indicate ion flux regulated by Na+/K+-ATPase and Cftr activity is a mechanism that drives asymmetric changes in KV cell volumes along the AP axis.

We next evaluated the impact of perturbing ion flux on cell shape changes during KV remodeling. The height (apical surfaces) of KV cells did not increase between 2 ss and 8 ss in embryos treated with inhibitors of Na+/K+-ATPase or Cftr activity as they did in controls (Figure 3—figure supplement 2A,B), which is consistent with a failure of lumen expansion. Analysis of length to width ratios (LWRs) of individual KV-ant and KV-post cells revealed that AP asymmetric cell shape changes observed in control embryos between 2 ss and 8 ss (Figure 3A) failed to occur in embryos treated with ouabain or Cftr inhibitors (Figure 3B–D). These results indicated that ion flux is necessary for both cell volume changes and cell shape changes during KV remodeling.

Since our ion flux inhibitor treatments were global, we wanted to test whether blocking ion channels altered other tissues in the embryo, including cells surrounding KV that could have an impact on KV cell shapes. Since the effect of loss of Cftr function on KV has already been determined genetically (Navis et al., 2013), we focused on the effects of ouabain treatments on external cells. The overall morphology of ouabain treated embryos was similar to controls at 8 ss, except the KV lumen was smaller (Figure 3—figure supplement 3A,B), indicating ouabain treatments did not cause severe developmental defects. To analyze cells surrounding KV, we ubiquitously expressed a membrane-localized mCherry (mCherry-CAAX) in Tg(sox17:GFP-CAAX) embryos that allowed us to simultaneously visualize both KV cells at the middle plane of KV and the surrounding external cells (Figure 3—figure supplement 3C). Due to lack of a reference frame (e.g. the lumen surface) to quantitate LWRs of surrounding cells, we used a different parameter called the ‘cell shape index, q’ (q = [(cell cross-sectional perimeter)/√(cell cross-sectional area)]) to define surrounding cell morphology (Bi et al., 2016). Analysis of external cells with clearly defined boundaries (mCherry-CAAX labeling) that were positioned either anterior or posterior of KV indicated that there was no significant difference in cell shapes at 2 ss or 8 ss stages between control or ouabain treated embryos (Figure 3—figure supplement 3D). These results indicate that ouabain does not alter the shapes of cells surrounding KV and suggest that defects in KV cell shape changes result from altered ion flux in KV. This is consistent with a previous study (Compagnon et al., 2014), in which blocking ion flux suggested forces associated with lumen expansion drive KV remodeling. However, because our 3D analyses showed that altering ion flux disrupts both lumen expansion and KV cell volume changes, it remained unclear whether failed cell shape changes were due to defects in lumen expansion or asymmetric cell volume dynamics or both.

Mathematical simulations of KV cell shape changes

To begin to tease apart how intrinsic cell size changes and extrinsic lumen expansion forces contribute to asymmetric KV cell shape changes, we developed and simulated a mathematical vertex model for cell shapes in KV. Vertex models, which have been used successfully by our group and others for predicting features of developing tissues (Fletcher et al., 2014; Bi et al., 2015; Farhadifar et al., 2007; Hufnagel et al., 2007; Wang et al., 2012), represent two-dimensional cross-sections of cells in a tissue as a network of edges and vertices, as shown in Figure 4A–D. Adhesion molecules and cytoskeletal machinery generate forces that affect cell shape in different ways. In the vertex model, the balance between these forces is represented by an effective interfacial tension parameter Λ. Positive interfacial tensions describe cytoskeletal forces that tend to decrease interface lengths, while negative interfacial tensions describe adhesion effects that tend to increase interface lengths. Additionally, cellular volume control is described in the 2D vertex model by a preferred cross sectional area A0 that the cells strive to attain. Deviations of a cell’s actual area from its preferred area correspond to cellular pressures. A ‘conjugate gradient’ computer algorithm was applied to alter the positions of the vertices based on the forces acting on them until a relaxed state is reached where all interfacial tensions are balanced by cellular pressures. Additional details about the model and the computer algorithm can be found in Materials and ethods.

Figure 4 with 4 supplements see all
Vertex model simulations for cell shapes during KV remodeling.

(A–D) Vertex model simulations with N = 10 KV-ant and KV-post cells. Upper and lower panels respectively show force-balanced states at 2 ss and 8 ss. All shown simulations start from the same initial cell positions, but the mechanical parameters differ. The full simulation box is cropped in order to focus on the KV. For the example of panel A, Figure 4—figure supplement 1B shows the respective full state. (A) Both KV and external cells are solid-like (interfacial tensions ΛKV-KV=-50 and Λext-ext=150), and the lumen cross-sectional area expands according to experimental measurements between 2 and 8 ss. (B) KV cells are solid-like (ΛKV-KV=50), external cells are fluid-like (Λext-ext=-120), and the lumen cross-sectional area expands. (C) Both KV and external cells are solid-like (ΛKV-KV=-50 and Λext-ext=150) and the lumen cross-sectional area stays constant between 2 and 8 ss. (D) KV cells are solid-like (ΛKV-KV=50), external cells are fluid-like (Λext-ext=-120), and the lumen cross-sectional area is constant. (E,F) Parameter scan for the anterior-posterior asymmetry, APA (LWR-ant - LWR-post), depending on the respective interfacial tensions of KV cells and external cells, which defines whether these cells are solid-like or fluid-like (FS = external cells fluid-like, KV cells solid-like; SS = both external and KV cells are solid-like; FF = both external and KV cells fluid-like; SF = external cells solid-like, KV cells fluid-like; hatched region = KV ant cells solid-like and KV-post cells fluid-like). For each pair of interfacial tensions, the APA was computed from the average of 100 separate simulation runs. When KV cells are solid-like, the standard error of the mean APA is typically on the order of 0.05. However, for fluid-like KV cells standard error of the mean APA can become much larger, which is reflected by the large mean APA fluctuations in this regime. (E) The lumen cross-sectional area changes normally between 2 and 8 ss. (F) The lumen cross-sectional area is fixed at a constant value between 2 and 8 ss. The parameter values corresponding to panels A-D are marked in E,F. For both E and F, a positive APA is robustly obtained only when KV and external cells are both solid-like. (G,H) Illustrations of how mechanical properties of external cells affect APA values in our simulations. For solid external cells, the interface between KV-ant and KV-post cells is prevented from moving posteriorly upon decreasing KV-post cell cross-sectional areas between 2 ss and 8 . As a consequence, the posterior KV cells flatten and obtain a smaller LWR-post value, which results in a positive APA. Conversely for fluid external cells, a decrease in KV-post cell cross-sectional area is accommodated by a posterior sliding of the interface between KV-ant and KV-post cells. Consequently, the APA does not increase and may even decrease. These mechanisms work both for increasing lumen cross-sectional area (G) and for constant lumen cross-sectional area (H).

https://doi.org/10.7554/eLife.30963.016

Using the vertex model, we studied whether the observed AP asymmetry in KV cell volume changes could act upstream of the observed AP asymmetry in KV cell shapes as described by aspect ratios. To this end, we focused our modeling efforts on the middle plane of the KV (Figure 4A–D). We first measured the cross-sectional areas of KV cells within this plane and found that changes in cell cross-sectional areas correlated with the corresponding cell volume changes (Figure 4—source data 1). Thus, to test whether KV cell volume changes can be sufficient to induce changes in KV cell aspect ratios, we prescribed the measured cross-sectional areas for KV lumen and KV cells (Figure 4—source data 2) as preferred areas A0 in our model and then studied the induced AP asymmetry of KV cell aspect ratio.

To simulate KV cell shape changes, we initialized a vertex model where a lumen is surrounded by 2N adjacent KV cells (split into KV-ant and KV-post cells with N chosen between 8 and 12) and the KV organ is surrounded by 100 ‘external’ cells. To understand how the volume changes between 2 ss and 8 ss affect cell shape, we do not take into account the full time-dependent evolution of the system. Rather, we performed quasi-static simulations consisting of two parts. Initially, preferred area values A0 for lumen, KV-ant cells, and KV-post cells were set equal to the respective 2 ss values reported in the first column of Figure 4—source data 2, and the system was relaxed to a force-balanced state using our computer algorithm (upper panels in Figure 4A–D). Subsequently, the preferred area values A0 were changed to their respective values at 8 ss chosen from the second column of Figure 4—source data 2 and the system was relaxed again (lower panels in Figure 4A–D). Such a quasi-static approach is appropriate if in the zebrafish KV, relaxation to mechanical equilibrium is faster than the volume changes of lumen and KV cells. In other developing epithelia, this relaxation timescale has been measured using laser ablation and is on the order of seconds (Fernandez-Gonzalez et al., 2009), which is significantly faster than the rate of lumen volume expansion, which is on the order of hours. Note that the images in Figure 4A–D are cropped to focus on the KV rather than the surrounding cells (same for Figure 4—figure supplements 2 and 3A–D). We show the full system for Figure 4A in Figure 4—figure supplement 1B.

There are still additional free parameters in the model, corresponding to the interfacial tension values for each cell, but these are not constrained by experimental data. Therefore, we decided to perform a wide parameter sweep to determine how these parameters affect cell shapes. In previous work (Wang et al., 2012), we demonstrated that AP asymmetric interfacial tensions were sufficient to drive KV cell shape remodeling even in the absence of asymmetric volume changes. To analyze whether asymmetric area changes alone are sufficient to drive the asymmetric KV cell shape changes, we choose the interfacial tensions between KV cells, ΛKV-KV, to be identical between KV-ant and KV-post cells. For simplicity, we assume that all cells external to the KV all have the same interfacial tension Λext-ext, which is allowed to differ from ΛKV-KV. For the purpose of illustration, we show two example simulations for different tension pairs (ΛKV-KV,Λext-ext) in Figure 4A,B. In Figure 4A with ΛKV-KV=-50 and Λext-ext=150, KV-ant cells at 8 ss appear narrow and elongated while KV-post cells appear wide and short, qualitatively reflecting the experimentally observed KV cell shape asymmetry at 8 ss. However, in Figure 4B with ΛKV-KV=50 and Λext-ext=-120, KV-ant and KV-post cells at 8 ss have very similar morphology, suggesting no cell shape change. To quantitatively compare simulations with experimental data, we developed a metric that captures the anterior-posterior asymmetry (APA) of KV cell shapes that is characteristic of KV remodeling. APA is defined as the difference between the length-width ratios (LWRs) of KV-ant and KV-post cells: APA = LWR ant – LWR-post (for the definition of the LWR, see Figure 4—figure supplement 1A). Note that because the definition of the APA is based on length-to-width ratios, it is size-independent. In our in vivo measurements, wild-type embryos at 8 ss correspond to an APA value of ~0.9. For the simulation shown in Figure 4B, we found an APA value of 0, suggesting no asymmetry, while the simulation in Figure 4A has APA value of 0.29, which is clearly asymmetric but not as high as in wild-type experiments.

Because the interfacial tensions ΛKV-KV and Λext-ext cannot be determined from our experimental data, an obvious question is whether there is any choice of those tension values in our model that would allow area changes alone to drive the observed shape changes. Figure 4E is a plot of APA as the interfacial tension in KV cells and external cells are varied. For each (ΛKV-KV,Λext-ext) parameter pair, the indicated APA value represents an average computed from 100 individual simulation runs. Blue areas indicate a positive APA corresponding to regions where KV-ant cells are more radially elongated than KV-post cells are, while red regions indicate negative APA corresponding to regions with more elongated KV-post cells. A first observation is that the APA is never above 0.31, which is much smaller than wild-type experimental observations. This suggests that changes to cross-sectional area may be an important contribution to shape remodeling, but alone they are not sufficient to generate the observed shape changes.

A second observation is that there are coherent regions in parameter space with similar values of APA, which suggests that our model may be able to identify a simple mechanism for how changes to cross-sectional area drive shape change. It has recently been discovered that as cells increase their interfacial tension Λ or increase their preferred area A0, the tissue transitions from fluid-like to solid-like behavior, undergoing a so-called rigidity transition (Bi et al., 2014, 2015; Park et al., 2015). Moreover, some of us have recently reported that a similar fluid-solid transition also occurs in bulk three-dimensional tissues (Merkel and Manning, 2017). Therefore, we expect that if we can identify a simple mechanism for 2D shape asymmetry that depends on the fluidity of the surrounding tissue, that same mechanism will also be present in 3D.

To do so, we map the results for 2D fluid-solid transitions onto our model, where black dashed lines in Figure 4E indicate phase boundaries between solid and fluid. In the upper right quadrant of Figure 4E,F, both the KV cells and external cells are solid-like (SS), in the lower right quadrant external cells are fluid-like while KV cells are solid-like (FS), in the upper left quadrant KV is fluid-like and external tissue is solid-like (SF), and in the lower left quadrant both tissues are fluid-like (FF). There is also a small, hatched region of parameter space where KV-ant cells are solid-like while KV-post cells are fluid-like. To test the significance of our APA values, we computed the standard error of the APA mean for each (ΛKV-KV,Λext-ext) parameter pair. We found that for solid-like KV cells, the error is typically on the order of 0.05, indicating that our results are robust in this regime. Conversely, for fluid-like KV cells, the standard error of the mean can become much larger, which is reflected in the higher APA fluctuations in this regime. They are a direct consequence of softer or floppier KV cells, leading to large fluctuations in their LWRs.

Interestingly, our model predicts that cross-sectional area changes drive the observed shape changes primarily in the solid-solid region, and our understanding of the fluid-solid transition helps us to understand this effect (Figure 4G). When the external cells are fluid-like, the KV cells are able to slide past the external cells, so that the interface between the KV-ant and KV-post cells (indicated by a thick black line in Figure 4G) moves towards the posterior as the area of the KV-post cells is reduced. In contrast, when the external cells are solid-like, the interface between the KV-ant and KV-post cells is pinned. In this second case, when the KV-post cells lose area they must maintain their lateral width and so the apico-basal extension must decrease. We note that this proposed mechanism works equally well in 3D as in 2D -- solid-like external tissue would pin the anterior-posterior interface so that cell volume changes would affect the area of lateral interfaces between KV cells but not the apical area in contact with the lumen. Therefore, in 3D asymmetric cell volume changes would lead to asymmetric cell shape changes and similar APA values to the ones we identified in our 2D model. Although we cannot rule out a more complex model for KV cell shape changes (e.g. with additional parameters characterizing the mechanical heterogeneities in each cell), our simple model suggests that asymmetric cell volume changes contribute to cell shape changes, though additional mechanisms are necessary to explain the very high APA values that are observed in experiments.

The number of epithelial cells in KV can vary in a wild-type population (Gokey et al., 2016), therefore we checked the robustness of our result (APA values) with respect to small changes in N. In particular, while Figure 4 shows the simulation results for N=10 KV-ant and KV-post cells, Figure 4—figure supplements 23 show the corresponding results for N=8 and N=12, respectively. In particular, independent of N, we observe positive APA only in the regime where both KV and external cells are solid-like. Moreover, there is a general trend of higher APA for a smaller KV cell number. Note that in addition to the mechanism creating the AP cell shape asymmetry illustrated in Figure 4G, which works largely independent of N, we have also discovered a quite different mechanism, which only works for small N if KV-ant cells are solid and KV-post cells are fluid (for details, see Materials and methods). In this case, KV-post cells are more easily deformed and accommodate the lumen expansion by increasing their apical lumen interface, which leads to flatter KV-post cells and thus a high APA at 8 ss (see Figure 4—figure supplement 2E). Note however that this mechanism depends on lumen expansion (compare Figure 4—figure supplement 2F).

Another benefit of the model is that we can test specific hypotheses prior to exploring them experimentally. First, to investigate whether lumen expansion is necessary to create an asymmetry in KV cell elongation, we repeated the numerical simulations shown in Figure 4A,B,E — which included both asymmetric cell cross-sectional area changes and increase in lumen cross-sectional area between 2 ss and 8 ss — except in this simulation we kept the lumen cross-sectional area fixed (Figure 4C,D,F). The APA values, shown in Figure 4F are generally smaller (max. APA value of 0.23) than those in Figure 4E (max APA value of 0.31). However, most of the regimes where both KV and external cells are solid-like still show positive APA values. These results suggest that in an environment in which cells have solid-like mechanical properties, asymmetric volume changes in KV cells can partially drive asymmetric KV cell shape changes even in the absence of lumen expansion (Figure 4H).

Second, using our model, we can explore whether heterogeneous mechanical properties of the external cells can have a significant effect on KV cell shape changes. So far, we have in our model described all external cells using the same parameters. However, given the presence of morphogenetic gradients along the AP axis within the presomitic mesoderm that include FGF and Wnt signals (Oates et al., 2012), it is plausible that the mechanical properties of the tailbud cells surrounding the KV may also show an AP-oriented gradient. Moreover, the KV is anteriorly abutting the notochord with likely different mechanical properties from the tailbud cells (Zhou et al., 2009). We thus wondered how our simulation results would depend on such heterogeneities of the external cells. To study this question, we performed simulations similar to that shown in Figure 4A where we additionally allowed all external cells on either the posterior side or the anterior side to be fluid-like (Figure 4—figure supplement 4A and B, respectively). These fluid-like subsets of the external cells have an interfacial tension of Λext-ext=-120 (as in Figure 4B). All other parameters were chosen as in Figure 4A, which included asymmetric cell cross-sectional area changes and lumen expansion. In particular, the solid-like external cells had Λext-ext=150. We found that AP cell asymmetry was much more pronounced if only the anterior external cells were solid-like (Figure 4—figure supplement 4A) than if only the posterior external cells were solid-like (Figure 4—figure supplement 4B). For the solid-like anterior external cells, the average APA computed from 100 simulations was 0.39 with a standard error of the mean of 0.03. Thus, solidity of only anterior external cells can be sufficient to introduce an asymmetry in KV cell shapes that can be slightly stronger than if all external cells were solid-like. Conversely, with solid-like external cells only in the posterior region the average APA was 0.07 with a standard error of the mean of 0.04. Thus, if there are solid-like cells present only in the posterior region, the induced AP asymmetry in KV cell shape was much weaker. Hence, asymmetry in the mechanical properties of the cells (and/or material) surrounding the KV can support asymmetric cell shape changes in KV.

Interfering with junction plakoglobin function inhibits KV lumen expansion

We were intrigued by our modeling results that predicted that given the right environment of surrounding cells, changes in KV cell volumes contribute to changes in KV cell shapes even when the lumen fails to expand. To test this prediction experimentally, we wanted to take an approach that would allow us to monitor cell shape changes in KVs in which ion flux and cell volume changes occur normally but lumen expansion is inhibited. Since coordinated remodeling of adherens junctions between epithelial cells plays important roles during lumen formation (Alvers et al., 2014), we chose to interfere with junctions between adjacent KV cells to disrupt lumen growth. The adherens junction component E-cadherin has been linked to cell junction stability and barrier function that maintains lumenal and tubular structures (Tay et al., 2013; Tunggal et al., 2005). E-cadherin is expressed in KV cells (Matsui et al., 2011; Tay et al., 2013), but loss of E-cadherin function in mutant embryos leads to early developmental defects during epiboly that preclude analysis of KV formation (Kane et al., 2005). We therefore needed tools that allow junctions to form, but with weakened integrity that allows fluid to leak out of the lumen. Transcriptome analysis of zebrafish KV cells (unpublished data) indicated that Junction plakoglobin (Jup; also called γ-catenin) is expressed in KV cells. Jup interacts in complexes at cell-cell adhesions (Fukunaga et al., 2005; Lewis et al., 1997) and is thought to link cadherins to the cytoskeleton (Kowalczyk et al., 1998; Leonard et al., 2008; Holen et al., 2012). Previous studies in cell cultures indicated Jup plays an essential role in maintaining cell-cell adhesions (Fang et al., 2014) and that perturbing Jup function results in increased epithelial permeability (Nottebaum et al., 2008). The zebrafish genome contains two jup genes, jupa and jupb. RNA in situ hybridizations confirmed jupa expression in KV cells and the population of precursor cells that give rise to KV called dorsal forerunner cells (DFCs) (Figure 5—figure supplement 1). Immunofluorescence experiments using Jup antibody (Martin et al., 2009) indicated Jupa protein is localized to lateral membranes of KV cells that are marked by GFP expression in Tg(sox17:GFP-CAAX) embryos (Figure 5A). Thus, we predicted that interfering with Jupa function would perturb KV cell-cell junction integrity such that the KV lumen would fail to expand properly.

Figure 5 with 3 supplements see all
Interfering with Junction plakoglobin inhibits KV lumen expansion.

(A) Immunostaining with Jup antibodies shows Jup enrichment at lateral membranes of KV cells marked by membrane-targeted GFP expression in Tg(sox17:GFP-CAAX) embryos. Embryos injected with jupa MO-1 showed reduced Jup protein levels. Boxes indicate enlarged regions shown as individual channels. Arrows point out representative lateral membranes. Scale = 20 μm. (B) Immunoblotting confirmed reduction in Jup protein level (arrowhead) in jupa MO-1 injected embryos relative to wild-type (WT) and control MO injected embryos. The graph shows normalized Jupa band intensities. Shown is the mean + SD for three independent experiments. (C) At 8 ss, control embryos showed an inflated KV lumen (red arrow) that was labeled using ZO-1 antibody staining. Embryos injected with jupa MO-1 to knockdown Jup expression in all cells (global knockdown) or specifically in DFC/KV cells (DFCjupa MO-1) appeared normal at 8 ss except that the KV lumen failed to expand. Interfering with Jup by injecting JUP-naxos mRNA also inhibited KV lumen expansion. Scale = 10 μm. (D) Quantification of KV lumen area in control and treated embryos at 8 ss. Co-injecting jupa MO-1 with jupa mRNA significantly rescued lumenogenesis defects. Shown are mean + SD for three independent experiments. (E) The number of ciliated KV cells was not different among the treatment groups. Shown is the mean + SD for results pooled from three independent experiments. (F) Representative images of at 8 ss in control and jupa MO injected embryos treated with vehicle (DMSO) or CFTRact-09. The graph shows KV lumen area (outlined by yellow line) in control and treated embryos. Scale = 10 μm. Shown is the mean + SD for two independent experiments.(G) Representative images of KV lumens of contro MO and jupa MO embryos injected with rhodamine-dextran. Scale = 20 μm. The graph shows percentage of embryos retaining and losing the fluorescent dye between 6 ss and 8 ss from two independent trials. N = number of embryos analyzed. *p<0.01 and ns = not significant (p>5% with Welch’s T-Test).

https://doi.org/10.7554/eLife.30963.023

To test the function of Jupa in KV morphogenesis we used an antisense morpholino (MO) to block jupa pre-mRNA splicing (jupa MO-1) or a previously reported MO that blocks translation (jupa MO-2) of jupa mRNA (Martin et al., 2009). Injection of either jupa MO efficiently reduced Jupa protein levels at KV cell junctions, as compared to embryos injected with a negative control MO (Figure 5A; Figure 5—figure supplement 2A). Reduction of Jupa expression was also confirmed using immunobloting. Jupa antibody detected a prominent band (arrowhead) around 75 kDa—consistent with Jupa proteins (~75–80 kDa) in other vertebrates (McKoy et al., 2000)—that was significantly reduced in jupa MO injected embryos (Figure 5B). Interfering with Jupa expression with MOs did not alter the gross morphology of embryos at 8 ss, but did disrupt KV lumen expansion relative to controls (Figure 5C,D) as predicted. ZO-1 immunostaining of apical tight junctions was used to assess the severity of lumen expansion defects jupa MO treated embryos (Figure 5C). Delivering jupa MO specifically to the DFCs (Amack and Yost, 2004) that give rise to KV also disrupted lumen expansion (Figure 5C,D), indicating Jup functions cell-autonomously during KV morphogenesis. Importantly, the effect on KV lumen expansion caused by MO injection was significantly rescued by co-injection of full-length jupa mRNA (Figure 5C,D; Figure 5—figure supplement 2B), indicating specificity for this phenotype. As a second approach to compromise Jupa function, we injected a human JUP mRNA with a mutation that causes naxos disease (McKoy et al., 2000). This mutant ‘JUP-naxos’ mRNA has previously been shown to encode a dominant-negative protein that interferes with Jup function in zebrafish (Asimaki et al., 2014). Similar to jupa MO treatments, expression of the JUP-naxos mRNA reduced KV lumen expansion without inducing other overt defects (Figure 5C,D). Importantly, the number of ciliated cells in KV in jupa MO and JUP-naxos mRNA treated embryos was similar to controls (Figure 5E; Figure 5—figure supplement 2C), which demonstrates that small KV lumen area was due to reduced lumen expansion rather than a reduced number of KV cells.

We next tested our prediction that loss of Jupa weakens cell-cell adhesions that are necessary for KV lumen expansion. First, we found that reducing Jupa expression moderately reduced E-cadherin enrichment (~22% decrease) along KV cell lateral domains relative to controls (Figure 5—figure supplement 3), which is consistent with previous results in cell culture studies (Fang et al., 2014). This finding suggested that although E-cadherin is maintained at levels sufficient for epiboly movements and KV formation, the cell-cell adhesions in KV might be weaker in Jupa depleted embryos than in wild-type. To test this functionally, we treated embryos with a small molecule activator of the Cftr channel (CFTRact-09) that increases Cl- ion flux and can over-inflate the KV lumen (Gokey et al., 2016). Treatment of control MO embryos with CFTRact-09 significantly increased KV lumen area (~50%) as compared to DMSO, but a similar increase was not observed in Jupa depleted embryos (Figure 5F). This suggested that fluids entering the lumen were leaking out through compromised cell-cell junctions. To test this directly, we injected a solution containing fluorescent dextran into the KV lumen. In 4 out of 5 control MO embryos the dextran remained in the lumen over a 1 hr time period. Conversely, in most Jupa depleted embryos the dextran gradually leaked out and only 1 out of 5 embryo retained significant amounts of dye (Figure 5G). Together, these results indicate Jupa functions to maintain KV cell-cell adhesion integrity that is critical for KV lumen expansion, and suggest Jupa depletion could provide a useful approach to block lumen expansion without affecting ion flux mediated cell volume changes in KV.

Asymmetric KV cell shape changes occur independent of lumen expansion

We next used Jupa depleted embryos as a tool to test our hypothesis that KV cell volume changes impact KV cell shape changes in the absence of forces exerted by the process of KV lumen expansion. Morphometric analyses of individual KV cells in mosaic-labeled Tg(sox17:CreERT2); Tg(ubi:Zebrabow) embryos treated with jupa MO-1 revealed that asymmetric KV cell volume changes occurred between 2 ss and 8 ss in Jupa depleted embryos (KV-ant cells increased volume from 2015 ± 534 μm3 to 2283 ± 414 μm3 and KV-post cells decreased volume from 1839 ± 612 μm3 to 1491 ± 310 μm3 that were similar to controls (KV-ant cells increased volume from 2108 ± 719 μm3 to 2709 ± 774 μm3 and KV-post cells decreased volume from 2273 ± 864 μm3 to 1434 ± 692 μm3) (Figure 6A,B). 3D analysis of cell shapes—assessed using LWRs—indicated that even though the lumen failed to expand (Figure 6D; Figure 6—figure supplement 1A), KV-ant and KV-post cells underwent normal asymmetric cell shape changes in Jupa depleted embryos just as observed in control embryos between 2 ss to 8 ss (Figure 6A,B). These results, which are consistent with predictions of the vertex models, provide in vivo evidence that cell shape changes can occur normally during KV remodeling in the absence of KV lumen expansion.

Figure 6 with 1 supplement see all
Asymmetric cell shape changes in KV are separable from lumen expansion.

(A) Mosaic labeled KV cells in control MO injected embryos showed asymmetric changes in cell volumes and cell shapes between at 2 ss and 8 ss. (B–C) Perturbing cell-cell junction integrity in KV by interfering with jupa (B) or lgl2 (C) expression inhibited KV lumen expansion, but asymmetric cell volume changes occurred that were similar to controls. In addition, asymmetric KV cell shape changes occurred normally in jupa and lgl2 MO embryos. (D) Quantification of 3D reconstructed KV lumen volumes (insets depict KV lumen in YZ axis) in control and treated live embryos at 8 ss. For quantitative analyses, the mean + SD is shown. The number of KV-ant and KV-post cells analyzed is indicated in the graphs in A-C. N = number of embryos analyzed. Data for control MO and jupa MO experiments are pooled from three independent experiments and lgl2 MO data are pooled from two experiments. Scale = 20 μm. *p<0.01 and ns = not significant (p>5% with Welch’s T-Test).

https://doi.org/10.7554/eLife.30963.027

To corroborate results obtained using Jupa depleted embryos, we took a second approach to inhibit lumen expansion by interfering with cell-cell adhesion. We chose to use a previously characterized MO that inhibits expression of the zebrafish Lgl2 (Lethal giant larvae 2) protein (Tay et al., 2013). Similar to Jupa depletion, loss of Lgl2 moderately reduces the accumulation of E-cadherin at lateral KV membranes and blocks KV lumen expansion (Tay et al., 2013). Analyses of mosaic-labeled KV cells in Lgl2 depleted embryos yielded results that were very similar to Jupa depleted embryos. Lgl2 depletion inhibited KV lumen expansion in mosaic labeled embryos (Figure 6D, Figure 6—figure supplement 1A), but KV cells completed normal asymmetric volume changes between 2 ss to 8 ss (KV-ant cells increased volume from 2057 ± 303 μm3 to 2329 ± 847 μm3 and KV-post cells decreased volume from 2127 ± 287 μm3 to 1617 ± 336 μm3) and normal asymmetric cell shape changes during KV remodeling (Figure 6C). Taken together (for statistical power analysis, see Figure 6—source data 1), these results are consistent with Jupa knockdown results and indicate that asymmetric epithelial cell shape changes that sculpt the KV organ are separable from the process of lumen expansion.

Discussion

The collective behavior of epithelial cells plays a key role in determining the architecture of tissues and organs. Studies of developmental processes in animal models have provided important insights into the biochemical signals and mechanical forces that regulate epithelial morphogenesis (Quintin et al., 2008; Schock and Perrimon, 2002). The zebrafish Kupffer’s vesicle (KV) is a simple organ that provides a useful model system to investigate mechanisms that regulate epithelial cell shape changes in vivo. Using a mosaic labeling approach and 3D morphometric analyses of single KV cells, we identified dynamic epithelial cell volume changes during morphogenesis that are asymmetric along the anteroposterior body axis: KV-ant cells become larger during development, whereas KV-post cells become smaller. Results from experimental perturbations (summarized in Figure 7) indicated that interfering with ion flux prevents KV lumen expansion, asymmetric changes in KV cell volume, and asymmetric changes in KV cell shape during KV remodeling. This indicated that KV cell shape changes depend on (1) lumen expansion, (2) KV cell volume changes or (3) both. Results from mathematical simulations (summarized in Figure 4G–H) indicate that mechanical properties of external tissues surrounding the KV can impact cell shape changes in the KV, and that when external tissues are solid-like, asymmetric cell volume changes in KV cells contribute to cell shape changes even in the absence of lumen expansion. Experimentally, we found that when we leave ion flux and asymmetric KV cell volume changes intact and only inhibit lumen expansion with leaky KV cell-cell junctions (summarized in Figure 7), AP asymmetric KV cell shape changes occur normally. Together, these studies identify asymmetric cell volume regulation as an intrinsic mechanism that guides cell shape changes during epithelial morphogenesis in KV. We propose this is a genetically programmed process that depends on the properties of surrounding cells, but can be separated from the biophysical forces of lumenogenesis.

Summary and working model for epithelial cell shape changes during KV morphogenesis.

Results from experiments and modeling suggest AP asymmetric cell volume changes contribute to asymmetric cell shape changes in the KV epithelium. Inhibiting ion flux blocks asymmetric cell volume changes, lumen expansion and shape changes in KV-ant (blue) and KV-post (red) cells. Vertex simulations predict that asymmetric volume (cross-sectional area) changes in KV cells can introduce AP asymmetry in KV cell shapes without lumen expansion. Consistent with this prediction, asymmetric changes in KV volume and shape occurred in the absence of lumen expansion in embryos with weakened KV cell junction integrity. These results suggest a model in which asymmetric cell volume changes contribute to cell shape changes in KV and that this process is separable from lumen growth.

https://doi.org/10.7554/eLife.30963.030

Asymmetric changes in cell size during KV epithelial morphogenesis

The finding that KV cells change volume during development is insightful for thinking about mechanisms of epithelial morphogenesis in KV since previous analyses (Compagnon et al., 2014; Wang et al., 2012) that were limited to 2D did not predict differences in KV cell size. Our previous analysis of 2D cell cross-sectional area (Wang et al., 2012) suggested cells slightly reduce their size during morphogenesis, but did not detect differences between KV-ant and KV-post cells. It is therefore striking that 3D analysis shows that KV cells do indeed change volume, and do so asymmetrically along the AP axis. We recently reported that the size of the KV organ is not under tight control during development, but rather must only exceed a size threshold to function normally during left-right patterning (Gokey et al., 2016). Thus, KV size can vary among wild-type embryos. Consistent with these findings, we observed variable KV cell sizes. However, it is clear that wild-type KV cells always change size in an asymmetric way along the AP axis. Anterior KV cells always increase their size, whereas posterior cells always decrease their size. Interfering with the asymmetry of these size changes by blocking ion flux prevents asymmetric cell shape changes that we know from previous studies (Compagnon et al., 2014; Wang et al., 2012) are critical for KV function. These results indicate the AP asymmetry of volume changes is important for KV morphogenesis and function.

The decrease in KV-post cell volume is mediated by ion channel activity that regulates fluid movement. Decrease in cell volume has also been observed during morphogenesis of zebrafish otic vesicle (Hoijman et al., 2015), where it was suggested that movement of fluids from epithelial cells into the lumen contributes to lumen expansion. It is generally thought that ion flux in epithelial cells sets up a transepithelial flow of fluids from outside the tissue into the lumen (Gin et al., 2007; Frizzell and Hanrahan, 2012). When ion flux was blocked in KV via Na+/K+-ATPase or Cftr inhibitors between the bud stage and 8 ss, KV-post cells did not shrink (but swelled) and the lumen failed to expand. This finding is consistent with a model in which intraepithelial fluid movement directly from KV-post cells into the lumen promotes lumen expansion. Since the amount of volume lost by KV-post cells does not fully account for the increase in lumen size, we propose ion flux in KV establishes both transepithelial flows and intraepithelial flows from KV-post cells to fill the lumen. Interestingly, brief treatments with the Na+/K+-ATPase inhibitor ouabain between 4–6 ss did not block lumen expansion. This may be because ouabain needs more time to penetrate deep inside the embryo to effectively block ion channel function in KV. Alternatively, these results may suggest ion channel function early in KV development (between bud stage and 4ss) is sufficient for lumen expansion or there are additional mechanisms independent of ion flux that contribute to lumen expansion and cell volume changes.

What makes KV-ant cells behave different from KV-post cells? This asymmetry likely results from a combination of intrinsic and extrinsic factors that differentially regulate KV-ant and KV-post cells. The increase in KV-ant cell size could involve cell growth. Previous studies have uncovered a role for TOR signaling in cell growth (hypertrophy) in non-dividing cells (Guertin and Sabatini, 2006). Interestingly, TOR signaling has been implicated in the morphogenesis of KV (Casar Tena et al., 2015; DiBella et al., 2009; Yuan et al., 2012). It will be interesting in future work to test for asymmetric expression/function of TOR pathway components in KV cells. Another possible intrinsic mechanism is that different KV cells develop different mechanical properties. Our previous mathematical models suggest that differential cell-cell interfacial tensions along the AP axis can generate AP asymmetric cell shape changes in KV (Wang et al., 2012). Interestingly, a recent study in Drosophila showed that contractile force induced cell shape changes are instituted via cell volume reduction (Saias et al., 2015), which indicates a link between cell volume regulation and mechanical force generation. In the KV system, it will be interesting to test in future work whether AP asymmetric volume changes result in differential cytoskeletal contractility between KV-ant and KV-post cells. Another possible contributing factor to asymmetric KV cell size is differential activation of ion channels in KV-ant and KV-post cells. For example, it is known that the Cftr localizes to the apical surface of all KV cells at all stages of KV development (Navis and Bagnat, 2015). A recent study has uncovered mechanosensitive activation of Cftr in response to membrane stretch (Zhang et al., 2010). Stretching the plasma membrane increased ion conductance and also the probability of open Cftr channels at cell membranes. During KV remodeling, apical membrane stretch in KV-post cells may lead to increased Cftr activity and higher ion-efflux with a loss of volume in these cells. An alternative possibility is that different KV cell fates (e.g. KV-ant and KV-post cells) may be determined early in development. By tracking the DFCs that give rise to KV, we have found that these cells maintain their relative spatial positions throughout KV development (Dasgupta and Amack, 2016). This suggests subpopulations of KV cells may differentiate early in development and become biochemically distinct during KV morphogenesis. Additional studies are warranted to test the hypothesis of distinct KV-ant and KV-post subpopulations of cells that have differential gene expression and/or ion channel activity.

The impact of mechanical forces on KV epithelial morphogenesis

Our vertex model simulations suggest that asymmetric volume changes are not alone sufficient to fully induce KV cell shape changes. In addition to cell-intrinsic mechanisms, biophysical forces likely guide the formation of the KV epithelium. These mechanical forces can also arise from extrinsic sources that stem from the mechanical properties of surrounding tissues or extracellular matrix (ECM) (Campàs et al., 2014; Chanet and Martin, 2014; Serwane et al., 2017; Etournay et al., 2015). Localized deposition of the ECM molecules laminin and fibronectin around anterior region of KV has been found to be important for asymmetric cell shape changes during KV morphogenesis (Compagnon et al., 2014). Interestingly, our simulations indicated that asymmetric cell shape changes are more pronounced when anterior external cells had solid-like properties and posterior external cells were fluid-like. In this case, the solid-like cells on the anterior side are able to ‘pin’ the interface between the KV-ant cells and the KV-post cells. An AP gradient of ECM, such as fibronectin (see Video 5), may help prevent neighbor exchanges and give rise to solid-like behavior only in anterior external cells. It is also possible that the solid-like ECM directly physically pins the KV-ant cells, so that the mechanism we have identified may operate even if the anterior cells beyond the ECM are fluid-like. Another possibility is that the notochord—which physically interfaces with KV-ant cells—may be a solid-like structure (Zhou et al., 2009) and this may provide the pinning mechanism. To further investigate these possibilities, important and technically challenging future work should focus on developing vertex models that interface with models for ECM fiber networks. In addition, fully three-dimensional models for confluent tissues, which have very recently been mechanically characterized (Merkel and Manning, 2017) should also be adapted for organogenesis.

Video 5
3D projection of KV in a fixed Tg(sox17:GFP) embryo stained with anti-fibronectin antibody at 8 ss.

Fibronectin (red) shows enrichment around the notochord and anterior region of the KV (green).. Anterior = top, Posterior = bottom. KV is rotating along its anteroposterior (AP) axis. Scale = 15 μm

https://doi.org/10.7554/eLife.30963.031

Other mechanisms, which remain unexplored, are mechanical forces generated by cells surrounding the KV as it advances towards the tailbud via convergent extension movements. Tissue fluidity in the tailbud plays an important role in controlling body elongation in zebrafish (Lawton et al., 2013) and may have an impact on KV. Our mathematical modeling suggests that solid-like behavior of surrounding tissue may play an important role in KV remodeling. In a previous study, we used DFC/KV specific knockdown of the Rho kinase Rock2b to test whether actomyosin contractility in KV cells vs. surrounding cells is involved in KV cell shape changes (Wang et al., 2012). KV cell shape changes failed to occur in embryos with Rock2b knocked down in KV cells, even though surrounding cells were normal, indicating that cell-autonomous actomyosin activity is important for KV cell shape changes. However, future studies are needed to explore how the mechanical properties of neighboring cells impact the establishment of asymmetric KV-ant and KV-post cell behaviors.

Lumen expansion occurs synchronously with changes in epithelial cell shapes during KV morphogenesis (Compagnon et al., 2014; Wang et al., 2012), which raises the possibility that contractile forces and/or intraluminal pressure contributes to KV cell shape changes. This idea is supported by the observation that blocking lumen expansion with ion channel inhibitors prevents KV shape changes (Compagnon et al., 2014). However, blocking ion channel activity also disrupts the previously unrecognized KV cell volume changes, making it unclear whether the lack of cell shape changes are due to reduced luminal forces or absence of cell volume changes. Our mathematical models suggest that KV cells can undergo asymmetric cell shape changes even in the absence of forces associated with lumenogenesis. This prediction was experimentally tested by perturbing KV junctions, which allowed us to block lumen expansion without altering ion channel activity or cell volume changes. 3D morphometric analyses revealed KV lumen expansion involves extension of apical surfaces of both KV-ant and KV-post cells in dorsoventral axis (represented here as cell height). But, lateral extension (represented here as cell width) happens only in KV-post cells, not in KV-ant cells due to tight packing and other mechanical influences. Interestingly, lateral extension of KV-post cells can be uncoupled from the dorsoventral extensions, which play a critical role in lumen expansion. Inhibiting lumen expansion by altering junctional integrity hinders dorsoventral expansion of apical surfaces of all KV cells, but KV-post cells still lose their volume via ion channel mediated fluid efflux and undergo lateral extension to facilitate asymmetric KV cell shape changes (Figure 6). Thus, ion channels mediate asymmetric cell shape changes via lateral extension of KV-post cells even when overall lumen expansion is inhibited. These results provide new mechanistic insight into KV epithelial morphogenesis and suggest a working model in which asymmetric KV cell shape changes depend on intrinsic ion flux-mediated fluid movements and do not depend on extrinsic forces generated by lumen expansion.

We propose that luminal forces have a nominal impact on KV cell shape changes. Previous experimental results support this idea. First, KV cells can fail to change shape even when lumen expansion is normal. Inhibiting Rock2b function or non-muscle myosin II activity had no effect on KV lumen expansion, but prevented cell shape changes during KV remodeling (Wang et al., 2011; Wang et al., 2012). This indicates that the mechanical forces generated during lumenogenesis are not sufficient to drive KV cell shape changes without active cytoskeletal contractility. Second, the degree of KV lumen expansion is highly variable in a population of wild-type embryos. Correlations between KV lumen size and KV function show that the lumen only needs to exceed a relatively low size threshold for the KV to be functional (Gokey et al., 2016). Together, these findings suggest forces exerted by expansion of the lumen play a minor role in cell shape changes during KV epithelial morphogenesis.

Cell volume changes in epithelial morphogenesis

Cell size is regulated by ion flux, but can also depend on progression through the cell cycle. Thus it is important to note that KV cells are post-mitotic epithelial cells that assemble a cilium (Amack et al., 2007). As discussed above, we consider the robust AP asymmetric changes in KV cell size that occurs with precise developmental timing (between 4 ss and 6 ss) as regulated cell volume changes that control KV organ architecture. In the zebrafish KV epithelium, we propose that cell volume changes work in concert with other mechanisms to drive KV remodeling. Other recent studies have also identified links between cell volume changes and epithelial morphogenesis (Kolahi et al., 2009; Saias et al., 2015; Hoijman et al., 2015). However, little is known about the influence of cell volume changes on cell shape regulation. As mentioned previously, during zebrafish otic vesicle development epithelial cells become thinner, suggesting intraepithelial fluid movement contributes to both lumen growth and cell/tissue shape change (Hoijman et al., 2015). This finding is consistent with studies in cell culture systems (Braunstein et al., 2004; Vázquez et al., 2001) that have shown that epithelial cells indeed undergo cellular fluid loss to regulate cell volume and cell shape. In the mouse embryo, a group of nonproliferative epithelial cells in the tooth primordium also decrease their volume and become thinner during tooth budding morphogenesis (Ahtiainen et al., 2016). Another recent study in Drosophila uncovered that during dorsal closure cells within the amnioserosa lose their volume by ~30% and change their shape (Saias et al., 2015). Additionally, in the egg chamber of Drosophila the follicle cell epithelium undergoes volume changes during oocyte development to attain distinct cell shapes (Kolahi et al., 2009). Taken together with our experimental results and mathematical models in KV, these examples suggest that cell volume change might be a common mechanism that impacts cell shape during epithelial morphogenesis in several tissues and organs.

Materials and methods

Key resources table
Reagent type (species)
or resource
DesignationSource or referenceIdentifiersAdditional information
Gene (zebrafish) junction plakoglobin a (jupa), cystic fibrosis transmembrane conductance regulator (cftr), lethal giant larvae homolog 2 (lgl2)PMID: 19101534, 23482490,23487313
Strain, strain background (zebrafish)Zebrafish (Danio rerio). Transgenic lines: this study-Tg(sox17:CreERT2), this study-Tg(sox17:GFP-CAAX), Tg(ubi:Zebrabow), Tg(sox17:GFP), this study-Tg(actb2:myl12.1-MKATE2). strain background (TAB)PMID: 23757414, 17008449See Materials and methods
Genetic reagent (zebrafish)p5E-sox17, pENTR/D_creERT2, pME-GFP-CAAX, pME-myl12.1-MKATE2, p3E-SV40-polyA, pDest-Tol2CG2, JUP-naxos.PMID: 22945937, 21138979,24920660Tol2 Kit V2, See Materials and methods
Antibodyanti-junction plakoglobin, anti-ZO-1, anti-E-cadherin, anti-acetylated tubulin, anti-GFP, anti-fibronectinPMID: 19101534, 23482490, 28315297
Commercial assay or kitTol2 Kit V2http://tol2kit.genetics.utah.edu/index.php/List_of_entry_and_destination_vectors
Chemical compound, drug4-hydroxy tamoxifen (4-OHT), ouabain, CFTRinh-172 (CFTRinh), CFTRact-09 (CFTRact)PMID: 25535919, 26442502,26432887
Software, algorithm2D vertex modelOwn code: Ph.D. thesis, Matthias Merkel, Technical University Dresden, 2014

Zebrafish husbandry and strains

Zebrafish strains were maintained using standard procedures. Wild-type TAB zebrafish were obtained from the Zebrafish International Resource Center. In addition, the following transgenic zebrafish lines were used: Tg(ubi:Zebrabow) (Pan et al., 2013), Tg(sox17:GFP) (Sakaguchi et al., 2006), Tg(sox17:GFP-CAAX)sny101 (this study), Tg(actb2:myl12.1-MKATE2)sny102 (this study), Tg(sox17:CreERT2)sny120 (this study). Embryos were staged as described (Kimmel et al., 1995).

Generation of transgenic lines

Transgene constructs were generated using the Gateway-based Tol2 kit (Kwan et al., 2007). To generate Tg(sox17:CreERT2) and Tg(sox17:GFP-CAAX) transgenics, gateway cloning was performed by combining p5E-sox17 (a generous gift from Stephanie Woo) (Woo et al., 2012), pENTR/D_creERT2 (Mosimann et al., 2011) or pME-GFP-CAAX (Tol2 Kit v2), p3E-SV40-polyA (Tol2 Kit v2), and pDest-Tol2CG2 (Tol2 Kit v2) plasmids and LR Clonase II Plus (Invitrogen). Verified constructs (25 ng/μl plasmid DNA) were injected separately with Tol2 Transposase mRNA (~25 ng/μl) into one cell stage TAB zebrafish embryos to generate Tg(sox17:CreERT2)sny120 or Tg(sox17:GFP-CAAX)sny101 F0 founders. Adult F0 animals were then crossed with wild-type fish to generate F1 heterozygotes. Tg(sox17:CreERT2)sny120 fish were then crossed with homozygous Tg(ubi:Zebrabow) (Pan et al., 2013) animals to generate a double Tg(sox17:CreERT2); Tg(ubi:Zebrabow) transgenic strain. To generate a Tg(actb2:myl12.1-MKATE2) transgenic fish, p5E-actb2 (Tol2 Kit v2), pME-myl12.1-MKATE2 (see below), and p3E-SV40-polyA (Tol2 Kit v2) plasmids were recombined into pDestTol2CG4 destination vector as described above. Wild-type TAB embryos were injected with verified constructs and Tol2 Transposase mRNA to generate Tg(actb2:myl12.1-MKATE2)sny102 F0 fish. Adult Tg(actb2:myl12.1-MKATE2)sny102 F0 fish were crossed with wild-type TAB to to generate F1 heterozygotes. Tg(actb2:myl12.1-MKATE2) fish were crossed with homozygous Tg(sox17:GFP-CAAX) animals to generate a double Tg(actb2:myl12.1-MKATE2); Tg(sox17:GFP-CAAX) transgenic strain.

Generation of pME-myl12.1-MKATE2 construct

The myl12.1 ORF was PCR amplified from cDNA pool generated from 8 ss zebrafish embryos using following primers- myl12.1F: 5′-ATTAATGGATCCATGTCGAGCAAACGCGCCAA-3′ myl12.1R: 5′-ATTAATGAATTCTGCATCGTCTTTGTCTTTGGCTC-3′. The PCR amplified myl12.1 ORF was sub-cloned into pCS2+MKATE2 vector using BamH1 and EcoR1 restriction enzymes to construct pCS2+ myl12.1-MKATE2 plasmid. The myl12.1-MKATE2 construct was PCR amplified from pCS2+ myl12.1-MKATE2 plasmid using following primers- attB1: 5′-GGGGACAAGTTTGTACAAAAAAGCAGGCTATGTCGAGCAAACGCGCCAA-3′ and attB2: 5′-GGGGACCACTTTGTACAAGAAAGCTGGGTTCATCTGTGCCCCAGTTT-3′. The PCR amplified myl12.1-MKATE2 construct was then cloned into pDONR221 vector using BP recombination to generate the middle entry pME-myl12.1-MKATE2 vector.

Whole-mount in situ RNA hybridization

A plasmid encoding full-length jupa was kindly provided by Maura Grealy’s lab (NUI, Galway) (Martin et al., 2009). It was subcloned into a pCS2+ vector and PCR amplified using following primers: jupaL- 5′-GGCTGGCCCTGTGTCCAGCC-3′ and jupaR- 5′-GTAGCCATCAAGCTCTTCAT-3′. The amplicon was TA cloned into pCRII TOPO vector and used to generate sense and antisense mRNA probes (DIG RNA labeling kit, Sigma) to detect jupa expression by in situ hybridization. RNA in situ hybridizations were performed as described (Wang et al., 2011).

Embryo injections

Morpholino oligonucleotides (MOs) were obtained from Gene Tools, LLC (Philomath, OR). We designed jupa MO-1 (5′-TTATGATTGTGTCTTCTCACCTGCA-3′) to interfere with jupa pre-mRNA splicing of exons 2 and 3. jupa MO-2 (5′-GAGCCTCTCCCATGTGCATTTCCAT-3′) designed to block jupa mRNA translation was previously described (Martin et al., 2009). Other previously characterized MOs used in this study were cftr MO (5′-CACAGGTGATCTCTGCATCCTAAA-3′) (Gokey et al., 2016), lgl2 MO-1 (5′-GCCCATGACGCCTGAACCTCTTCAT-3′) (Tay et al., 2013) and a standard negative control MO (5′-CCTCTTACCTCAGTTACAATTTATA-3′) (Gene Tools). MOs were injected into wild-type TAB embryos between the 1- and 2 cell stages. Dose curves were performed to determine optimal MO treatments: 2.5 ng of jupa MO-1, 2.5 ng of jupa MO-2, 1 ng of cftr MO (Gokey et al., 2016), 4.4 ng of lgl2 MO-1 (Tay et al., 2013) and 2.5 ng of control MO. All MOs were co-injected with 4 ng p53 MO (5′-GCGCCATTGCTTTGCAAGAATTG-3′) to diminish off-target effects as described (Tay et al., 2013). To conduct rescue experiments, pCS2+ vector containing full-length jupa was digested with NotI restriction enzyme and the linearized plasmid was used as a template to synthesize capped jupa mRNA using SP6 mMessage mMachine kit (ThermoFisher Scientific, Waltham, MA. For Jup rescue experiments, jupa MO-1 was co-injected with 75 pg jupa mRNA. A construct that encodes a dominant negative JUP-naxos protein (Asimaki et al., 2014) was a kind gift from the Saffitz Lab. To over-express the JUP-naxos protein, 120 pg of JUP-naxos mRNA was injected into 1 cell stage wild-type TAB embryos.

Fluorescent dextran injections into KV

Control MO or jupa MO-1 treated Tg(sox17:GFP-CAAX) embryos were dechorionated and mounted in 1% low melting agarose at 6 ss. KV lumens were microinjected with ~1 nL of 10 kDa dextran, Alexa Fluor-568 (Molecular Probes; Eugene, OR, Lot: 1120095) at 6 ss and imaged using a Zeiss Imager M1 microscope immediately (at 6 ss). Successfully injected embryos were then incubated at 28.5 degrees for one hour and then imaged again at 8 ss.

Immunofluorescence and microscopy

For whole mount immunofluorescent staining experiments, embryos were fixed in 4% paraformaldehyde in 1X PBS with 0.5% Triton X-100 at 4°C overnight and then dechorionated in 1X PBS. Embryos were permeabilized in blocking solution containing 1X PBS, 0.1% Triton X-100, 0.1%DMSO, and 5% goat serum for 4 hr. Primary antibodies were diluted in fresh blocking solution and incubated with embryos at 4°C overnight. Primary antibodies used: mouse anti-junction plakoglobin (1:200, BD Transduction Laboratories, San Jose, CA), mouse anti-ZO-1 (1:200, Invitrogen, Carlsbad, CA), mouse anti-E-cadherin (1:200, BD Transduction Laboratories), mouse anti-acetylated tubulin (1:200, Sigma, St. Loius, MO), mouse anti-GFP (1:200, Molecular Probes), rabbit anti-GFP (1:200, Molecular Probes) and rabbit anti-fibronectin (1:200 Sigma, F3648). Embryos were then washed in 1X PBS with 0.1% Triton X-100, 0.1% DMSO, and 1% BSA at room temperature. AlexaFluor 488- and 568- conjugated anti-rabbit and anti-mouse secondary antibodies (Invitrogen, Molecular Probes) were used at 1:200 dilutions in blocking solution overnight. Stained embryos were then washed in 1X PBS with 0.1% Triton X-100, 0.1% DMSO, 1% BSA at room temperature. Embryos were imaged using either Zeiss Imager M1 microscope or a Perkin-Elmer UltraVIEW Vox spinning disk confocal microscope. Quantification based on fluorescent immunostaining was performed using ImageJ software. KV lumen areas were measured using maximum projections of ZO-1 staining. E-cadherin levels at KV cell junctions were measured by determining the mean gray level (per pixel) along KV cell lateral membranes as described (Tay et al., 2013). This mean gray level (fluorescence intensity) was normalized to GFP intensity along lateral membranes of KV cells.

Immunoblotting

Protein extracts from approximately 30 zebrafish embryos at 8 ss were prepared as described (Martin et al., 2009). 30 μL of 2X SDS sample buffer was added and samples were boiled for 5 min. Extract from approximately 10 embryos was loaded into each lane of commercially prepared 12% gels (Bio-Rad laboratories, Hercules, CA, 456–1044) and ran at 100 V for 2 hr. Semi-dry transfers were performed at 15 V for 45 min. onto a nitrocellulose membrane (Millipore, Billerica, MA HATF00010). Membranes were blocked in blocking solution (3% BSA, 100 mM NaCl, 20 mM Tris with pH 7.6, 0.2% Tween-20 in distilled water) over night at 4°C. Membranes were cut and anti-Jup (BD Transduction Laboratories) and anti-alpha tubulin antibodies (Sigma T-6199) were used at 1:1000 dilutions in primary antibody block (0.3% BSA and tris-buffer saline with Tween-20 or TBST) and incubated at 4°C over night. Membranes were washed 4 × 15 min in TBST. Anti-mouse (Bio-Rad laboratories 166–2408) secondary antibodies were used at a 1:10,000 dilution in TBST for 2 hr at room temperature. After 4 washes for 15 min in TBST (10 mM Tris with pH 8, 150 mM NaCl, 0.05% Tween-20 in distilled water) membranes were incubated in ECL (Bio-Rad laboratories 170–5060) for 1 min and imaged on a ChemiDoc MP (Bio-Rad laboratories) imager. Band intensities were quantified using ImageJ software.

Pharmacological treatments

To induce low levels of Cre recombinase activity in Tg(sox17:CreERT2; Tg(ubi:Zebrabow) double transgenic embryos, these embryos were treated with a working concentration of 5 μM 4-hydroxy tamoxifen (Sigma) in 0.1% DMSO from the dome stage to the shield stage. To inhibit ion transport, embryos were either treated with a working concentration of 1 mM ouabain (Sigma) dissolved in water or 30 μM CFTRinh-172 (Tocris, Catalog No. 3430) in 0.1% DMSO from the bud stage to 2 ss or 8 ss. To activate Cftr channels, control MO and jupa MO injected embryos were treated with a working concentration of 10 μM CFTRact-09 (Chem Bridge, San Diego, CA) from the bud stage to 8 ss. 0.1% DMSO was used as a vehicle control for all experiments. After pharmacological treatments, embryos were thoroughly washed with embryo medium, mounted in 1% low melting agarose and imaged using either a Perkin-Elmer UltraVIEW Vox spinning disk confocal microscope or a Zeiss Imager M1 microscope.

Live imaging and morphometric analysis of KV cells

To image live KV cells, embryos were dechorionated and mounted in 1% low-melting point agarose on a glass-bottom MetTek dish at specific stages. Time-lapse imaging of KV was performed using 2 μm step-scan captured at 5 min. intervals for 105 min. using a Perkin-Elmer UltraVIEW Vox spinning disk confocal microscope. The acquired 3D datasets were processed and volume rendered using surface evolver tool in Imaris (Bitplane, Belfast, UK). Imaris was used to measure the length, width and height of reconstructed KV cells. The surface of the lumen was used to establish the axes of KV cells such that lateral axis (cell width) is parallel to the tangent of the curved lumen surface. To measure KV-lumen, KV-ant and KV-post cell cross-sectional areas, captured 3D images were oriented and maximum cross-sectional area from the middle plane perpendicular to the DV axis of individual cells were measured using clipping plane function in Imaris (Bitplane).

KV cell volume measurements

Single mosaic labeled cells (YFP+) in the KV was 3D reconstructed using ‘Create Surface’ tool in Imaris (Bitplane) software. From 3D reconstructed cells the ‘cell volume’ was measured. To measure total KV cellular volume, double transgenic Tg(actb2:myl12.1mKATE2); Tg(sox17:GFP-CAAX) embryos were used to 3D reconstruct the KV lumen and the total KV cellular component. The 3D lumen was split into equal anterior and posterior halves and the cellular component associated with the two halves of the lumen were defined as the ‘total KV-ant cellular volume’ and ‘total KV-post cellular volume.’

Analysis of cells external to the KV

55 pg of mRNA encoding a membrane-targeted mCherry (mCherry-CAAX) was injected into Tg(sox17:GFP-CAAX) embryos at the 1 cell stage. Confocal images were captured from live embryos at 2 ss and 8 ss. The cell shape index, q = [(cell cross-sectional perimeter)/√(cell cross-sectional area)] (Bi et al., 2016) was used to define morphology of cells surrounding KV at the middle plane of the KV organ in control and ouabain treated embryos at 2 ss and 8 ss. On average, 5 cells were measured from the anterior and posterior regions per embryo.

Vertex model simulation of KV

We simulate KV morphogenesis using the Vertex Model with periodic boundary conditions (Bi et al., 2015; Farhadifar et al., 2007; Fletcher et al., 2014; Hufnagel et al., 2007). Because fully 3D models introduce a larger number of variables and parameters and were not until very recently well-characterized (Merkel and Manning, 2017) we choose a two-dimensional description where each cell i is represented as a polygon with area Ai and perimeter Pi. We focus our description on the plane through the center of the KV perpendicular to the dorso-ventral axis, and represent lumen, KV-ant cells, KV-post cells, and cells external to the KV as different cell types, which may differ in their mechanical properties. We choose to have an equal number N of KV-ant and KV-post cells, respectively, and 100 external cells. Force-balanced states are defined by minima of the following effective energy functional

(1) E=12i[KA(AiA0)2+KPPi2]+ij, i<jΛijlij .

Here, the first sum is over all cells. The first term in it describes a cell area elasticity, where KA is the associated spring constant and A0is the preferred area. The second term in the first sum describes cell perimeter elasticity, where KP is the associated spring constant. The second sum in Equation (1) is over all interfaces ij between adjacent cells i and j. It accounts for the interfacial tensions between cells, where Λij denotes the interfacial tension between cells i and j and lij denotes the interface length. Note that in order to facilitate comparison with experimental data, we choose micrometers as length units for our vertex model simulations.

For our simulations, we choose the values of A0 displayed in Figure 4—source data 2. The listed values for lumen and KV cells are experimentally measured average cross-sectional areas (see Materials and methods). We assumed the external cells to be about as big as the KV cells, so we set the value of the external cells at 2 ss to the average of KV-ant and KV-post cells. The total preferred area is computed as the total sum of the preferred areas A0 of all cells at 2 ss. The preferred area of the external cells at 8 ss is chosen such that the total preferred area stays constant between 2 ss and 8 ss, which corresponds to only a small preferred area change of these cells (Figure 4—source data 2). We set KA=1000and KP=1 for all cell types. We have chosen a very high ratio (KAL2)/KP with L being a typical cell diameter in order to ensure that the measured cross-sectional area values in Figure 4—source data 2 are largely fulfilled by the cells.

The values for the line tensions depend on both involved cell types. We have set the line tension between any two KV cells i and jto the same value Λij=ΛKV-KV, independent of whether the KV cells are anterior or posterior cells. Similarly, the line tension between two external cells i and j is set toΛij=Λext-ext. Since we have no measured values for these interfacial tensions, we vary both interfacial tension parameters, ΛKV-KV and Λext-ext, in Figure 4; Figure 4—figure supplements 2 and 3. The interfacial tension between a KV cell i and an external cell j is set to the average of both homotypic interfacial tensions with an additional offset: Λij=ΛKV-ext=ΛKV-KV+Λext-ext/2+200. The tension offset serves to prevent KV cells from being extruded from the KV epithelium and to allow for a smoother basal interface between KV and external cells. Between KV cells and lumen, the interfacial tension is set to a positive value of Λijlumen-KV=100 to ensure that the lumen surface is roughly spherical.

The system is initialized using the Voronoi tessellation of a pattern of cell positions. The 2N KV cell positions are arranged equidistantly on a circle around the central lumen ‘cell’ position. The radius of this circle is computed as the estimated lumen radius plus half of the estimated KV cell height. The positions for the 100 external cells are drawn randomly from a uniform distribution with the condition of having at least a distance of lumen radius plus estimated KV cell height from the lumen cell position. Then, preferred cell areas are set to their 2 ss values and the system is relaxed by minimizing the energy functional. Afterwards, the preferred areas are set to their respective 8 ss values and the system is relaxed again. Note that the Voronoi tessellation is only used to facilitate the initialization. The subsequent energy minimizations are carried out using varying vertex positions. Also note that the dimensions of the periodic box were also allowed to vary during the minimizations. We use the conjugated gradient algorithm from the GSL library (https://www.gnu.org/software/gsl/) for the energy minimization (Press, 2007).

Computation of the KV cell length-width ratio (LWR) in the simulations

We compute the LWR of a given KV cell as the quotient of its length L divided by its width W (Figure 4—figure supplement 1). We define the width W as the distance between the midpoints of the respective interfaces with the two adjacent KV cells. The length L is defined as the distance from the midpoint of the interface with the lumen to the midpoint between points P and Q, which are the respective endpoints of the interfaces with the adjacent KV cells. In Figure 4E–F we plot the respective average LWR. During the energy minimizations, KV cells occasionally lose contact with the lumen. For the averaging, we thus only take into account the KV cells that are in still contact with the lumen.

Definition of separation of solid from fluid regimes

Earlier work on the vertex model suggested that a shape index computed from cell perimeter and area can be used to differentiate between solid and fluid regime (Bi et al., 2015). However, these simulations only studied vertex model tissues with a single cell type randomly arranged, while in our simulations, there are several cell types and a very distinct geometrical arrangement. Thus, to differentiate solid cell from fluid ones, we choose a different measurement, which is based on the actual cell perimeter Pi and the interfacial tensions Λ. Based on the interfacial tensions, one can define another parameter, which characterizes a preferred perimeter P0= -Λ/(2KP). It has been observed that fluidity also correlates with the difference between actual and preferred perimeter Pi-P0 (Bi et al., 2015). Solid vertex model tissues have PiP0>0 while fluid vertex model tissue has Pi-P0=0. Correspondingly, we use this criterion to differentiate between solid and fluid cells to define the positions of the dashed black lines in Figure 4E,F and in Figure 4—figure supplement 2E,F and, Figure 4—figure supplement 3E,F. Note that as a consequence, the positions of these lines slightly vary for different conditions.

Case of solid anterior and fluid posterior KV cells

In our vertex model we have discovered a second mechanism that can lead to a positive APA for small N, which is different from the mechanism illustrated in Figure 4G. This mechanism is at work for instance in the hatched region in Figure 4—figure supplement 2E, where anterior KV cells are solid-like and posterior KV cells are fluid-like. This difference arises even though both anterior and posterior KV cells have the same interfacial tension ΛKV-KV, because anterior KV cells have a much higher preferred area at 8ss than posterior KV cells (Figure 4—source data 2). Because the condition of fluidity in the vertex model depends on both preferred area and interfacial tension (Bi et al., 2015), there is an intermediate regime when increasing ΛKV-KV where anterior KV cells are already solid, but posterior KV cells are still fluid. When the lumen area increases in this regime, both anterior and posterior cells together have to accommodate a larger total apical interface with the lumen. However, because the anterior cells are solid while the posterior cells are fluid, the latter are more easily stretched laterally. This induces an asymmetry in cell shape that corresponds to a positive APA. Note that the effect of this mechanism appears to extend further into the region where also the posterior KV cells are solid, likely because close to the hatched region they are still more easily deformable than the anterior KV cells.

Simulations with asymmetric properties of the external cells

To simulate asymmetric properties of external cells (see Figure 4—figure supplement 4), we proceeded as before with the following changes. We divide all external cells into an anterior and a posterior subset based on the randomly drawn initial Voronoi cell positions. If the initial Voronoi position of a cell is anterior (posterior) of the initial lumen Voronoi position, we regard it as one of the anterior (posterior) external cells. The interfacial tensions between two anterior (posterior) external cells are defined by the parameter Λext,A-ext,A (Λext,P-ext,P). To set the anterior cells solid (fluid) and the posterior cells fluid (solid), we choose the parameter values Λext,A-ext,A=150 and Λext,P-ext,P=-120 (Λext,A-ext,A=-120 and Λext,P-ext,P=150). The interfacial tension between an anterior and a posterior external cell was set to the average: Λext,A-ext,P=Λext,A-ext,A+Λext,P-ext,P/2.

Percentage cell volume, cell cross-sectional area and cell height change quantifications

Percentage changes were measured using the following method: If we consider, cellular properties at 8 ss = y with standard deviation δy and cellular properties at 2 ss = x with standard deviation δx, then ‘% change (z)’ = {(y – x)/x}*100. The standard deviations can be used as the uncertainty in the measured values. Thus, the uncertainty δz in z can be represented as:

δz=100xδy2+δx2yx21/2

Statistical power analysis

For the ion channel inhibition in Figure 3A–D, we found no significant AP differences in cell volume and LWR except for the DMSO control at 8 ss as discussed in the main text, where we declared an AP difference non-significant if p>5%. To verify whether the number of measured cells was large enough to conclude that the true AP differences were smaller than for the control at 8 ss, we performed a statistical power analysis. In particular, we tested against the alternative hypothesis (H1) that the true AP average cell volume difference (or LWR difference) in a given case was the same as for the DMSO control at 8 ss. Given our measured averages and standard deviations of the control case at 8 ss, and using Welch’s t-test, we computed for all other cases the so-called type II error rate β (or false negative rate), i.e. the probability of wrongly identifying an AP difference as not significant. Results are shown in Figure 3—source data 1 and the respective statistical power correspond to 1-β. The type II error probability is always below the conventionally chosen value of 20%, and for the LWR it is always at most 3%.

For the perturbation of epithelial junctions in Figure 6A–C, we found no significant AP differences only for 2 ss. Analogous to above, we compute the type II error rate β for all 2 ss cases based on measured averages and standard deviations for the MO control at 8 ss (Figure 6A). Results are shown in Figure 6—source data 1, and the type II error rate is always at most 8%.

Statistics

The significance of pairwise differences between groups of biological data was computed by Welch’s two-tailed t-test.

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Decision letter

  1. Marianne Bronner
    Reviewing Editor; California Institute of Technology, United States

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

[Editors’ note: a previous version of this study was rejected after peer review, but the authors submitted for reconsideration. The first decision letter after peer review is shown below.]

Thank you for submitting your work entitled "Asymmetric cell volume changes regulate epithelial remodeling of the left-right organizer" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by a Senior/Reviewing Editor.

Our decision has been reached after consultation between the reviewers. Based on these discussions and the individual reviews below, we regret to inform you that your work will not be considered further for publication in eLife.

Specifically, all three reviewers were enthusiastic about many aspects of your manuscripts but also asked for rather detailed changes that are likely to take considerable time. This includes improving the illustrations, improving the quality of the results, increasing statistical significance, and improving the writing. It is the policy of eLife to reject papers if it would take longer than two months to make the necessary revisions. Therefore, we have no choice but to decline the present version of your manuscript. However, if you feel you can address the comments of the reviewers, detailed below, we would be happy to consider a new version of the paper as a new submission and every effort would be made to return the paper to the same reviewers. I hope you find the attached reviews useful in revising the manuscript.

Reviewer #1:

This paper describes experiments and a model exploring the relationships among geometric changes and the forces of morphogenesis, an understudied topic that deserves more exposure. I believe this paper could be a model of an important way to study morphogenesis. My recommendation is for this elegant series of hypothesis-driven experiments, which combine mechanics with molecular tools and mathematical hypothesis generation, to be published with some revision.

Strengths: Adding the third dimension to studies of morphology changes raises the bar in an important way. The study itself examines interactions among the players, including an epithelium, a lumen, and the surrounding cells, rather than characterizing a single component of the system, such as one ion channel. The trajectory of the paper, from observations of 3D morphology, to a model used to generate hypotheses about the underlying mechanisms including force generation and material properties, to explicit tests of those hypotheses using modern molecular techniques, is a good model of how these types of experiments can be conducted.

Weaknesses: the illustrations are not as clear as they need to be, and labels are inconsistent making it difficult to follow, particularly the first half, (up through the description of the model). The first half of the document is difficult to read in a way that I associate with an author who knows a subject so well that she or he forgets to provide the reader with the "obvious" background needed.

1) There is no discussion of total system volume. I wanted to know what is getting pushed where as the lumen increases in volume. I quickly counted the number of external cells in Figure 3A, and there were fewer at 8ss. Where did they go? It doesn't seem like the decrease in volume of posterior KV cells would be sufficient for the KV as whole to have a constant volume. A clearer explanation of the boundary conditions is needed in the text.

2) I think it is imperative that they add a description of the effect of the drugs and the MOs on the rest of the embryo, especially the external cells. That, plus the differences in the apparent extent of the KV in Figures 2A – E, also made me want to know about total volumes and relative versus absolute changes in shape and size.

3) It was not clear to me why knocking out E-cadherin directly is lethal, but knocking out its accumulation is not. Is it just a dosage effect, or is something else going on?

4) The manuscript should be revised one more time, preferably by someone not intimately familiar with the project, who can identify where and why clarity is lacking. I've made some specific suggestions below, but it needs to be revised for consistency in terms (x,y,z or l,w,h or apical-basal etc.) and illustrations (line graphs vs. bar graphs, consistency of colors – in different places red means lumen, untransformed KV cell, anterior cells only, etc.). The section describing the model, in particular, needs to include more information up front – even with long experience interpreting models, I had to go back and forth to the supplements many times before finding the background I needed to understand the model and its predictions.

Highly recommended: Since the authors have the 3D data, I was disappointed that the model was nevertheless simplified to 2D. I have to assume the authors are heading in that direction, so I was torn about whether to list this under required. I chose to recommend it highly on the basis that the model as is clearly provides useful testable predictions, which are cleverly explored experimentally in the latter half of the paper. But, it would be great if the model could be that much more representative of the actual geometry.

Reviewer #2:

The study by Dasgupta and colleagues show for the first time asymmetric volume changes in the KV and argue that this asymmetry contributes to the asymmetric cell shape changes contribute to KV cell shape changes. They also provide experiments to show that inhibiting lumen expansion does not affect cell shape changes associated with KV morphogenesis. Although asymmetric volume change represents an interesting potential mechanism, the paper is severely hampered by the quality of the results, and specifically, the huge standard deviations for some experiments. I also have some doubts about the impact of the study considering that the described process lead to change in 20% of the size of the KV and that the Amack group showed that left-right patterning occurs normally even though KV size can change vary more than 30% between embryos (see Gokey et al. 2015).

First main point of paper: There are asymmetric cell volume changes that correlate with asymmetric cell shape changes.

The presented data are not completely convincing. The reason is the approach used by the authors is not validated. For example the authors never made sure that the results are not simply due to photobleaching, changes in depth, etc. The authors should perform alternative cell volume measurements in KVs where cells are labeled in a mosaic with a membrane marker either by late mRNA injection, DNA injection or even cell transplantation and assess if the signal and therefore the cell volume measurements affected by the location of cells within the KV or by the stage of the embryos (the organ is deeper?) or by photo-bleaching (prolonged imaging). These points need to be addressed since the volume change reported here is only of about 20% between 2 and 8 ss. Overall the best would be to provide a comprehensive analysis of cell surface or volume and present convincing results (and significant data for statistical analysis) of cell volume changes throughout the KV. At present the authors are dependent on mosaic experiments where the cells location is random and we are left with data presenting huge variability with scales on the plots that are not consistent and comparable between conditions (see Figures: 1, 2 and 5).

Given that their simulations are in 2D, it might help to also perform a simpler experiment where they label cells exactly as they did in Wang 2012, and see if they see the expected differences in cross-sectional area between anterior and posterior cells. They could also see if their cross-sectional area measurements match up with their volume data (i.e. assume that the cells are columnar and calculate volume).

In addition, the authors need to clarify the following points:

- Sample size and number of independent experiments;

- Experimental design; e.g. cell volume changes between 2 and 8 ss: did they follow the same cells overtime and therefore show single cell growth or shrinkage or simply measured the volume of different cells in embryos at different stages and compared average cell volumes?

- Explain better the statistical tests and how the analysis was done; e.g. How was the cell volume change quantified?

They write in the Discussion:

"The finding that KV epithelial cells change volume during development is surprising and insightful since previous 2D analyses had not identified any statistically significant changes in KV cell size."

Their previous study showed that cells reduced their cross-sectional area during morphogenesis (Wang 2012), suggesting that cells reduced their volume, but this study shows for first time that these volume changes are asymmetric. This point should be clarified.

Second main point of paper: Inhibiting ion channels inhibits cell shape changes.

Large error bars and inconsistent scales are a problem.

Third main point of paper: In simulations, asymmetric cell volume changes could account partially for cell shape changes, provided that the mechanical properties of the cells satisfy certain conditions.

The issue about mechanical properties of cells is important. It is unclear if the anterior and posterior mechanical properties of the KV environment are different and how this contributes to KV morphogenesis. If the mechanical properties of surrounding cells are important, could they simulate what happens when there are asymmetric differences in the environment (e.g. If the anterior surrounding cells are solid like, but basal surrounding cells are fluid like, etc.), or when there are asymmetric differences in the KV cells (when anterior KV cells are solid like, but basal KV cells are fluid like, etc.).

Given that the effect of cell volume changes relies on the condition that surrounding cells are solid like, it would be important to perform an experiment to test that. It brings a different light to the Wang et al., 2012 paper, which used blebbistatin and MOs to disrupt mechanical properties of KV cells. Could some of the effects they saw be due to the changes in mechanical properties of surrounding cells rather than the KV cells themselves?

Given that their simulations show that cell volume changes alone could not account for cell shape changes, it is weird that in Figure 2 cell shape changes are completely gone.

Is cell volume change somehow a prerequisite for cell shape changes via actomyosin contractility? They suggest there could be a link in the Discussion, referencing Saias et al., 2015.

Considering the importance of actomyosin in the process of KV morphogenesis, it is important to show what happens to KV cell volume (anterior and posterior) when myosinII/rock2b is inhibited. It might be also good to check phosphorylated myosin via antibody staining (as they did in Wang 2012) to see if inhibiting ion channels somehow affects actomyosin contractility.

Repeat experiments in Figure.5 with DFC specific knockdown and possibly in lgl mutant, which is available.

Fourth main point: Lumen expansion does not affect KV cell shape change.

This is Figure 5. Once again, the huge error bars and differences between controls are problematic.

Fifth and concluding point: Inhibiting cell volume changes disrupts cell shape changes.

Here key experiments are missing. The authors never demonstrate that asymmetric (anterior-posterior) cell volume changes are necessary or contribute KV morphogenesis. The authors should directly test the impact of cell volume change on KV morphogenesis and in particular the impact of asymmetric volume change between anterior and posterior. Photomorpholinos could provide temporal and spatial specificity. If they can get it to work, blocking cftr in anterior vs. posterior cells, and blocking cftr in left vs. right cells to see if an additional axis of asymmetry arise (they could also input this into their simulations) would be necessary.

Finally, the authors need to rethink their final drawing because it is really misleading.

Reviewer #3:

The work of Dasgupta et al. uses the Kupffer's vesicle of zebrafish as a model to study the epithelial morphogenesis, in particular the possible role of programmed cell volume changes and the relationship between lumenogenesis and epithelial remodeling.

Positive aspects of the manuscript include: Methodology: the use of an elegant mosaic labelling approach to perform 3D volumetric and shape analysis of single epithelial cells in vivo. Results: (a) The description of AP asymmetry in cell volume in the KV, with A cells increasing while P decreasing volume between 2 and 8 somites (this data add to the previously described AP asymmetry in cell shape, seen in 2D, which the authors called "tissue remodeling"). (b) The finding that ion flux is required not only for KV lumen expansion (as previously described) but also for the establishment of AP polarity in cell volume. (c) The experimental separation of AP changes in cell shape and volume from lumen expansion, which indicate that the former changes might be autonomous and not a response to external forces generated during lumen expansion. Results from a-c led show that changes in KV cell volume might contribute to epithelial remodeling independent from the forces generated by lumen expansion.

My main concerns of the paper are:

Methodology:

a) The authors mix results and discussion of experimental data of cell shape (measured in 2D) with volume changes (in 3D). This is not correct and could lead to wrong interpretations. For making a clearer connection between the two, parameters of cell shape should incorporate the 3 principal axes. Also, the authors equal the term "epithelial remodeling" with the changes in cell shape in 2D. The authors should restrict the use of the term epithelial remodeling when addressing the epithelial changes in a broader manner, for example, in the introduction and discussion, and use "changes in cell shape" in the other cases.

b) The authors developed a mathematical approach using a 2D vertex model to test the relationship between changes in cell shape and volume and lumen expansion. The model works well for testing the relationship between cell shape changes and lumen expansion in 2D. However, the authors extend the interpretation of the results to the changes in cell volume, assuming that changes in cell cross sectional areas (2D) are equivalent to the changes in cell volume (3D). However, this is not necessarily the case, and thus all the conclusions and predictions that involve changes in cell volume are not valid. The authors should be very careful and make more explicit the limitations of the model. Although the use of 2D models for 3D data is in many cases useful, it is not in this particular case, where the changes of cell volume are central for the mechanisms.

Results:

It remains unclear what the main conclusions of the paper are. The authors propose that changes in cell volume work upstream of the changes in tissue remodeling but there is no direct evidence of this, nor they explore the mechanisms that could mediate this process.

Also, it remains unexplored what makes the changes in cell volume to be AP asymmetric, and whether these changes are really "programmed autonomously" or if the mechanisms that control cell volume are symmetric but modulated in an asymmetric manner by non-autonomous forces (e.g. asymmetric ECM deposition).

Finally, the lack of requirement of lumen expansion for KV tissue remodeling and changes in cell volume is an interesting finding, but again this is not explored further.

In summary, the paper provides interesting methodology and data, but in its current state has a lack of focus, which makes unclear its main contribution to our understanding of epithelial morphogenesis.

[Editors’ note: what now follows is the decision letter after the authors submitted for further consideration.]

Thank you for submitting your article "Asymmetric cell volume changes regulate epithelial morphogenesis in zebrafish Kupffer's vesicle" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by Marianne Bronner as the Senior and Reviewing Editor. The following individual involved in review of your submission has agreed to reveal her identity: Dany S Adams (Reviewer #1).

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

While the reviewers felt that the manuscript was much improved, there remain important issues that need to be addressed, as outlined below in the specific comments of reviewers 2 and 3. We hope you find their comments useful and look forward to receiving your revised manuscript.

Reviewer #1:

The clarity of the manuscript is greatly improved, and extra data that I requested has been collected, analyzed thoughtfully, and integrated. I believe it is a strong paper that makes a contribution to our understanding of KV development and, more generally, the interplay of different forces during morphogenesis. I also believe it makes a strong argument for the importance of addressing questions about morphogenetic forces and shape generation, and I think the structure of the project could serve as a useful model for how to conduct such studies. Specifically, the sequence they present, careful description of 3D morphology at multiple time points, followed by hypothesis generation (in their case using modeling), followed by experimental testing of those hypotheses, is a useful outline for these kinds of studies. I look forward to following this project, especially if these authors are able to expand their models into 3D the way they have expanded their descriptions. I recommend publication.

Reviewer #2:

The manuscript by Dasgupta et al. describes how asymmetric cell shape changes between the anterior and posterior cells of the zebrafish Kupffer's vesicle (KV). Using 3D analysis of single cells at defined single time points, the authors show that asymmetric cell shape changes in length, width and volume (but not in height, xz) occur between 4 and 6 somite-stages (ss) and that lumen expansion does not primarily contribute to asymmetric cell shape changes.

Although the conclusion from the latter part is convincing, what drives asymmetric cell shape changes is largely unclear from the 3D analysis other than involvement of ion channels. The authors emphasize that live embryos are imaged at one stage to avoid potential photobleaching. This approach is good to initially identify the crucial parameters and duration. However, to identify the mechanisms underlying asymmetric cell shape changes requires the analysis of single cell behavior continuously during the critical time points (from 4ss to 6ss) based on time-lapse movies.

There are several concerns to be clarified.

1) It is confusing about the term "asymmetric cell volume changes", which reflects asymmetric cell shape changes in length and width but not in height (Figure 2). What is the biological significance of asymmetric cell volume changes from a mechanistic point of view? Would it be all explained by changes in length and width (2D)? Also, it is ambiguous to include this term in the title.

2) Despite the fact that the authors found out the crucial time point in this process between 4 ss and 6 ss (Figure 2), the majority of their analyses have been done at 2 ss and 8 ss stages. The problem is that the experiments using ion channel inhibitors have been treated from tailbud to 2ss or tailbud to 8ss. Because of the effects of those ion channel inhibitors on height of both anterior and posterior cells at 2 ss, the authors should treat with those inhibitors from 4ss to 6ss to see if there is the effect only on the morphometric properties of posterior cells. How could ion channels mediate asymmetric cell shape changes if lumen expansion (apical expansion) were not important in this process?

3) I am uncertain about how informative the modeling part is (Figure 4). The vertex model simulations are basically done in 2D without measurements for the tensions in anterior and posterior cells and the external forces during the transition between 4ss and 6ss. The limitation is there is no data showing shape changes of a single cell in the anterior or posterior KV during this crucial transition (4ss to 6ss) based on time-lapse movies. However, I cannot assess on the mathematical modeling using solid-like or fluid-like cells that eventually leads to the conclusion that cell shape changes do not depend on lumen expansion.

4) There is inconsistency of the data upon ouabain treatment at 2ss (Figure 3B vs. Figure 3—figure supplement 2). The former shows a reduction of the lumen size, whereas the latter shows that the lumen size looks unaffected.

5) What is the biological significance of cell height changes at 2 ss in embryos treated with ion channel inhibitors or lgl2-morphant embryos? It is confusing, as cell height does not significantly change between anterior and posterior cells from 2ss to 8ss.

Reviewer #3:

The study by Dasgupta and colleagues show for the first time asymmetric volume changes in the KV and argue that this asymmetry contributes to the asymmetric cell shape changes contribute to KV cell shape changes. They also provide experiments to show that inhibiting lumen expansion does not affect cell shape changes associated with KV morphogenesis. The authors use a nice combination of experiments and modeling to assess this interesting issue. This study proposes a new process that could at work in the process of KV morphogenesis that promises to start exciting new avenues of investigation. Overall the article is a great piece of work and has been significantly improved since the last revision. We have a few comments.

1) Inhibiting ion channels inhibits cell shape changes/asymmetric cell changes in KV are separable from lumen expansion.

The statistical analysis. Ouabain treatment and cftr knock down lead to similar conclusion which is great. However, both experiments are based on the fact that the results are not significant in the treated embryos at 2SS and 8SS. For this type of analysis, the authors should provide a power analysis to demonstrate that the sample size is enough to conclude about not significant results. Similar comment for jupa and igl2 treatments.

2) Mechanical properties of the surrounding cells. In simulations, asymmetric cell volume changes could account partially for cell shape changes, provided that the mechanical properties of the cells satisfy certain conditions.

The issue about mechanical properties of cells is important. The mechanical properties of surrounding cells are important and need to be solid like at least anteriorly. Please provide reasonable explanation as to why anterior and posterior could be solid like and more generally, what are the arguments implying that mesodermal cells are solid like (this can be part of the Discussion). The 'pinning' hypothesis is interesting in that aspect and could be discussed. Lance Davidson’s work could be discussed as it seems that the notochord is a pretty rigid structure from his work. In addition, could the fact that fibronectin is enriched in the anterior side of the KV could increase the possibility that cells are solid like anteriorly?

3) Cell volume changes and actomyosin. Given that their simulations show that cell volume changes alone could not account for cell shape changes, it is weird that in Figure 3 cell shape changes are completely gone. A way to interpret these data is that ouabain does not specifically target cell volume but also affects elements modulating cell shape, such actomyosin. This raises the question of the effect of ouabain on the actomyosin network. The demonstration is that the actomyosin network is not altered by the treatment. The authors should check it with phosphorylated myosin via antibody staining (as they did in Wang 2012) to see if inhibiting ion channels somehow affects actomyosin contractility. Or is it a problem with the model?

4) The conclusion 'Asymmetric cell volume changes regulate epithelial morphogenesis'. It should be toned down providing that we do not know if the surrounding cells are solid like and that we do not know how the actomyosin network is compromised by the treatments used. The term 'regulate' seems too strong at this point.

https://doi.org/10.7554/eLife.30963.036

Author response

[Editors’ note: the author responses to the first round of peer review follow.]

Reviewer #1:

[…] Weaknesses: the illustrations are not as clear as they need to be, and labels are inconsistent making it difficult to follow, particularly the first half, (up through the description of the model). The first half of the document is difficult to read in a way that I associate with an author who knows a subject so well that she or he forgets to provide the reader with the "obvious" background needed.

Thank you for making this point. We have made significant revisions to the text and figures to improve clarity and consistency. Additional background has been added to the text and new Figure 1.

1) There is no discussion of total system volume. I wanted to know what is getting pushed where as the lumen increases in volume. I quickly counted the number of external cells in Figure 3A, and there were fewer at 8ss. Where did they go? It doesn't seem like the decrease in volume of posterior KV cells would be sufficient for the KV as whole to have a constant volume. A clearer explanation of the boundary conditions is needed in the text.

Our apologies that the manuscript did not make this clear. We do not change the

number of cells in our simulations. Therefore, in the simulations, the increase in lumen area is accommodated by a (small) decrease in the preferred area for each of the external cells. The force-balanced states depicted in former Figure 3(now Figure 4A-D) and in the related figure supplements do not represent the whole system. We cropped these images to focus on the KV itself rather than all the external cells. Therefore, the number change you observed is due to cropping. We now added the explanation of the cropping into the main text and the caption of Figure 4. In addition, we now also include Figure 4—figure supplement 1B, where the full system is depicted as an example for the case of Figure 4A. Also, Figure 4—source data 2now includes the explanation that the preferred areas of the external cells at 8 ss is chosen such that the total preferred area of the whole system stays constant. Also, we agree that it is an interesting question how the KV volume change is accommodated by the external cells in the experiments. From preliminary experiments in which we have tracked cell nuclei (expressing mCherry) in the tailbud region, it is clear that there are at least 10-20 cell diameters between the KV and the edge of the embryo and the nuclei density is not significantly disturbed, so it seems plausible that the volume change is distributed throughout the external cells and does not change much about the external cell geometry. We plan to focus on the fluidity, geometry, and mechanical properties of external cells in future work, but we think this large endeavor is beyond the scope of this manuscript.

2) I think it is imperative that they add a description of the effect of the drugs and the MOs on the rest of the embryo, especially the external cells. That, plus the differences in the apparent extent of the KV in Figures 2A-E, also made me want to know about total volumes and relative versus absolute changes in shape and size.

Thank you for making this suggestion. We have now examined the effect of the

pharmacological inhibitor ouabain on the rest of the embryo and specifically on external cells surrounding the KV. We focused on ouabain treatments since it is already known that Cftr is expressed specifically in KV cells (and not external cells around the KV) and that loss of Cftr function does not alter tissues other than KV between 2 ss and 8 ss (Navis et al., 2013). This analysis is included in new Figure 3—figure supplement 2and has been described in the Results:

“Since these treatments were global, we wanted to test whether blocking ion channels altered other tissues in the embryo, including cells surrounding KV that could have an impact on KV cell shapes. […] These results indicate that ouabain does not alter the geometry of cells surrounding KV and suggest that defects in KV cell shape changes result from altered ion flux in KV”.

We did not repeat this analysis for Jup MO and Lgl2 MO treatments (Figure 6) since KV cells (shape changes and volume changes) are normal in these treatments. The differences in the apparent extent of the KV arise from natural variations in the size of KV from embryo to embryo. We recently investigated KV variation and found that KV size can vary significantly among wild-type embryos (Gokey et al., 2016). The reason for this is that KV size is not tightly regulated, but rather only needs to exceed a certain (relatively small) threshold to function normally to establish left-right asymmetry. This source of variability has been clarified in the revised text. To corroborate size changes at the single-cell level, we measure the total volume of the KV cellular component and KV lumen at 2 ss and 8 ss stages and found asymmetric volume changes that were similar to our single cell analysis (Figure 2—figure supplement 2). We report the raw data (absolute size/volumes of single cells) at 2 ss and 8 ss stages (Figure 3 and Figure 6) from all our control and treatment groups. However, given the variability in KV size (discussed above), we have focused the manuscript on relative changes rather than absolute changes. In other words, it is not the amount of volume change, but rather the AP asymmetry of the volume changes—anterior KV cells always increase in size and posterior cells always decrease in size—that is important for KV morphogenesis.

3) It was not clear to me why knocking out E-cadherin directly is lethal, but knocking out its accumulation is not. Is it just a dosage effect, or is something else going on?

Yes, this is a dosage effect. Loss-of-function mutations in E-cadherin result in the arrest of epiboly movements of deep cells at the 70-80% epiboly stage development (8 hpf) prior to the appearance of KV (11 hpf) (Kane et al., 2005). With the MO dose we used in Jup knockdown embryos, E-cadherin enrichment at lateral KV membranes is only moderately reduced (~22% decrease) (Figure 5—figure supplement 3). These knockdown embryos complete gastrulation and appear similar to controls except for KV (Figure 5C), indicating Ecadherin is maintained at levels sufficient for epiboly movements and KV formation. However, our functional results (see Figure 5F, G) suggest loss of Jup that leads to a moderate reduction in E-cadherin expression weakens the cell-cell adhesions between KV cells relative to wild-type. This has been clarified in the text.

4) The manuscript should be revised one more time, preferably by someone not intimately familiar with the project, who can identify where and why clarity is lacking. I've made some specific suggestions below, but it needs to be revised for consistency in terms (x,y,z or l,w,h or apical-basal etc.) and illustrations (line graphs vs. bar graphs, consistency of colors – in different places red means lumen, untransformed KV cell, anterior cells only, etc.).

Thank you for these suggestions. We recruited two additional readers that have now critically evaluated our manuscript and figures and made helpful comments. We have made significant revisions to the text and figures to improve clarity and consistency. Additionally, in the schematics and in the graphs we have now consistently pseudo-colored KV-ant and KV-post cells associated with the middle plane of the organ as blue and red, respectively.

The section describing the model, in particular, needs to include more information up front – even with long experience interpreting models, I had to go back and forth to the supplements many times before finding the background I needed to understand the model and its predictions.

We have now extensively revised the modeling section and the corresponding

supplement. See below for details. We hope that clarity has significantly improved.

Highly recommended: Since the authors have the 3D data, I was disappointed that the model was nevertheless simplified to 2D. I have to assume the authors are heading in that direction, so I was torn about whether to list this under required. I chose to recommend it highly on the basis that the model as is clearly provides useful testable predictions, which are cleverly explored experimentally in the latter half of the paper. But, it would be great if the model could be that much more representative of the actual geometry.

This is an excellent point, which we carefully considered ourselves in writing this

manuscript. We are in fact headed in the direction of 3D models and we agree that it is important, but in order to use such a model for the KV, we first have to understand the simple, bulk mechanical properties of 3D models for tissues, which has not really been studied before.

Some of us have a preprint that just appeared online (Merkel and Manning, 2017), showing that there is indeed a fluid-solid transition in the 3D model. We next need to investigate whether adding additional interfacial tensions (as we’ve done here in 2D) are numerically stable, as there are some subtleties about numerical stability in 3D that we are actively working through now. Therefore, we didn’t want to use a model we do not yet fully understand here, and, as discussed below, we do not think it is necessary to explain the science.

For us, the question one must always ask with modeling is whether the mechanism that one is proposing as biologically relevant can survive realistic perturbations to the model. In this case, the mechanism we propose is that solid-like behavior of the surrounding tissue can “pin” the interface between the anterior and posterior KV cells so that it remains at the mid-plane despite changes to cell volume and cross-sectional area, and this helps to generate the observed shape changes. We have added a new figure and text to highlight this mechanism.

Now, we must ask whether this mechanism would survive a (major) perturbation to the model, which is going from 2D to 3D. We provide data that demonstrates that the 3rd dimension (e.g. the length of the cell along the dorsal-ventral axis, which we call the “height”) contributes much less differential changes to cell volume than changes in the cross-sectional areas. We have added a table Figure 4—source data 1, which shows that the AP-asymmetry in cell volumes at 8ss significantly alters the AP-asymmetry of cross sectional areas, but only marginally alters the height asymmetry. This suggests that KV remodeling in the middle plane perpendicular to the DV axis can be understood independently of the precise processes that control cellular DV extension. Specifically, one can imagine that the mechanism we have identified in 2D works the same way in 3D: when pinning the A-P interface at the middle plane, the average apical areas of the A and of the P cells are also pinned respectively. Thus, cell volume changes would directly affect the apico-basal dimension of these cells, leading to a change in aspect ratio. Of course, this relies on the fact there is solid-like pinning of the interface in 3D, and we have added a note and a reference in the modeling section to note the work from our preprint showing that solid-fluid transitions do exist in a 3D version of the model.

Reviewer #2:

The study by Dasgupta and colleagues show for the first time asymmetric volume changes in the KV and argue that this asymmetry contributes to the asymmetric cell shape changes contribute to KV cell shape changes. They also provide experiments to show that inhibiting lumen expansion does not affect cell shape changes associated with KV morphogenesis. Although asymmetric volume change represents an interesting potential mechanism, the paper is severely hampered by the quality of the results, and specifically, the huge standard deviations for some experiments. I also have some doubts about the impact of the study considering that the described process lead to change in 20% of the size of the KV and that the Amack group showed that left-right patterning occurs normally even though KV size can change vary more than 30% between embryos (see Gokey et al. 2015).

The sizeable standard deviations that can come with measuring KV cells are due to the wide variability in the size of KV from embryo to embryo. These standard deviations are important, as they accurately reflect the degree of variability in the system. We now report all raw cell volume data as an average + one standard deviation in the revised text (Results section). As noted, we recently reported a comprehensive study of the natural variation of KV size and number of ciliated cells in several wild-type and transgenic strains (Gokey et al., 2016).

Direct comparisons of KV size with left-right patterning (a readout for KV function) in individual embryos revealed that KV size only needed to exceed a certain (relatively small) threshold to function normally. Thus, we agree with the reviewer that the amount that the KV cells change in size/volume, which varies from embryo to embryo, may not be that important. However, it is clear from our results that wild-type KV cells always change size in an asymmetric way along the anterior-posterior (AP) axis. Anterior KV cells always increase in size and posterior cells always decrease in size. These asymmetric cell size changes are linked to asymmetric cell shape changes along the AP axis that we know from previous studies are critical for KV function.

We have modified the Results section and added a new paragraph to the Discussion section to clarify the point that it is the AP asymmetry rather than the absolute value of volume changes that are important for KV morphogenesis and function.

First main point of paper: There are asymmetric cell volume changes that correlate with asymmetric cell shape changes.

The presented data are not completely convincing. The reason is the approach used by the authors is not validated. For example the authors never made sure that the results are not simply due to photobleaching, changes in depth, etc. The authors should perform alternative cell volume measurements in KVs where cells are labeled in a mosaic with a membrane marker either by late mRNA injection, DNA injection or even cell transplantation and assess if the signal and therefore the cell volume measurements affected by the location of cells within the KV or by the stage of the embryos (the organ is deeper?) or by photo-bleaching (prolonged imaging). These points need to be addressed since the volume change reported here is only of about 20% between 2 and 8 ss.

We apologize that there was confusion about our methodology. We have now substantially revised the text and added new figures and movies (Figure 1 and Video 2) to clarify our approach to 3D measurements of KV cells.

We did not use an alternative labeling approach (mRNA injection, DNA injection, cell transplantation), since each approach would be subject to the same potential imaging artifacts (e.g. photobleaching or tissue depth). Instead, we took several precautions to avoid these artifacts. In our initial experiments, time-lapse imaging (Video 3) was used to identify cell dynamics during KV development. However, to avoid potential photobleaching problems, live embryos in subsequent experiments were imaged only once at one stage of development – not continuously or at multiple stages. Thus, using Figure 2Bas an example, different live embryos were imaged and analyzed at 2 ss, 4 ss, 6ss and 8 ss stages and representative cell morphologies from each stage are depicted.

To avoid differences in fluorescence signal due to differences in imaging depth, cells were not selected at random positions in the KV, but rather had to reside at the middle plane along the AP axis (defined as where the lumen diameter is largest) of the KV in all embryos analyzed. This was not made clear in our original manuscript and is now emphasized in the text. All images were also visualized in an YZ orientation (Figure 2A) to determine that signals were not affected by depth. This methodology has now been clarified in the text, figures and videos (Figure 2A and Video 2).

Overall the best would be to provide a comprehensive analysis of cell surface or volume and present convincing results (and significant data for statistical analysis) of cell volume changes throughout the KV. At present the authors are dependent on mosaic experiments where the cells location is random and we are left with data presenting huge variability with scales on the plots that are not consistent and comparable between conditions (see Figures: 1, 2 and 5).

Mosaic labeling indeed randomly labeled KV cells. By selecting enough labeled cells from several embryos, we sampled KV cells from all positions along the middle plane of KV.

This is now shown in new Figure 2—figure supplement 1. To corroborate size changes observed in randomly labeled single cells, we also measure the total volume of the KV cellular component and KV lumen at 2 ss and 8 ss stages and found asymmetric volume changes that were similar to our single cell analysis Figure 2—figure supplement 2.

We also adjusted the scales on our graphs so that they are consistent and all data are comparable from figure to figure. We have emphasized that AP asymmetry, rather than the specific amount of volume change, is important for KV morphogenesis. This has been clarified in the text and added to the Discussion.

Given that their simulations are in 2D, it might help to also perform a simpler experiment where they label cells exactly as they did in Wang 2012, and see if they see the expected differences in cross-sectional area between anterior and posterior cells. They could also see if their cross-sectional area measurements match up with their volume data (i.e. assume that the cells are columnar and calculate volume).

We did this analysis (see Figure 4—source code 1) and KV cell cross-sectional areas do correlate with KV cell volume changes. In particular, Figure 4—source code 1clearly shows that the AP volume difference at 8ss is fully accounted for by the AP difference in cross-sectional areas, but not by an AP cell height difference.

We also performed an analysis to see if the cross-sectional area matches with volume data and cell height data. As the reviewer suggests, we first thought to assume that the cells were columnar, but we quickly realized that simple assumptions like cuboidal cell shapes were not enough to account for cell volumes based on cell cross-sectional areas and cell heights, but that an effective geometrical factor was needed, which accounted for effects like cell-cell interfacial curvature. The need for such a factor was consistent with our observation of curved cell interfaces and its value was always approximately two, but the precise value varied depending on experimental stage and cell type (A/P). Thus, interpreting these results in a systematic way was difficult and so we left this computation out of the manuscript.

In addition, the authors need to clarify the following points:

- Sample size and number of independent experiments;

Thank you for pointing this out. This information has now been clarified in the figures and figure legends:

Figure 2: Results are pooled from three independent experiments. The number of cells and embryos analyzed is indicated in the graph in panel C and explained in figure legend.

Figure 3A-D: All the experiments including DMSO control, pharmacological treatments and morpholino injections were repeated two separate times at each stage (2 ss and 8 ss). The number of cells and embryos analyzed is indicated in the graphs and figure legend.

Figure 5A, B: Immunofluorescence experiments were performed three times and representative images are shown here. For immunoblotting experiment, quantitative analyses of band intensities were performed on three independent control and knockdown experiments. Average normalized band intensity data from those experiments are represented in the graph.

Figure 5C-E: Control MO and jupa MO knockdown experiments were repeated four independent times and rescue experiments were performed three independent times. DFC specific knockdown of Jup was performed two independent times. Dominant negative Jup-naxos mRNA was also injected two separate times to validate MO knockdown phenotype. This is clarified in the figure legend. The number of embryos (N) analyzed for each experiment is indicated under the graphs.

Figure 5F: Each experiment was repeated two times and the total number of embryos analyzed (N) is denoted in the graph.

Figure 5G: Each experiment was repeated two times and the total number of embryos analyzed (N) is denoted in the graph.

Figure 6A-C: Control MO and jupa MO knockdown experiments were repeated three independent times and lgl2 MO experiment was repeated two separate times at each stages (2 ss and 8 ss). The number of embryos (N) and cells analyzed are mentioned in the graphs and in the figure legend.

Figure 2—figure supplement 2: Total KV anterior and posterior cellular component volumes were measured from two independent experiments at each stage (2 ss and 8 ss) and average values are plotted. Number of embryos (N) analyzed at each stage is denoted in the graph.

Figure 3—figure supplement 2: Experiments were repeated two independent times at each stage (2 ss and 8 ss) and number of embryos (N) and cells (n) analyzed are mentioned in the graphs.

Figure 5—figure supplement 3: Control MO and jupa MO knockdown experiments were repeated two independent times and number of embryos analyzed (N) is denoted in the graph.

- Experimental design; e.g. cell volume changes between 2 and 8 ss: did they follow the same cells overtime and therefore show single cell growth or shrinkage or simply measured the volume of different cells in embryos at different stages and compared average cell volumes?

We used both approaches, but all data presented in the paper are averages that

come from pooling results obtained from imaging different cells in different embryos at different stages. This method was selected to guard against potential imaging artifacts as discussed above. Our experimental design has now been clarified in the revised text:

“To investigate the dynamics of KV cells in 3D, we first performed time-lapse imaging of mosaic labeled KVs in live Tg(sox17:CreERT2;ubi:Zebrabow) embryos from the 2-somite stage (2 ss) when the lumen first forms to the 8 somite stage (8 ss) when the lumen is fully expanded (Amack et al., 2007, Wang et al., 2012, Gokey et al., 2016). […] By analyzing enough embryos, we sampled KV cells from all positions along the middle plane of KV at different stages of development (Figure 2—figure supplement 1).”

- Explain better the statistical tests and how the analysis was done; e.g. How was the cell volume change quantified?

We have modified our images and now show the raw data (previously represented in the supplementary images) of KV cell volumes at 2 ss and 8 ss instead of percentage volume changes (Figure 3A-D, Figure 6A-C). In wild-type embryos at 2 ss the KV-ant and KV-post cells have very similar volumes and cell shapes. This changes at 8 ss where KV-ant cells increase their volume and KV-post cells decrease their volume and become significantly asymmetric. These asymmetric cell volume changes result in asymmetric in cell shapes. Inhibiting ion flux perturbs asymmetric cell volume changes and asymmetric cell shape changes. This has been clarified in the main text.

They write in the Discussion:

"The finding that KV epithelial cells change volume during development is surprising and insightful since previous 2D analyses had not identified any statistically significant changes in KV cell size."

Their previous study showed that cells reduced their cross-sectional area during morphogenesis (Wang 2012), suggesting that cells reduced their volume, but this study shows for first time that these volume changes are asymmetric. This point should be clarified.

We have revised the Discussion to clarify this point:

“The finding that KV epithelial cells change volume during development is insightful for thinking about mechanisms of KV morphogenesis since previous analyses (Compagnon et al., 2014, Wang et al., 2012) that were limited to 2D had not predicted any differences in KV cell size. […] It is therefore a striking that 3D analysis shows that KV cells do indeed change volume, and do so asymmetrically along the AP axis.”

Second main point of paper: Inhibiting ion channels inhibits cell shape changes.

Large error bars and inconsistent scales are a problem.

As discussed above, the error bars represent the natural variation in KV observed in a population of embryos. Even with variability, sampling enough embryos allowed us to identify statistically significant differences. We have changed the scales in our graphs such that they are now consistent throughout the paper.

Third main point of paper: In simulations, asymmetric cell volume changes could account partially for cell shape changes, provided that the mechanical properties of the cells satisfy certain conditions.

The issue about mechanical properties of cells is important. It is unclear if the anterior and posterior mechanical properties of the KV environment are different and how this contributes to KV morphogenesis. If the mechanical properties of surrounding cells are important, could they simulate what happens when there are asymmetric differences in the environment (e.g. If the anterior surrounding cells are solid like, but basal surrounding cells are fluid like, etc.).

Thank you for this suggestion. We now additionally studied the influence of

asymmetries in the external cells. As may have been expected from our proposed mechanism (that solid-like cells “pin” the interface between the anterior and posterior cells at the middle plane during volume changes), we find that it does make a difference whether only anterior or only posterior external cells were solid or all external cells were solid.

In particular, if only posterior external cells were solid (Figure 4—figure supplement 4B), the AP asymmetry in KV cell shape was much weaker than if only anterior external cells were solid (Figure 4—figure supplement 4A), because they do a poor job of “pinning” the interface between anterior and posterior cells. In contrast, when only anterior external cells are solid-like, the pinning works well and the APA is very slightly larger than in the case where all external cells were solid. We now added a discussion of these simulations to the main text.

In future work, we are planning to extensively study the fluidity, geometry, and mechanical properties of the external cells in more detail, in order to better constrain their effects on KV mechanics, but we believe this is beyond the scope of the current manuscript.

Or when there are asymmetric differences in the KV cells (when anterior KV cells are solid like, but basal KV cells are fluid like, etc.).

The case of solid anterior KV cells and fluid posterior KV cells is already included in our simulations. For small numbers of cells, the APA is also positive. We have added a brief discussion of this case to the main text and a more extensive discussion to a dedicated section in the supplement.

Given that the effect of cell volume changes relies on the condition that surrounding cells are solid like, it would be important to perform an experiment to test that. It brings a different light to the Wang et al., 2012 paper, which used blebbistatin and MOs to disrupt mechanical properties of KV cells. Could some of the effects they saw be due to the changes in mechanical properties of surrounding cells rather than the KV cells themselves?

This is an interesting point. Our previously reported blebbistatin treatments and

global MO knockdowns of the Rho kinase Rock2b (Wang et al., 2012) likely impact the mechanical properties of both KV cells and surrounding cells. We previously did an experiment to test whether actomyosin contractility in KV cells vs. surrounding cells is involved in KV cell shape changes (Wang et al., 2011). To test this, MOs against Rock2b were injected into midblastula stage embryos for DFC/KV specific knockdown. KV cell shape changes failed to occur in embryos with Rock2b knocked down in KV cells, even though surrounding cells were normal. This indicates that cell-autonomous function of Rock2b and actomyosin activity is important for KV cell shape changes. Of course, this doesn’t rule out a role for surrounding cells, and future studies are needed to test this possibility. This point is now addressed in the Discussion.

Given that their simulations show that cell volume changes alone could not account for cell shape changes, it is weird that in Figure 2 cell shape changes are completely gone.

Is cell volume change somehow a prerequisite for cell shape changes via actomyosin contractility? They suggest there could be a link in the Discussion, referencing Saias et al., 2015.

Considering the importance of actomyosin in the process of KV morphogenesis, it is important to show what happens to KV cell volume (anterior and posterior) when myosinII/rock2b is inhibited. It might be also good to check phosphorylated myosin via antibody staining (as they did in Wang 2012) to see if inhibiting ion channels somehow affects actomyosin contractility.

We agree that is interesting and important to tease apart the relationships among ion flux, cell volume changes and cytoskeletal contractility. The suggested experiments are excellent and we plan to pursue this line of investigation in future work. As suggested, there are several possible outcomes. First, as mentioned in our Discussion section, contractile forces that generate cell shape changes may be directly linked to cell volume changes. In the KV system, it will be interesting to test whether AP asymmetric volume changes result in differential cytoskeletal contractility between KV-ant and KV-post cells. To do this, we are generating fluorescent reporters of actomyosin dynamics in KV cells to analyze this in real time.

A second possibility is that actomyosin contractility is necessary for cells to change volume. Here, we can use treatments (e.g. blebbistating, Rock2b MO, and mypt1 mRNA) to perturb contractility and then assay volume changes. In addition, we are building genetic tools to address this possibility. Since we have established the sox17:CreERT2 transgenic line, we are using CRISPR to introduce loxP sites into cytoskeletal regulatory genes such as Rock2b. This system would allow us to delete Rock2b specifically in KV and then follow contractility, volume changes and shape changes.

A third possibility, as pointed out, is that ion flux might influence contractility independent of volume changes. This may be difficult to separate out, but we will use antibody staining and fluorescent reporters to assess contractility in KV at early stages prior to volume changes.

Finally, it is possible that the relationship between cell volume changes, ion flux and cytoskeletal contractility is not linear, but rather more complex and interdependent on one another. Additional experiments and potentially new mathematical models will be needed if this is the case.

Of note, in response to this comment, we did few blebbistatin treatments in mosaic labeled embryos to get an idea of what might be happening. Preliminary results were inconsistent, suggesting either technical problems (variable drug efficacy or wrong dose) or that the relationship between contractility and cell volume is complex (see possible outcomes discussed above).

We feel that the experiments proposed to test the interplay between cell volume changes and cytoskeletal contractility represent a new line of investigation that goes beyond the scope of this paper and any new results would not change the conclusions of the present manuscript.

Repeat experiments in Figure 5 with DFC specific knockdown and possibly in lgl mutant, which is available.

As suggested, we used DFC/KV-specific knockdown to determine whether Jup

functions cell-autonomously in KV cells to regulate KV lumen expansion. Similar to global Jup knockdowns, the DFC/KV-specific knockdowns resulted in a reduced KV lumen size without affecting the number of ciliated KV cells. These new data, presented in Figure 5C-E, indicate Jup functions in KV cells cell-autonomously to regulate KV lumen expansion. This has been added to the Results section.

We did not repeat experiments in old Figure 5(which is now Figure 6) with DFC/KV-specific knockdowns for two reasons: 1) global and DFC/KV-specific knockdowns have the same effect/phenotype (Figure 5C) and 2) KV cell behaviors (shape changes and volume changes) in knockdown embryos are normal, similar to control embryos (Figure 6A-C). If we had observed a defect in KV cell shape or volume changes in global knockdowns, we would want to test whether this was a cell-autonomous effect using DFC/KV-specific knockdowns. But this is not the case here.

Unfortunately, available lgl2 mutants are not useful for KV studies. As previously described (Sonawane et al., 2005) lgl2 mutants do not have phenotypes during early development (prior to 4 days post-fertilization) due to maternal contribution. Only MOs that target both maternal and zygotic expression develop KV phenotypes (Tay et al., 2013).

Fourth main point: Lumen expansion does not affect KV cell shape change.

This is Figure 5. Once again, the huge error bars and differences between controls are problematic.

A discussion of error bars and differences among control embryos appears above

and is now included in the revised text.

Fifth and concluding point: Inhibiting cell volume changes disrupts cell shape changes.

Here key experiments are missing. The authors never demonstrate that asymmetric (anterior-posterior) cell volume changes are necessary or contribute KV morphogenesis.

Our work with mosaic labeled cells has uncovered asymmetric volume changes

along the AP axis of KV that occur at the same developmental timing as AP asymmetric cell shape changes that were previously described. A previous study that used ouabain treatments in inhibit ion flux and lumen expansion (Compagnon et al., 2014) predicted that the forces associated with rapid KV lumen growth impacts asymmetric KV cell shape changes. Here, we show that in addition to blocking lumen expansion, inhibiting ion flux also blocked asymmetric changes in KV cell volume. Thus, the failure of cells to change shape when ion flux was altered could be due to blocked lumen expansion or blocked volume changes or both.

To test this, we blocked lumen expansion by creating a leaky KV. In these embryos (Figure 6), asymmetric volume changes occurred normal in KV even though the lumen didn’t expand. KV cells also completed normal cell shape changes in the absence of normal lumen expansion.

From these experiments, we conclude that asymmetric cell shape changes that occur in KV depend on ion flux-mediated cell volume changes and do not depend on extrinsic mechanical forces associated with lumen expansion. We also used vertex models to test whether changes in cell volume (cross-sectional area) contribute to KV cell shape changes. This has been clarified in the Results section:

“First, to investigate whether lumen expansion is necessary to create an asymmetry in KV cell elongation, we repeated the numerical simulations shown in Figure 4A, B, E – which included both asymmetric cell cross-sectional area changes and increase in lumen cross-sectional area between 2 ss and 8 ss – except in this simulation we kept the lumen cross-sectional area fixed (Figure 4C, D, F). […] These results suggest that in an environment in which cells have solid-like mechanical properties, asymmetric volume changes in KV cells can partially drive asymmetric KV cell shape changes even in the absence of lumen expansion (Figure 4H).”

Taken together, these results indicate asymmetric cell volume changes contribute to asymmetric cell shape changes in KV. These results have been emphasized for clarity in the revised manuscript.

The authors should directly test the impact of cell volume change on KV morphogenesis and in particular the impact of asymmetric volume change between anterior and posterior. Photomorpholinos could provide temporal and spatial specificity. If they can get it to work, blocking cftr in anterior vs. posterior cells, and blocking cftr in left vs. right cells to see if an additional axis of asymmetry arise (they could also input this into their simulations) would be necessary.

Whether volume changes in anterior KV cells or posterior KV cells make equal

contributions to overall KV morphogenesis is very interesting question that we plan to address in future work. In addition to the suggested photo-MO experiments, we are currently generating genetic tools to test the contributions of asymmetric volume changes. We have identified a handful of genes that are asymmetrically expressed in KV– either in anterior cells or posterior cells – that we are using to generate transgenic lines that can overexpress or interfere with ion channels. As suggested, we plan to integrate results from these experiments with our models to fully test the contribution of asymmetric volume changes to KV form and function. We agree with the reviewer and predict that asymmetric ion channel activity is driving asymmetric volume changes. However, we feel that rigorously testing this hypothesis is beyond the scope of this paper.

Finally, the authors need to rethink their final drawing because it is really misleading.

We have significantly simplified Figure 7and have clarified that this is a working

model. In this model, cell volume changes (represented in 2D by changes in cell cross-sectional area) introduce cell shape changes along the AP axis of the KV in wild-type embryos. Perturbations of ion flux inhibit lumen expansion, prevent cell volume (or cell cross-sectional area) changes and prevent KV cell shape changes. Weakening junction integrity inhibited normal lumen expansion, but asymmetric cell volume and cell shape changes occurred normally, albeit to a lesser extent compared to wild type.

Reviewer #3:

[…] My main concerns of the paper are:

Methodology:

a) The authors mix results and discussion of experimental data of cell shape (measured in 2D) with volume changes (in 3D). This is not correct and could lead to wrong interpretations.

We have worked to ensure that we more clearly separate now between our

experimental findings about the 3D KV and our 2D modeling of a cross section through the center of the KV along a plane that is perpendicular to the dorso-ventral (DV) axis. We also explain more clearly now that volume changes are correlated with cross-sectional area changes, and that our simulations show that asymmetric changes in the KV cell areas can induce asymmetric KV cell shape changes.

For making a clearer connection between the two, parameters of cell shape should incorporate the 3 principal axes.

Cell parameters for all 3 axes (L, W and H) were measured using 3D rendered cells and are reported in the paper. We chose to use a length to width ratio (LWR) to describe cell shapes for two reasons: 1) to make connections with our previous work (done in 2D) that used LWRs to describe cell shape changes (Wang et al., 2011, Wang et al., 2012) and 2) to make connections with simulations using vertex models. In our simulations, we quantify cell shape anisotropy based on a LWR, where the length corresponds to the apico-basal extension of the cells, and the width corresponds to the lateral extension perpendicular to the DV axis. Thus, the LWR quantified in the simulations corresponds to the LWR quantified in the experiments.

Since the 2D simulations focus on a plane perpendicular to the DV axis, the cell extension along the DV axis (called “height”) is not included in the modeling. Please see our response to reviewer 1 Point #6 for a detailed discussion of our justification for this choice. We have also edited the paper significantly to address this point.

Also, the authors equal the term "epithelial remodeling" with the changes in cell shape in 2D. The authors should restrict the use of the term epithelial remodeling when addressing the epithelial changes in a broader manner, for example, in the introduction and discussion, and use "changes in cell shape" in the other cases.

We agree and have reformulated. We now are explicit about ‘changes in cell shape’ and ‘changes in cell volume.’

b) The authors developed a mathematical approach using a 2D vertex model to test the relationship between changes in cell shape and volume and lumen expansion. The model works well for testing the relationship between cell shape changes and lumen expansion in 2D. However, the authors extend the interpretation of the results to the changes in cell volume, assuming that changes in cell cross sectional areas (2D) are equivalent to the changes in cell volume (3D). However, this is not necessarily the case, and thus all the conclusions and predictions that involve changes in cell volume are not valid. The authors should be very careful and make more explicit the limitations of the model. Although the use of 2D models for 3D data is in many cases useful, it is not in this particular case, where the changes of cell volume are central for the mechanisms.

We agree that one has to be careful when studying the effect of cell volume changes using a 2D model. However, we have explicitly measured the changes in cell volume and cell cross-sectional area in the plane perpendicular to the dorsal-ventral axis (e.g. where our 2D model resides) as well as the “height” of cells – their length along the DV axis. These are shown in the main text in Figure 2Cand Figure 4—source data 1 and 2. These data demonstrate that differential cell volume changes between anterior and posterior cells correlate with cell cross-sectional area changes while heights of anterior and posterior cells remain approximately same. This gives us confidence to base our 2D simulations on the change of the cell cross-sectional areas. We have clarified this in the main text. Please also see our detailed response to reviewer 1 Point#6 for a discussion of why we expect that the mechanism identified by our 2D model is valid in 3D.

Results:

It remains unclear what the main conclusions of the paper are. The authors propose that changes in cell volume work upstream of the changes in tissue remodeling but there is no direct evidence of this, nor they explore the mechanisms that could mediate this process.

In this work, we found experimental conditions that separate the process of lumen expansion (and its associated biophysical forces) from the process that drives cell shape changes in KV. We conclude that ion flux and its associated asymmetric volume changes contribute to cell shape changes during epithelial morphogenesis in KV. We have explored possible mechanisms of how asymmetric KV cell volume changes create asymmetric KV cell shapes. However, we might not have explained this clearly enough.

To improve our manuscript, we have modified the main text to more clearly explain our conclusions and the mechanism that is responsible for asymmetric KV cell shapes in the simulations when the external cells are solid. In addition, we created illustrations of these mechanisms for both presence and absence of lumen expansion (Figure 4G, H). Moreover, we added to the main text a brief explanation of another mechanism that can create asymmetric KV cell shapes in the case where anterior KV cells are solid, posterior KV cells are fluid, and there is only a small number of KV cells. A more detailed discussion of this second mechanism was added to the supplement.

Also, it remains unexplored what makes the changes in cell volume to be AP asymmetric, and whether these changes are really "programmed autonomously" or if the mechanisms that control cell volume are symmetric but modulated in an asymmetric manner by non-autonomous forces (e.g. asymmetric ECM deposition).

Finally, the lack of requirement of lumen expansion for KV tissue remodeling and changes in cell volume is an interesting finding, but again this is not explored further.

As discussed above in response to reviewer #2, we are very interested in

determining how KV cell volumes change in an asymmetric way and how this asymmetry contributes to KV form and function. We are generating several genetic tools to perturb volume changes and test whether they are cell-autonomous. We will also perform experiments and simulations to test the role of external forces that include ECM and the mechanical properties of surrounding cells. Multiple approaches will be needed to tease apart contributions from cell autonomous and non-cell autonomous effects, which we feel is beyond the scope of this paper.

In summary, the paper provides interesting methodology and data, but in its current state has a lack of focus, which makes unclear its main contribution to our understanding of epithelial morphogenesis.

Thank you for this feedback. We have revised the manuscript text to more sharply focus on the goals and the main contributions of this study. We set out to develop new mathematical models and experimental methods that allow morphometric analysis of epithelial cells in the simple KV organ in order to investigate the interplay between intrinsic and extrinsic mechanisms that contribute to cell shape changes during epithelial morphogenesis. Using this combination of modeling and experimental approaches, we uncovered new mechanistic insights that contribute to our understanding of epithelial morphogenesis:

1) Epithelial cells in KV undergo volume changes that are asymmetric along the AP axis during morphogenesis. KV-ant cells increase in volume and KV-post cells decrease in volume. These AP asymmetric volume changes coincide with AP asymmetric cell shape changes.

2) Ion flux is an intrinsic mechanism that regulates asymmetric cell volume changes and cell shape changes in KV.

3) Mathematical models indicate that mechanical properties of external cells surrounding the KV can impact cell shape changes in the KV. Models predicted that when external cells are solid like, asymmetric cell volume changes in KV cells contribute to cell shape changes even in the absence of lumen expansion. In particular, we have uncovered a possible mechanism for KV cell shape change: if the interface between KV-ant and KV-post cells was pinned, the volume changes directly affects the apico-basal dimension of the cells while the apical area would remain constant. Hence, asymmetric cell volume changes can induce asymmetric cell shape changes.

4) Experiments determined that asymmetric cell volume and shape changes are separable from extrinsic biophysical forces associated with lumen expansion.

Taken together, these results demonstrate ion flux serves as an intrinsic mechanism to regulate asymmetric epithelial morphogenesis in the KV organ and that changes cell morphology can be uncoupled from mechanical forces exerted during lumen expansion.

These take-home points have been clarified throughout the revised text.

[Editors' note: the author responses to the re-review follow.]

Reviewer #2:

[…] Although the conclusion from the latter part is convincing, what drives asymmetric cell shape changes is largely unclear from the 3D analysis other than involvement of ion channels. The authors emphasize that live embryos are imaged at one stage to avoid potential photo-bleaching. This approach is good to initially identify the crucial parameters and duration. However, to identify the mechanisms underlying asymmetric cell shape changes requires the analysis of single cell behavior continuously during the critical time points (from 4ss to 6ss) based on time-lapse movies.

Thank you for your comments and this suggestion. We have now performed time-lapse imaging of mosaic labeled KVs between 4 ss and 6 ss and quantified volume dynamics of single cells at 5 minute intervals during this critical time point. This has allowed us to track and quantify changes in KV-ant and KV-post cell morphology in real time. Consistent with our results from analyzing ‘snap shots’ at specific developmental stages (e.g. 2 ss, 4 ss, 6 ss or 8 ss), we found that individual KV-anterior cells increase their volume between 4-6 ss and KV-posterior cells decrease their volume. This live imaging approach sheds light on KV cell behavior and how these cells change their morphology during morphogenesis. Our new results are presented in Figure 2—figure supplement 2 and Video 4.

There are several concerns to be clarified.

1) It is confusing about the term "asymmetric cell volume changes", which reflects asymmetric cell shape changes in length and width but not in height (Figure 2). What is the biological significance of asymmetric cell volume changes from a mechanistic point of view? Would it be all explained by changes in length and width (2D)? Also, it is ambiguous to include this term in the title.

We use the term ‘asymmetric cell volume changes’ to describe regional changes in KV cell size along the anterior-posterior (AP) body axis: KV-anterior cells increase in volume over developmental time, whereas KV-posterior cells decrease in size. We apologize for the confusion, and have now clarified this in the Results section of the manuscript (subsection “3D analysis of single cells reveals asymmetric cell volume changes during KV morphogenesis”, last paragraph).

Changes in cell volume are not necessarily tied to changes in cell shape: cells can change volume but remain the same shape or change shape without changing volume. Results presented in this paper indicate KV cell volume changes contribute to KV cell shape changes. Inhibitors of ion flux inhibit lumen expansion and AP asymmetric KV cell volume changes. This indicates ion flux, which drives fluid movements, is necessary for both lumen expansion (previously published) and KV cell volume changes (not surprising and consistent with a role for ion flux in cell volume changes in other systems). When we looked at KV cell shape changes by length and width (2D) in embryos treated with ion flux inhibitors, they did not occur. This indicates that KV cell shape changes depend on 1) lumen expansion, 2) KV cell volume changes or 3) both. Mathematical modeling suggested that KV cell shapes could change if KV cell volume changes occurred without lumen expansion. We next tested this experimentally. When we leave ion flux and asymmetric KV cell volume changes intact and only inhibit lumen expansion, KV cell shape changes (measured by length and width in 2D) occur normally. Taken together, this suggests that KV cell shape changes do not depend on full lumen expansion, but do depend on asymmetric KV cell volume changes. Thus, mechanistically, KV cell volume changes are required (drive) for cell shape changes. This has been clarified in the first paragraph of the Discussion section.

As suggested, we have changed the title to “Cell volume changes contribute to epithelial morphogenesis in zebrafish Kupffer’s vesicle.”

2) Despite the fact that the authors found out the crucial time point in this process between 4 ss and 6 ss (Figure 2), the majority of their analyses have been done at 2 ss and 8 ss stages. The problem is that the experiments using ion channel inhibitors have been treated from tailbud to 2ss or tailbud to 8ss. Because of the effects of those ion channel inhibitors on height of both anterior and posterior cells at 2 ss, the authors should treat with those inhibitors from 4ss to 6ss to see if there is the effect only on the morphometric properties of posterior cells.

Thank you for suggesting this experiment to test the effect of inhibiting ion channels during the critical stages of 4 to 6 ss. Our rationale for starting pharmacological treatments at the tailbud stage was to allow the compounds to penetrate inside the embryo and block ion channel function in KV throughout the lumen expansion process. We have now repeated the mosaic-labeling experiments with ouabain treatments that started at 4 ss and then imaged KVs at 6 ss. Over the course of several trials, we found, unfortunately, that treatments between 4-6 ss did not reduce KV lumen expansion (see Author response image 1 for a typical result). This indicates longer treatments are indeed necessary to effectively block ion channels, and therefore we are unable to assess the effect of acutely blocking ion flux between 4-6 ss.

Author response image 1
Ouabain treatments between 4-6 ss do not block KV lumen expansion.

(A-B) Mosaic-labeled KV in a control embryo (A) and an embryo treated with ouabain from 4ss to 6ss (B). (C) Measurements of maximum KV lumen area indicated that ouabain treatments between 4-6 ss were not effective at blocking KV lumen expansion. N=number of embryos analyzed. Error bars=one standard deviation. ns=not significant, (Welch’s T-Test).

https://doi.org/10.7554/eLife.30963.034

How could ion channels mediate asymmetric cell shape changes if lumen expansion (apical expansion) were not important in this process?

We apologize for not making this clear. In wild-type embryos, KV lumen expansion involves extension of apical surfaces of both KV-ant and KV-post cells in dorsoventral axis (represented here as cell height, Figure 2C). But, lateral extension (represented here as cell width,) happens only in KV-post cells, not in KV-ant cells due to tight packing and other mechanical influences. Interestingly, lateral extension of KV-post cells can be uncoupled from the dorsoventral extensions, which play a critical role in lumen expansion. Inhibiting lumen expansion by altering junctional integrity hinders dorsoventral expansion of apical surfaces (Figure 6—figure supplement 1A, B)of all KV cells but KV-post cells still lose their volume via ion channel mediated fluid efflux and undergo lateral extension to facilitate asymmetric KV cell shape changes (Figure 6B, C). Thus, ion channels mediate asymmetric cell shape changes via lateral extension of KV-post cells even when overall lumen expansion is inhibited (Figure 6D). This has been clarified in the Discussion section.

3) I am uncertain about how informative the modeling part is (Figure 4). The vertex model simulations are basically done in 2D without measurements for the tensions in anterior and posterior cells and the external forces during the transition between 4ss and 6ss. The limitation is there is no data showing shape changes of a single cell in the anterior or posterior KV during this crucial transition (4ss to 6ss) based on time-lapse movies. However, I cannot assess on the mathematical modeling using solid-like or fluid-like cells that eventually leads to the conclusion that cell shape changes do not depend on lumen expansion.

The purpose of our vertex model simulations was not a literal simulation of the precise time-dependent cell shape changes in vivo. As the referee correctly points out, we would be missing a lot of information for that (cell-cell interfacial tensions, external forces, tracking of cell shapes). Rather, the purpose of our simulations was to identify potential mechanisms that could induce an asymmetric LWR in the KV cells. Since we do not know the values of the cell-cell interfacial tensions and the properties of the surrounding cells, we scanned the parameter space for many possible combinations of both, and we found there was a large regime where asymmetric cell volume change drives asymmetric KV cell shape change. Based on these simulations, we could identify a potential mechanism that translates volume asymmetry into cell shape asymmetry (Figure 4G). Then we wanted to check whether this mechanism depended on the lumen expansion, and found that even without an expanding lumen, asymmetric cell volume change can drive asymmetric cell shape change (Figure 4C, D, F, H).

4) There is inconsistency of the data upon ouabain treatment at 2ss (Figure 3B vs. Figure 3—figure supplement 2). The former shows a reduction of the lumen size, whereas the latter shows that the lumen size looks unaffected.

Thank you for pointing this out. We have now quantified the effect of ouabain on KV lumen expansion and present these results in Figure 3—figure supplement 2B. The results were consistent with previous experiments, as reported in Figure 3B. We have therefore included a more representative image in Figure 3—figure supplement 2C.

5) What is the biological significance of cell height changes at 2 ss in embryos treated with ion channel inhibitors or lgl2-morphant embryos? It is confusing, as cell height does not significantly change between anterior and posterior cells from 2ss to 8ss.

Height changes reflect reduced lumen expansion as compared to controls (see also the response to point #2 above). This difference is already significantly different at 2ss in ouabain, CFTR inh, CFTR MO treated embryos and becomes significant by 8ss in Lgl2 MO treated embryos. In wild-type and all treated embryos, height responds similarly in both anterior and posterior cells to lumen expansion (or lack thereof).

Reviewer #3:

[…] 1) Inhibiting ion channels inhibits cell shape changes/asymmetric cell changes in KV are separable from lumen expansion.

The statistical analysis. Ouabain treatment and cftr knock down lead to similar conclusion which is great. However, both experiments are based on the fact that the results are not significant in the treated embryos at 2SS and 8SS. For this type of analysis, the authors should provide a power analysis to demonstrate that the sample size is enough to conclude about not significant results. Similar comment for jupa and igl2 treatments.

We have repeated the treatments with ouabain, cftr MO and lgl2 MO to increase the sample size of both KV-ant and KV-post cells. We have updated Figure 3B, D and Figure 6C accordingly. Moreover, we dedicate a new Materials and methods section and two new tables (Figure 3—source data 1 and Figure 6—source data 1) to a statistical power analysis to substantiate our results of non-significance in Figures 3A-D (ion channel inhibition) and 6A-C (jupa and lgl2). In particular, for the ion channel inhibition, we test the non-significance against the alternative hypothesis that the AP volume and LWR differences are as high as in the unperturbed case (DMSO control) at 8ss. We find false error rates for the volume always below the conventional chosen value of 20%, and for the LWR even below 5%. This corresponds to a statistical power of more than 80% and 95%, respectively. We performed a similar analysis for the jupa and lgl2 treatments and always found a statistical power of at least 90%.

2) Mechanical properties of the surrounding cells. In simulations, asymmetric cell volume changes could account partially for cell shape changes, provided that the mechanical properties of the cells satisfy certain conditions.

The issue about mechanical properties of cells is important. The mechanical properties of surrounding cells are important and need to be solid like at least anteriorly. Please provide reasonable explanation as to why anterior and posterior could be solid like and more generally, what are the arguments implying that mesodermal cells are solid like (this can be part of the Discussion). The 'pinning' hypothesis is interesting in that aspect and could be discussed. Lance Davidson’s work could be discussed as it seems that the notochord is a pretty rigid structure from his work. In addition, could the fact that fibronectin is enriched in the anterior side of the KV could increase the possibility that cells are solid like anteriorly?

Our modeling results indicate that asymmetric cell shape changes in KV are more pronounced if only the anterior external cells have solid-like mechanical properties than if only the posterior external cells are solid-like, as the solid-like cells on the anterior side are able to "pin" the interface between the KV-ant cells and the KV-post cells. In order to determine general mechanisms, we have indeed kept the model very simple – for example, it does not take into account effects of additional tissues/structures (e.g. notochord) or extra-cellular matrix (ECM). It is possible that the notochord – which physically interfaces with KV-ant cells – may be a solid-like structure, and this may provide the pinning mechanism. Additionally, an AP gradient of ECM may help prevent neighbor exchanges and give rise to solid-like behavior only in anterior external cells. It is also possible that the solid-like ECM directly physically pins the KV-ant cells, so that the mechanism we have identified may operate even if the anterior cells beyond the ECM are fluid-like. These possibilities have been expanded upon in the discussion. In addition, we have provided new Video 5 that depicts enrichment of the ECM molecule fibronectin associated with the notochord and anterior external cells, which is consistent with previous findings (Campagnon, et al. 2014. Dev Cell. PMID: 25535919). Our future work will investigate the contributions of notochord and ECM to the mechanical properties of external cells and asymmetric cell shape changes in KV.

3) Cell volume changes and actomyosin. Given that their simulations show that cell volume changes alone could not account for cell shape changes, it is weird that in Figure 3 cell shape changes are completely gone. A way to interpret these data is that ouabain does not specifically target cell volume but also affects elements modulating cell shape, such actomyosin. This raises the question of the effect of ouabain on the actomyosin network. The demonstration is that the actomyosin network is not altered by the treatment. The authors should check it with phosphorylated myosin via antibody staining (as they did in Wang 2012) to see if inhibiting ion channels somehow affects actomyosin contractility. Or is it a problem with the model?

Thank you for bringing up this important point. We propose that cell volume changes contribute to epithelial morphogenesis in KV and are likely linked to additional important factors (e.g. mechanical properties of cells, tissue-tissue interactions between notochord and KV, and actomyosin cytoskeletal contractility) that are not currently in our model. We are quite intrigued about the relationship between cell volume changes and actomyosin contractility in KV epithelial morphogenesis. As suggested here, we used antibodies that detect phosphorylated myosin light chain (pMLC) to test whether treating embryos with ouabain alters non-muscle myosin II activity in KV cells. Our group (Wang, et al. 2012. Developmental Biology. PMCID: PMC3586254) and the Heisenberg group (Compagnon, et al. 2014. Developmental Cell. PMID: 25535919) have previously used this pMLC antibody to qualitatively assess myosin II activity in KV. Over the course of several trials, we detected cortical pMLC staining in control and ouabain treated KV cells that localized to the cortex of KV cells marked by the tight junction protein ZO1 (see Author response image 2A). However, it was not clear to us whether pMLC levels were different in treated KVs as compared to controls. Despite significant efforts to make this assay quantitative, we have not overcome variabilities in the antibody staining that appear to be due to variable penetration of the antibody into deep tissue layers where KV is located.

As an alternative approach to measure myosin II activity in KV, we took advantage of our stable transgenic Tg(actb2:myl12.1-MKATE2) embryos that express the fluorescent mKate2 protein fused to myosin light chain (Myl12.1). We and others (Compagnon, et al. 2014. Developmental Cell. PMID: 25535919) have found that the accumulation and intensity of Myl12.1-fusion proteins correlates with phosphorylated myosin II antibody staining intensity, and thereby serve as a good proxy for active myosin II. Embryos from homozygous Tg(actb2:myl12.1-MKATE2) parents were treated with ouabain starting at the tailbud stage and then KVs in live treated and control embryos were imaged at the 6 somite stage (see Author response image 2B). Imaging live KVs eliminated the need to fix embryos and perform the antibody staining procedure. Blind measurements of Myl12.1-mKate2 intensity at randomly selected cell-cell interfaces did not detect a difference between control and ouabain treated KVs (see Author response image 2C). Similar results were obtained in multiple experiments, and measurements of KV area (see Author response image 2D) were used to test efficacy of ouabain treatments.

Although these results suggest actomyosin is not dramatically changed in KVs treated with ouabain, we cannot rule out more subtle effects. Thus, we do not feel it is appropriate to draw conclusions from this one experimental approach and include these data in the present manuscript. In future work that is beyond the scope of this paper, we will investigate more rigorously the relationships among ion flux, cell volume changes and cytoskeletal contractility.

Author response image 2
Effect of ouabain treatments on the actomyosin network in KV.

(A) Antibodies that recognize phosphorylated myosin light chain (pMLC) were used to detect active myosin II at the apical cortex of KV cells marked by ZO1 staining. pMLC staining was present in both control and ouabain treated KVs, but this signal proved difficult to quantify. (B) Expression of fluorescent myosin light chain (myl12.1)-mKate2 fusion proteins at the apical cortex of KV cells in live embryos. Dashed yellow lines outline KV lumen. Scale bars=20 mm. (C) Expression intensity of myl12.1-mKate2 measured at cell-cell interfaces was found to be similar between control (n=15 interfaces) and ouabain (n=25 interfaces) treated embryos. (D) Measurements of maximum KV lumen area indicated that ouabain treatments were effective at blocking ion flux and lumen expansion in these embryos. N=number of embryos analyzed. Error bars=one standard deviation. *p=0.001. ns=not significant, (Welch’s T-Test).

https://doi.org/10.7554/eLife.30963.035

4) The conclusion 'Asymmetric cell volume changes regulate epithelial morphogenesis'. It should be toned down providing that we do not know if the surrounding cells are solid like and that we do not know how the actomyosin network is compromised by the treatments used. The term 'regulate' seems too strong at this point.

We have toned down the conclusions and changed the title to “Cell volume changes contribute to epithelial morphogenesis in zebrafish Kupffer’s vesicle.”

https://doi.org/10.7554/eLife.30963.037

Article and author information

Author details

  1. Agnik Dasgupta

    Department of Cell and Developmental Biology, State University of New York, Upstate Medical University, Syracuse, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-0860-1006
  2. Matthias Merkel

    Department of Physics, Syracuse University, Syracuse, United States
    Contribution
    Conceptualization, Software, Investigation, Writing—original draft, Writing—review and editing
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9118-1270
  3. Madeline J Clark

    Department of Cell and Developmental Biology, State University of New York, Upstate Medical University, Syracuse, United States
    Contribution
    Investigation, Methodology, Writing—review and editing
    Competing interests
    No competing interests declared
  4. Andrew E Jacob

    Department of Cell and Developmental Biology, State University of New York, Upstate Medical University, Syracuse, United States
    Contribution
    Validation, Investigation, Methodology
    Competing interests
    No competing interests declared
  5. Jonathan Edward Dawson

    Department of Physics, Syracuse University, Syracuse, United States
    Contribution
    Conceptualization, Software, Investigation
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0001-9770-8475
  6. M Lisa Manning

    Department of Physics, Syracuse University, Syracuse, United States
    Contribution
    Conceptualization, Resources, Supervision, Funding acquisition, Writing—original draft, Writing—review and editing
    For correspondence
    mmanning@syr.edu
    Competing interests
    No competing interests declared
  7. Jeffrey D Amack

    Department of Cell and Developmental Biology, State University of New York, Upstate Medical University, Syracuse, United States
    Contribution
    Conceptualization, Resources, Supervision, Funding acquisition, Writing—original draft, Writing—review and editing
    For correspondence
    amackj@upstate.edu
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5465-9754

Funding

Simons Foundation (#446222)

  • Matthias Merkel
  • M Lisa Manning

Research Corporation for Scientific Advancement (Cottrell Scholar program)

  • Matthias Merkel
  • M Lisa Manning

Gordon and Betty Moore Foundation

  • Matthias Merkel
  • M Lisa Manning

National Institutes of Health (R01GM117598)

  • M Lisa Manning

National Institutes of Health (R01HL095690)

  • Jeffrey D Amack

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank Jeffrey Saffitz, Stephanie Woo and Maura Grealy for sharing reagents. We also thank members of the Amack and Manning groups for helpful discussions and Gonca Erdemci-Tandogan for providing critical feedback on the manuscript. A special thank you to Fiona Foley and Sharleen Buel for outstanding technical support. This work was supported by NIH grants R01HL095690 (JDA) and R01GM117598 (MLM). Additional support was provided by a grant from the Simons Foundation (#446222 MLM and MM), the Research Corporation for Scientific Advancement through the Cottrell Scholars Program (MLM and MM), and through the Gordon and Betty Moore Foundation (MLM and MM). Computing infrastructure support was provided through NSF ACI-1541396.

Reviewing Editor

  1. Marianne Bronner, California Institute of Technology, United States

Publication history

  1. Received: August 5, 2017
  2. Accepted: January 26, 2018
  3. Accepted Manuscript published: January 29, 2018 (version 1)
  4. Version of Record published: February 6, 2018 (version 2)

Copyright

© 2018, Dasgupta et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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