Organ morphogenesis depends on the precise orchestration of cell migration, cell shape changes and cell adhesion. We demonstrate that Notch signaling is an integral part of the Wnt and Fgf signaling feedback loop coordinating cell migration and the self-organization of rosette-shaped sensory organs in the zebrafish lateral line system. We show that Notch signaling acts downstream of Fgf signaling to not only inhibit hair cell differentiation but also to induce and maintain stable epithelial rosettes. Ectopic Notch expression causes a significant increase in organ size independently of proliferation and the Hippo pathway. Transplantation and RNASeq analyses revealed that Notch signaling induces apical junctional complex genes that regulate cell adhesion and apical constriction. Our analysis also demonstrates that in the absence of patterning cues normally provided by a Wnt/Fgf signaling system, rosettes still self-organize in the presence of Notch signaling.https://doi.org/10.7554/eLife.21049.001
Organ morphogenesis relies on the integration of complex processes, such as cell migration, cell shape changes and cell specification to generate the correct three-dimensional geometry necessary for function. Additionally, these cell behaviors must be coupled with mechanisms that regulate the final size of the organ for correct integration into the organism. Understanding the mechanisms that underlie these phenomena is a major focus of modern biology. Even though the intracellular mechanisms leading to cell shape changes are fairly well understood, we are only beginning to elucidate how signaling pathways coordinate cell shape changes with the development of a whole tissue or embryo. Additionally, the importance of multicellular, epithelial rosettes in Drosophila axis elongation and retina development, mouse pre-implantation embryo morphogenesis, pancreas development, brain tumors or the neural stem cell niche has only fairly recently been recognized (Bedzhov and Zernicka-Goetz, 2014; Blankenship et al., 2006; Harding et al., 2014; Martin and Goldstein, 2014; Wippold and Perry, 2006).
The zebrafish lateral line is a powerful model to study sensory organ morphogenesis, as it develops superficially in the skin and is amenable to experimental manipulation and in vivo imaging. The lateral line is a sensory system for the detection of water movements and consists of rosette-shaped sensory organs (neuromasts) that are arranged in lines along the body of the animal. Each neuromast is composed of sensory hair cells surrounded by support cells. Lateral line hair cells are homologous to vertebrate inner ear hair cells and are specified by the same molecules (Nicolson, 2005). The lateral line system on the trunk develops from an ectodermal placode posterior to the ear that migrates to the tail tip. The migrating placode (now called primordium) periodically deposits clusters of cells that mature into neuromasts, thus forming a line of sensory organs. This migrating primordium consists of a mesenchymal leading region and a trailing region in which cells apically-basally polarize and then apically constrict to form garlic bulb/rosette-shaped proneuromasts (reviewed in [Harding et al., 2014]). The two domains are maintained by a feedback mechanism between the Wnt and Fgf pathways (Aman and Piotrowski, 2008). Activation of the Wnt pathway in the leading region induces Fgf ligands that activate the Fgf pathway in the trailing region. Fgf ligand expression is uniform in the leading region but then becomes restricted to one central cell as organs (proneuromasts) begin to form. Fgf ligand expression by a central proneuromast cell that activates Fgf signaling in surrounding cells is crucial for proneuromast formation and maintenance (Durdu et al., 2014; Ernst et al., 2012; Harding and Nechiporuk, 2012; Lecaudey et al., 2008; Nechiporuk and Raible, 2008). Fgfr-Ras-Mapk signaling is thought to directly induce apical constriction and rosette formation via the activation of the actin-binding protein shroom3. shroom3 leads to apical localization of Rock2a kinase that phosphorylates non-muscle myosin II (pNMII) driving actomyosin constriction (Ernst et al., 2012; Harding and Nechiporuk, 2012).
Of all the mutants/manipulations thus far analyzed that affect lateral line development only very few lead to an increase in organ size. The only manipulation described that causes an increase in neuromast size is the upregulation of Wnt signaling, while the size of the primordium increases after inhibition of the Hippo pathway member amotl2a (Agarwala et al., 2015; Head et al., 2013; Wada et al., 2013; Wada and Kawakami, 2015; Jacques et al., 2014). The Hippo pathway controls organ size via a kinase cascade that leads to the phosphorylation and degradation of the transcriptional co-activators Yap/Taz (Sun and Irvine, 2016). In the absence of pathway activation, Yap/Taz are translocated to the nucleus where they activate proliferation and survival genes. The Wnt pathway affects organ size via controlling proliferation, which, at least partially, is regulated by amotl2a (Agarwala et al., 2015). In contrast, loss of the transcriptional co-activator yap1 rescues the amotl2a overproliferation phenotype but does not interact with the Wnt pathway. Thus, the details of how Wnt, amotl2a and yap1 are integrated to affect proliferation are not well understood. Here we describe how the upregulation of Notch signaling by overexpression of the Notch1a intracellular domain (ICD; NICD, [Scheer and Campos-Ortega, 1999]) in the primordium, leads to a significant increase in neuromast size. The increased neuromast size in both Notch and Wnt overexpressing embryos is yap1-independent, and in case of NICD the organ size increase is even proliferation-independent.
Apical constriction is driven by cortical tension-generating actomyosin contractions that are transmitted to neighboring cells via apically localized cadherins (reviewed in [Heisenberg and Bellaïche, 2013; Martin et al., 2009; Pilot and Lecuit, 2005; Schwayer et al., 2016]). We demonstrate that Fgf-dependent Notch signaling cell-autonomously, and downstream of Fgf signaling upregulates adherens and tight junction molecules. These molecules together form an apical junctional complex associated with a circumferential actomyosin belt (Niessen, 2007). Thus, Notch signaling increases organ size independently of proliferation by coordinating actomyosin activation and cell adhesion.
When we constitutively activate Notch signaling in the lateral line by driving the NICD in (Tg(cldnB:lynGFP);Tg(cldnB:gal4) x Tg(UAS:nicd)) embryos, we observe that deposited neuromasts are substantially larger than in sibling embryos (Figure 1A–B’, Figure 1—figure supplement 1A–A'; Video 1). The NICD transgene is strongly induced when the lateral line placode is forming at 13 hr post fertilization (hpf) (data not shown). It is also myc-tagged and and shows widespread expression in the lateral line and skin (Figure 1C–D). The neuromast size increase is reminiscent of Wnt overexpressing apc mutant neuromasts (Figure 1F–F’, [Wada et al., 2013]). In contrast, a reduction of Notch signaling in mib1 mutants causes fragmentation of neuromasts (Figure 1E,E’; Video 2; [Matsuda and Chitnis, 2010]).
Labeling of the first deposited trunk neuromast (L1) with the actin marker phalloidin shows that the NICD apical F-actin meshwork is larger compared to wildtype neuromasts, suggesting that NICD neuromasts are composed of more apically constricted cells (Figure 1G–H’). The robust induction of NICD in (Tg(cldnB:lynGFP);Tg(cldnB:gal4) x Tg(UAS:nicd)) transgenic embryos, is also evidenced by upregulation of the Notch targets notch3, her4, strong activation of the Notch reporter Tp1bglob:EGFP and downregulation of delta ligands (Figure 1I–K’ and Figure 1L–N’). On the other hand, in mib1 mutants notch3 and her4 are reduced but not completely absent and delta genes are upregulated (Figure 1I’’–J’’ and L’’–N’’; Matsuda and Chitnis, 2010). NICD primordia migrate more slowly and deposit fewer neuromasts than wildtype primordia and 80% of the NICD primordia do not reach the tail tip, as they deposit too many cells and run out of cells (Figure 1O and Figure 1—figure supplement 1A–C).
During early posterior lateral line development, Notch determines the proportion of placodal cells that contribute to the lateral line ganglion posterior to the ear and how many cells will become part of the migrating primordium (Mizoguchi et al., 2011). Consequently, ectopic activation of Notch signaling generates an initially larger primordium at the expense of the ganglion, which could explain the increase in neuromast size (Figure 1O and Figure 1—figure supplement 1D). Indeed primordium size limits how large NICD neuromasts can grow. NICD primordia become increasingly smaller during migration and NICD neuromasts accordingly decrease in size the further posterior they are located along the trunk of the embryo (Figure 1O and P). However, by 32 hpf NICD and sibling primordia consist of the same number of cells but NICD primordia still deposit larger L3 neuromasts (Figure 1O and P; Video 3). Also, heat-shock activation of Notch signaling after the primordium and ganglion have separated significantly increases the neuromast cell number (Figure 1Q–R’, Figure 1—figure supplement 1E). These results indicate that, although the primordium size determines the maximum size of an enlarged neuromast, the increased NICD neuromast size is independent of the early role of Notch in allocating cells to the primordium.
The size of the migrating primordium is partially controlled by the Hippo pathway components amotl2a (a tight junction-associated scaffolding protein) and yap1 (a transcriptional co-activator) (Agarwala et al., 2015). amotl2a inhibits proliferation, whereas yap1 is required in the primordium to reach its normal size. To test if the Hippo pathway might modulate neuromast size downstream of Notch or Wnt signaling we tested if (a) Manipulating Wnt or Notch signaling affects the expression of Hippo pathway genes and (b) if the downregulation of yap1 rescues neuromast size in NICD and apc embryos. yap1 is expressed in the leading 2/3 of the wildtype primordium (Figure 2A’) and is upregulated in NICD (Figure 2B’). yap1 is also increased in apc primordia correlating with the increased proliferation observed in these embryos (Figure 2D’; [Aman et al., 2011]). In contrast, yap1 is downregulated in mib1 and Wnt depleted hs:dkk1b primordia (Figure 2C’,E’). However, yap1 is not expressed in deposited wildtype, NICD, mib1, apc or hs:dkk1b neuromasts suggesting that proliferation in deposited neuromasts is yap1-independent (Figure 2A–E). We also analyzed the canonical Hippo pathway inhibitor stk3 (the ortholog of hippo kinase in Drosophila) and amotl2a, which inhibits Wnt-induced proliferation (Figure 2—figure supplement 1A–J'; Agarwala et al., 2015). The expression of these genes does not correlate with proliferation in the primordia with manipulated Wnt or Notch signaling, showing that we do not yet fully understand the integration of Hippo signaling with other mitogens and that further studies are needed.
However, the upregulation of yap1 in primordia correlates with the increased neuromast cell numbers in apc and NICD. In addition, yap1 morpholino injections inhibit the ectopic proliferation in the primordium that is induced by loss of amotl2a, phenocopying the yap1 mutant (Agarwala et al., 2015). We therefore injected yap1 morpholino into NICD and apc embryos and counted the number of cells in the deposited neuromasts, as well as migrating primordium. yap1 morpholino reduces the number of primordium cells in its expression domain in the leading 2/3 of wildtype, apc and NICD primordia (Figure 2F). As morpholino injections can cause cell death in the lateral line (Aman et al., 2011), we injected the yap1 morpholino into homozygous p53 mutant embryos (Figure 2—figure supplement 1K). The primordium cell number decreases from approximately 100 to 80, which is similar to what we find in yap1 morpholino-injected wildtype embryos (Figure 2H), suggesting that the decrease in cell number is not caused by morpholino-induced p53 activation (Kok et al., 2015). Although yap1 downregulation significantly reduces the size of the primordium, it does not rescue neuromast size in apc or NICD embryos to a wildtype level (Figure 2G). The reduction in NICD L1 and L2 pro-neuromast size in yap1 morphants (Figure 2F and G) can be attributed to the role of yap1 in primordium size regulation and that the primordium size limits the number of cells available for allocation into forming neuromast (Figure 1O and P). The above experiments show that the increase in neuromast size in apc and NICD is yap1 independent.
Activation of Wnt signaling produces larger neuromasts via hyperproliferation and we therefore tested if proliferation is upregulated in a yap1-independent fashion in NICD primordia and neuromasts (Wada et al., 2013). In contrast to apc neuromasts, which show a significant increase in proliferation, NICD neuromasts show a significant decrease (Figure 3A–D). Accordingly, apc neuromasts gain a significant number of cells after deposition between 32–56 hpf, whereas NICD neuromasts do not significantly grow in cell number (Figure 3E). The BrdU index and number of primordium cells in 31 hpf apc primordia is not significantly higher than in the siblings (Figure 3F,F’,H,I), although 33 hpf apc primordia possess a significant increase in proliferating cells (Aman et al., 2011). Similarly, the BrdU index and the number of cells in 35 hpf NICD primordia is unchanged (Figure 3G–I).
The striking increase of the BrdU index in deposited apc neuromasts compared to the primordium suggests that apc neuromasts only grow larger once deposited, whereas the increase in the size of NICD neuromasts is controlled by a proliferation-independent process within the primordium, prior to deposition. To further test the hypothesis that aberrant proliferation does not contribute to increased neuromast size in NICD embryos, we inhibited proliferation by treating embryos with the DNA replication inhibitors hydroxyurea and aphidicolin (HUA and Aph, Figure 3J–L). Indeed, the size of the last deposited NICD neuromast is not significantly reduced, even though the primordium cell number is smaller (Figure 3K–L). On the other hand, apc neuromasts were rescued to a wildtype size and apc primordia became smaller (Figure 3K–L). To further test if Wnt and Notch affect neuromast size via independent mechanisms we simultaneously activated Wnt and Notch signaling. The induction of Wnt signaling in NICD embryos causes even bigger neuromasts than in untreated NICD embryos, suggesting an additive effect of the two pathways (Figure 3M–O). These results demonstrate that Wnt signaling affects neuromast size via upregulating proliferation in the deposited neuromasts, whereas activation of Notch causes an increase in neuromast size independently of the proliferation rate.
As proliferation does not contribute to the large proneuromast size in NICD primordia and Notch is crucial for cell fate specification via lateral inhibition, we asked if in NICD primordia a switch in cell fate could be responsible for the increase in organ size. As yet, four cell types have been identified in the primordium: (a) mesenchymal, unpatterned cells in the leading region, (b) hair cell precursors, (c) support cells in the center and (d) future interneuromast cells that are deposited in between neuromasts (Figure 1A’, arrows). Interneuromast cells are lateral line stem cells that postembryonically give rise to additional sensory organs (Grant et al., 2005; López-Schier and Hudspeth, 2006; Lush and Piotrowski, 2014). To test if Notch changes the fate of one cell population into another we counted the number of cells in these four populations in wildtype and NICD primordia.
In the ear and lateral line, Delta/Notch signaling is essential for the specification of sensory hair and support cells (Haddon et al., 1999; Itoh and Chitnis, 2001; Riley et al., 1999; Millimaki et al., 2007). The deltad- (and atoh1a) – expressing cell differentiates into a hair cell, whereas the surrounding Notch-expressing cells are specified as support cells (Matsuda and Chitnis, 2010). As Notch signaling inhibits atoh1a, NICD neuromasts consist only of support cells (Figure 4A–L). However, within the primordium only one atoh1a-labeled hair cell precursor exists per proneuromast (Figure 4K) and their cell fate switch to a support cell cannot account for an increase of an average of 15 or 11 cells in the first proneuromast (orange, Figure 4M–N’) and an increase of 12 or 19 cells in the last proneuromast cells at 26 hpf and 30 hpf (green, Figure 4M–N’). Nevertheless, we tested if the ectopic generation of hair cells in NICD neuromasts would rescue the neuromast size. We induced hair cells in NICD primordia and neuromasts by activating the hair cell specification gene atoh1a using a heat-shock inducible transgenic line (Figure 4—figure supplement 1A–E’’). Hair cell containing neuromasts in heat-shocked Tg(hs:atoh1a;NICD) larvae are not significantly different in size from sibling NICD neuromasts (Figure 4—figure supplement 1F). Thus, the loss of hair cells in NICD neuromasts does not cause the increase in NICD neuromast size. Interestingly, hs:atoh1a alone induces a significant increase in neuromast size (Figure 4—figure supplement 1C'’ and F). The increase of her4 expression in response to ectopic atoh1a activation suggests that Atoh1a possibly induces larger neuromasts through Notch signaling activation in the primordium (Figure 4—figure supplement 1B–E).
We then asked if a cell fate switch in mesenchymal leading cells contributes to the large neuromast phenotype in NICD. We determined that the number of mesenchymal cells in the leading region is initially normal in 26 hpf embryos and is only reduced from an average of 9 to 5 cells in 30 hpf NICD primordia (Figure 4M and N’). Therefore, the reduction in the number of leading region cells is equally insufficient to explain the increase in organ size. The last potential cell fate switch that could contribute to the organ size increase is the loss of future interneuromast cells from the primordium. Future oblong-shaped interneuromast cells are located at the periphery of the primordium (Figure 4M’, arrow) but their number is not affected in NICD embryos (Figure 4O–P arrows, Figure 4Q). The only significant difference we detected between NICD and sibling primordia is that NICD primordia possess on average fewer, larger proneuromasts at the expense of a normal-sized proneuromast (Figure 4M and M’, grey cells, and Figure 4R). We therefore conclude that cell fate changes are not major contributors to the increase in organ size in NICD primordia but that the allocation of support cells into fewer but larger proneuromasts is the major cause.
The loss of Fgf signaling leads to the loss of proneuromasts/rosettes (Lecaudey et al., 2008; Nechiporuk and Raible, 2008). It has therefore been proposed that the Fgf-expressing central cell in a proneuromast acts as a signaling center that recruits surrounding support cells by inducing Fgf signaling in these cells (Lecaudey et al., 2008; Nechiporuk and Raible, 2008). We therefore tested if in NICD proneuromasts, Fgf ligand expression or Fgf signaling is increased. Surprisingly, even though fgfr1 is normally expressed, fgf3, fgf10a and the Fgf targets pea3 and dkk1b are substantially abrogated in NICD primordia (Figure 5A,A’,B,B’,C,C’,D,D’, Figure 8L,L’). Fgf signaling is likely reduced because: (a) Notch inhibits atoh1a, which is required for fgf10a expression in the most mature rosettes in the primordium (Figures 4J,L and 5C,C’ arrows; [Matsuda and Chitnis, 2010]) and (b) because Notch inhibits Wnt, which normally induces Fgf signaling in the first forming rosette (see below, Figure 8I’–K’). Therefore, increased Notch signaling causes decreased Wnt and Atoh1a dependent Fgf signaling. The loss of Notch signaling in mib1 mutants also leads to loss of Fgf signaling in central cells, however fgfr1a and pea3 are strongly expressed in peripheral cells of the primordium (Figure 5A’’,D’’). In mib1 primordia Fgf signaling is reduced in central cells because of the upregulation of atoh1a, which is normally restricted to a central cell by Notch signaling. Atoh1a, inhibits fgfr1a, thus causing the loss of Fgf signal transduction (Figure 5A’’–D’’; [Matsuda and Chitnis, 2010]).
As the expression of Fgf pathway genes is not completely abrogated in NICD primordia, we depleted Fgf signaling in NICD embryos with (a) the pharmacological Fgfr1 inhibitors SU5402 and PD173074 (Figure 5E–F’’), (b) treated NICD embryos with an inhibitor of MAPKK/Di-pErk (PD0325901) (Figure 5E’’’–F’’’, Figure 5—figure supplement 1A-B’) and (c) heat-shock induction of dominant-negative fgfr1 in NICD embryos (Figure 5G–H’). Irrespective of by which method we deplete Fgf signaling in NICD embryos, proneuromasts form, even though the primordium still eventually stalls (Figure 5F’,F’’,F’’’,H’). Also, the molecular analysis of Fgf-depleted wildtype and NICD embryos shows that, although the Fgf signal transduction protein di-pErk is absent from all embryos (Figure 5J’’, L’’), only in NICD embryos Rock2a and pMRLC are still properly localized to the apical constrictions of proneuromast cells (Figure 5L',P'). These results demonstrate that in the absence of Fgf signaling, Notch signaling is sufficient to apically localize Rock2a and pMRLC and induce/maintain apical constrictions (Figure 5I-P').
To test if Notch signaling acts downstream of Fgf signaling we performed in situ analyses with the Notch target her4 in Fgf-depleted sibling and NICD embryos (Figure 5Q-T'). Indeed, Notch signaling is highly reduced in the siblings in the absence of Fgf signaling (Figure 5Q–S’; [Nechiporuk and Raible, 2008]) but the loss of Fgf signaling has no effect on her4 expression in NICD primordia (Figure 5R–T’). We determined that Notch acts downstream of the RAS/MAPK pathway, as the MAPKK inhibitor PD0325901 leads not only to the loss of the Fgf targets pea3 and fgfr1a but also of notch3 and her4 (Figure 5—figure supplement 1C–F’).
To functionally test if Fgf signaling affects organ morphogenesis via the induction of Notch signaling, we crossed Notch-depleted mib1 mutants with a heat-shock line that allows us to constitutively upregulate Fgf signaling (Tg(hs:cafgfr1), Figure 5X–Y’’’). The analysis of the resulting embryos demonstrates that in the absence of Notch signaling, the activation of Fgf signaling is not sufficient to rescue proneuromast formation (Figure 5Y–Y’’). Also, the expression of ZO1, a component of the apically localized tight junctions is also still reduced in mib1;hs:cafgfr1 embryos (Figure 5Y',Y''') confirming that Notch acts downstream of Fgf signaling in rosette formation.
shroom3 is an Fgf-dependent scaffolding protein thought to be required for preneuromast cell apical constriction (Figure 5U, [Ernst et al., 2012]). shroom3 is lost in Fgf-depleted primordia, reduced in NICD primordia, completely lost in Fgf-depleted NICD primordia and upregulated in mib1 primordia (Figure 5U’, V, V’, W). As NICD primordia are forming proneuromasts in the absence of shroom3 (Figure 5V’, arrows), shroom3 is not required for rosette formation in the presence of Notch. We also expressed dominant negative shroom3 in wildtype primordia and did not observe any rosette or lateral line defects (Figure 5—figure supplement 1G–I). Likewise, in contrast to published data (Durdu et al., 2014; Ernst et al., 2012), shroom3 morpholino injections did not delay rosette formation. To investigate if other shroom genes might act redundantly with shroom3 we performed in situ hybridization with shroom1, shroom2a, shroom2b and shroom4 (Figure 5—figure supplement 1J–K''). shroom2b and shroom4 are not expressed in the lateral line. shroom1 and 2a are not expressed in NICD primordia and we therefore conclude that shroom3 does not act redundantly with other shrooms in NICD rosette formation.
Our results show that in wildtype primordia Fgfr activation induces Ras-MAPK signaling, which in turn activates Notch signaling. Notch leads to the apical localization of Rock2a and pMRLC in a shroom3-independent fashion, leading to apical constriction of the cell.
As Notch is sufficient to induce apical constrictions in the absence of Fgf signaling, we wondered if apical constriction is affected in mib1 mutant primordia compared to wildtype and NICD primordia (Figure 6). Like in wildtype proneuromasts, cells in NICD proneuromasts constrict apically (Figure 6A,B,E). In 36 hpf mib1 mutant primordia cells in younger proneuromast constrict normally (R2), however cells in the most trailing, about to be deposited rosette (R3) lose apical constrictions and no longer constrict apically (Figure 6C–E; Video 4). We measured cell constrictions in the R3 proneuromast by calculating the ratios between the narrowest cell surface area value by the widest area value. Results confirmed that mib1 R3 cells are much less constricted than wildtype or NICD R3 cells (Figure 6E). We also visualized and quantified apical constriction volumes by coloring the constriction volume in magenta using Imaris (Bitplane) (Figure 6F). Subtracting the constriction volumes of R2 from R3 of WT and mib1 proneuromasts reveals that the difference is positive for wildtype proneuromasts, whereas it is negative for mib proneuromasts. Therefore, the constriction volumes decrease in mib1 as they mature and before they are deposited. These observations support the findings by Matsuda et al., that mib1 proneuromasts fall apart (Matsuda and Chitnis, 2010). The disintegration of apical constrictions becomes more apparent between 32–36 hpf, as the loss of Notch signaling in mib1 mutants becomes progressively more severe, likely because of depleted maternal stores (Matsuda and Chitnis, 2010).
To test if NICD expressing cells might act as signaling centers that recruit surrounding wildtype cells into a proneuromast, we tested if the phenotype is non cell-autonomous. We transplanted cells (magenta color) from NICD transgenic embryos or wildtype embryos (as a control) into wildtype Tg(cldnb:lynGFP) embryos and compared neuromast size (green color) and cell composition between these two conditions (Figure 7A–B’).
We observed that neuromasts only get bigger (green dots) than the average wildtype neuromast (grey line) if they contain NICD clones but not if they contain wildtype clones (Figure 7B). If NICD cells acted as a signaling center enlarged neuromasts should not only contain NICD-positive cells but also possess an above average increase in wildtype cells (black dots). The quantification of all cell transplantation experiments confirmed that the significant increase in neuromast size in NICD>WT experiments (Figure 7B’, green bars) is only due to the excessive incorporation of transplanted NICD cells into proneuromasts, as the number of wildtype host cells (black bars) is not significantly changed between two types of cell transplantation strategies. Therefore, we conclude that wildtype cells are not recruited to contribute to larger neuromasts (Figure 7A–B’; Video 5). Thus, the increase in organ size after Notch activation is cell-autonomous and Notch transducing cells do not act as a signaling center.
It was suggested that self-organization into rosettes depends on cell adhesion molecules, such as e-cadherin (cdh1) and n-cadherin (cdh2), and a reduction in e-cadherin expression correlates with lateral line adhesion defects in mib1 mutants (Matsuda and Chitnis, 2010). The loss of e-cadherin expression was attributed to the inhibitory effect of the expanded atoh1a domain in mib1 primordia. Correspondingly, we found that in NICD primordia e-cadherin mRNA and E-cadherin protein are highly upregulated, accompanied by an increase in the length of adherens junctions in deposited neuromasts (Figure 7C–C’,D–E’’,F–H).
To determine if e-cadherin is regulated by Atoh1a, Wnt, Fgf or Notch signaling we analyzed e-cadherin expression after different signaling pathway manipulations (Figure 7C–CC’’). Our data does not support the hypothesis that atoh1a negatively regulates e-cadherin and that therefore the upregulation of e-cadherin could be caused by the loss of atoh1a in NICD primordia. Ectopic expression of atoh1a in wildtype primordia causes increased e-cadherin levels and neuromast size (Figure 7CC’’ and Figure 4—figure supplement 1F). Simultaneously, her4 is also induced (Figure 4—figure supplement 1C and F). These data suggest that heat-shock induction of atoh1a leads to the upregulation Notch signaling which induces e-cadherin.
Wnt overexpressing apc mutant primordia also show an increase in e-cadherin mRNA and protein expression suggesting that e-cadherin is regulated by Wnt signaling as in many other cell types (Figure 7CC and Figure 7—figure supplement 1A,B; Heuberger and Birchmeier, 2010). Our data show that Wnt signaling regulates e-cadherin via the induction of Notch signaling, as e-cadherin is reduced in mib1 primordia and the upregulation of Wnt with the GSK3β inhibitor BIO is not sufficient to induce e-cadherin in Notch-deficient primordia (Figure 7C’’ and 7CC’). Likewise, Fgf signaling indirectly upregulates e-cadherin expression via the induction of Notch signaling. Loss of Fgf signaling leads to the loss of e-cadherin expression but if Notch is simultaneously activated, e-cadherin is expressed (Figure 7—figure supplement 1C–F'). n-cadherin, on the other hand, is regulated by Wnt in a Notch-independent fashion, as it is not affected in mib1 or NICD primordia (Matsuda and Chitnis, 2010) but it is upregulated in apc mutants (Figure 7—figure supplement 1G–J). N-cadherin also localizes to apical constrictions (Revenu et al., 2014), however, we did not observe a rosette defect in n-cadherin mutants (pactm101b, cdh2hi3644, data not shown). Together these experiments show that e-cadherin is induced by Notch signaling downstream of Wnt, Fgf and Atoh1a.
To test the function of e-cadherin in rosette formation we transplanted cells from e-cadherin morpholino-injected embryos into wildtype embryos, as e-cadherin mutants die during epiboly (Figure 7—figure supplement 1K–M’’; [Kane et al., 2005]). E-cadherin-deficient cells behave normally in wildtype primordia and neuromasts, even if almost the entire neuromast consists of mutant cells, suggesting that other adhesion molecules act redundantly (Figure 7—figure supplement 1M–M’’; Video 6).
To determine if cell adhesion is affected at all in the NICD lateral line, we analyzed if transplanted NICD cells sort out from wildtype cells. First we analyzed if transplanted NICD cells preferentially contribute to neuromast or interneuromast cells and observed that NICD and wildtype cells contributed equally to either lateral line cell type, again demonstrating that cell fate is not affected (Figure 7J). However, NICD cells that contribute to interneuromast cells cluster significantly more often than wildtype cells, which are usually aligned in a string (Figure 7I–I’’’, arrow and Figure 7K; Figure 7—figure supplement 1N,O). Almost 76% of NICD cells formed clusters of 2 or more cells, compared to 13% of wildtype cells. Combined with the finding that transplanted NICD cells contribute to wildtype neuromasts in larger clusters we conclude that Notch overexpression changes cell adhesion properties.
To identify genes that are transcriptionally controlled by Notch signaling during proneuromast formation, we isolated primordium cells from dissected tails of 36 hpf Tg(cldnB:lynGFP);Tg(cldnB:gal4) x Tg(UAS:nicd) and sibling Tg(cldnB:lynGFP);Tg(cldnB:gal4) embryos by FACS and performed RNASeq analysis. We identified 187 genes that are upregulated in the lateral line of NICD embryos (Figure 7—source data 1). GO term analysis revealed that enriched cellular components are: ‘apical junction complex’, containing the tight junction components cldnb, cldna, cldne and cingulinb and ‘actin cytoskeleton’ (Figure 7O–P’ and R–S’). Upregulated actin cytoskeleton genes are formin1 (fmn1), myo5c, baiap2b and cingulinb (cgnb) (Figure 7—figure supplement 2, Figure 7—source data 1 and 2). cgnb is a tight-junction associated protein that links tight junctions to the actomyosin cytoskeleton and regulates RhoA signaling (Aijaz et al., 2005; Terry et al., 2011; Van Itallie and Anderson, 2014). Other molecules that interact with this complex are also upregulated, such as rab25a, a Rab11 GTPase family member, as is celsr2, an atypical cadherin (Figure 7X–Y’). Celsr1, if reduced, causes otic defects because of loss of apical constrictions due to disturbed actomyosin recruitment to the apical junctional complex (Sai et al., 2014). RNASeq analysis also revealed the induction of cd9b by Notch, a Tetraspanin family member implicated in cell-matrix adhesion and Sdf1 (Cxcl12a) mediated migration (Figure 7L–M’, [Arnaud et al., 2015; Leung et al., 2011]).
By performing in situ hybridization experiments with a number of candidate adhesion molecules, we identified that epithelial cell adhesion molecule (epcam), which regulates claudin expression and tight junctions is also strongly upregulated in NICD embryos (Wu et al., 2013; Figure 7U–V’). Conversely, in situ expression analyses demonstrated that molecules induced by NICD are downregulated in mib1 mutants supporting the finding that Notch is regulating the expression of tight junction and adhesion molecules (Figure 7N–Z’).
To determine if tight junction associated genes are responsible for the large organ phenotype we injected a cldnb morpholino into wildtype embryos, and also tested celsr2rw71 mutants for lateral line phenotypes (Kwong and Perry, 2013; Wada et al., 2006). But like with e-cadherin mutant cells, we did not observe apical junction disassembly phenotypes, likely due to the presence of other claudins and celsr1a/1b in the primordium (data not shown). Likewise, a mutation in the adhesion molecule epcam does not cause a lateral line phenotype (Slanchev et al., 2009). cd9b has been previously knocked down by morpholino injections. Primordium migration is normal but neuromast formation is affected (Gallardo et al., 2010). As morpholino injections can be toxic (Aman et al., 2011), the function of cd9b is currently unclear and has to be reinvestigated.
Our data show that (a) cell adhesion is upregulated in NICD cells, as transplanted cells cluster and (b) cell adhesion molecules act redundantly as their individual downregulation does not cause a phenotype. Given that adhesion and tight junction genes belong to the most highly enriched GO term in NICD primordia, and NICD-positive cells form clusters, we conclude that Notch signaling induces an increase in organ size by activating a combination of adherence and tight junction molecules.
Collective cell migration and organ morphogenesis are coordinated in the primordium via signaling interactions between Wnt and Fgf signaling (Aman and Piotrowski, 2008; Chitnis et al., 2012). It is therefore important to determine how Notch signaling fits into this gene interaction network. Wnt signaling in the leading primordium region activates Fgf signaling in the trailing region. Both pathways repress the other pathway via the activation of inhibitors in their respective expression domain (dkk1b and sef/dusp6).
Notch signaling is activated by Fgf signaling in the primordium (Figure 5S’ and Figure 9A,B; [Matsuda and Chitnis, 2010; Nechiporuk and Raible, 2008]). Fgf signaling activates the transcription of notch3 in hair cell progenitors and support cells and of atoh1a and delta ligands just in central hair cell progenitors (Figure 9B). Notch signaling, in turn, inhibits atoh1 expression (Baker et al., 1996; Baker and Yu, 1997; Matsuda and Chitnis, 2010; Millimaki et al., 2007). However, if Notch signaling also acts downstream of Wnt signaling has not been investigated.
In apc mutants or in embryos treated with the GSK3β inhibitor BIO, the Fgf pathway genes pea3 and atoh1a and the Notch target her4 are upregulated (Figure 8A–A’, B-B', C-C' and Figure 8—figure supplement 1). As Notch is activated by Fgf signaling we asked if Wnt activates Notch via the activation of Fgf signaling. We treated sibling and apc mutants with the Fgfr1 inhibitor PD173074. The loss of Fgf signaling attenuates pea3, atoh1a and her4 in sibling and apc mutants, especially in the youngest proneuromast (Figure 8D–F’), which demonstrates that Wnt signaling is not sufficient to activate either atoh1a or her4 to wildtype levels in the absence of Fgf (Figure 8E,E' and F,F'). Also, rosette formation still occurs in NICD primordia with depleted Wnt signaling by heat-shock activation of dkk1b demonstrating that Notch signaling acts downstream of Wnt signaling (Figure 8G–H’). In conclusion, Wnt activates Fgf signaling, which in turn activates atoh1a, delta ligands and Notch signaling (Figure 9A,B).
To test if Notch signaling negatively feeds back on Wnt signaling as in mature neuromasts (Romero-Carvajal et al., 2015), we assessed the expression of Wnt target genes in NICD and mib1 mutant primordia. In NICD primordia, lef1, wnt10a and sef (an Fgf inhibitor that is regulated by Wnt; (Aman and Piotrowski, 2008)) are reduced, whereas in mib1 the expression domains of these genes expand to the trailing region of the primordium (Figure 8I-K'',P). This finding was surprising as the Fgf target and Wnt inhibitor dkk1b is strongly reduced in NICD primordia and wnt10a should expand. dkk1b can therefore not be responsible for the repression of Wnt signaling in NICD primordia (Figure 8L–L’). A clue was provided by our observation that in apc primordia the Wnt target wnt10a is not expanded uniformly in the trailing region but is mostly upregulated in a group of central cells suggesting that Wnt signaling is inhibited in more peripheral cells (Figure 8N,N', arrows). To test if Notch signaling restricts wnt10a expression in peripheral cells, we treated apc mutant embryos with the γ-secretase and Notch inhibitor DAPT. Indeed, wnt10a is now expressed in the most trailing cells demonstrating that Notch inhibits wnt10a, a feedback interaction not previously described in the primordium (Figure 8O,O'). A Fgf-independent inhibition of Wnt signaling by Notch is also supported by our finding that wnt10a expression is still inhibited in NICD primordia treated with Fgf inhibitor, even though loss of Fgf signaling in wildtype embryos leads to expanded wnt10a expression (Figure 8Q–R'). The inhibition of wnt10a by Notch also explains why wnt10a is restricted to a single central cell in proneuromast of a wildtype primordium (Figure 8M,M'). wnt10a seems to be fluctuating in central cells, as it is not obvious in all primordia imaged (Figure 8K).
In conclusion, as illustrated by the schematic representation of gene expression in forming neuromasts, changes in the wnt10a domain only correlate with the presence or absence of Notch (notch3, Figure 8M’–R’). Thus, the analysis of NICD primordia revealed that Wnt signaling is normally not only inhibited by the Fgf target dkk1b but also independently by Notch signaling.
The fact that Notch signaling negatively feeds back on Wnt signaling also explains why Fgf signaling is reduced in NICD primordia. Fgf signaling in the first forming proneuromast depends on Wnt signaling, whereas the expression of fgf10a in the central cell of formed proneuromasts depends on atoh1a (Aman and Piotrowski, 2008; Matsuda and Chitnis, 2010). Therefore, the loss of Fgf signaling in NICD primordia can be attributed to both the inhibition of Wnt signaling and the inhibition of atoh1a expression by Notch in the trailing region. The conclusion that the loss of Fgf signaling in NICD primordia is mostly caused by Notch-mediated repression of Wnt signaling is confirmed by our finding that Fgf signaling (pea3) can be activated in NICD primordia by activating Wnt signaling with the GSK3β inhibitor BIO (Figure 8S,S'). Combined our pathway analyses revealed that Notch is a component of the Wnt/Fgf signaling interaction network that controls migration with organ morphogenesis.
Even though overexpression of both Wnt and Notch signaling induce larger neuromasts, they produce this effect by different mechanisms. Wnt signaling upregulates proliferation in deposited neuromasts, whereas Notch signaling induces an increase in organ size as they form within the primordium in a proliferation-independent manner. Accordingly, cell cycle inhibition in NICD embryos does not rescue the neuromast size, in line with our previous study that demonstrated that proliferation does not influence wildtype neuromast size (Aman et al., 2011). Likewise, although inhibition of the Hippo pathway component amotl2a causes an increase in proliferation in the primordium (Agarwala et al., 2015) and yap1 morpholino injections reduce the primordium size, neither manipulation affects the size of the forming sensory organs in wildtype, apc or NICD embryos (Figure 2G–H; [Agarwala et al., 2015]). Thus, primordium size and lateral line organ size are regulated by independent mechanisms. However, in amotl2a mutant embryos on average one more neuromast develops (Agarwala et al., 2015). Thus, primordium size does not affect neuromast size in a wildtype embryo but rather affects neuromast number. yap1 is upregulated and amotl2a is reduced by Notch signaling without affecting the primordium cell number suggesting that yap1 also possesses proliferation-independent functions that need to be further explored (Figures 3H–I, 9A).
RNASeq and GO term analyses of FACS isolated primordium cells and the analyses of candidate genes revealed that members of the epithelial junctional complex are upregulated in the lateral line of NICD embryos. Epithelial junctional complexes consist of cadherin-based adherens junctions and tight junctions that encompass a large number of transmembrane proteins connected to the actomyosin belt via cytoplasmic scaffolding proteins (González-Mariscal et al., 2003). Because of the enrichment of cell adhesion molecules, such as E-cadherin and because transplanted NICD cells coalesce (Figure 7K, Figure 7—figure supplement 1O), we hypothesize that an increase in cell adhesion is contributing to the increase in organ size. However, transplanted E-cadherin mutant cells do not show a phenotype likely due to redundancy, as the primordium expresses several other adhesion molecules, such as epcam and claudins. Epcam is a calcium-independent, epithelial adhesion molecule and its reduction in mutant zebrafish and cultured cells changes their adhesiveness and induces ectopic, localization of apical junctional complexes (Slanchev et al., 2009; Wu et al., 2013). epcam (tacstd) is required for zebrafish gastrulation movements, and its loss also does not cause a lateral line phenotype suggesting that it acts redundantly with other adhesion molecules in the primordium. A previously reported morpholino-induced lateral line defect was likely caused by morpholino toxicity (Villablanca et al., 2006).
The upregulated Claudin proteins could act redundantly with E-cadherin and Epcam adhesion molecules. Although, Claudins are tight junction molecules and play an important role in controlling permeability of epithelia, they also affect cell adhesion as demonstrated in Xenopus development and cell culture (Brizuela et al., 2001; Kubota et al., 1999). In addition, their loss leads to epithelial to mesenchymal transition and the induction of epithelial characteristics if overexpressed (Bhat et al., 2015). Interestingly, loss of claudin-6 leads to the loss of apical actin accumulation in the developing Xenopus pronephros demonstrating that Claudins regulate a variety of morphogenetic processes (Sun et al., 2015). Thus, it is possible that Claudins are also involved in apical constriction, as has been demonstrated for the tight junction protein ZO-1 (Bhat et al., 2015; Tornavaca et al., 2015). The finding that the knockdown of individual adhesion and tight junction molecules does not cause a phenotype is reminiscent of the collectively migrating Drosophila border cells, which have only been shown to fall apart when JNK signaling is inhibited, which has many downstream targets (Llense and Martín-Blanco, 2008).
Other molecules previously implicated in proneuromast formation also act redundantly. For example, the combined downregulation of lgl1 and lgl2 leads to loss of the leading-most rosettes in the primordium (Hava et al., 2009). Lgl proteins act upstream of Notch signaling, as in the zebrafish retina loss of lgl1 leads to an increase in the apical domain of neuroepithelia, which induces the upregulation of Notch signaling (Clark et al., 2012). Likewise, in the mouse brain lgl1 regulates Notch via the localization of the Notch inhibitor Numb (Klezovitch et al., 2004). In NICD lateral line primordia lgl2 expression is normal (data not shown) suggesting that lgl2 acts upstream of Notch signaling in the lateral line as well. If lgl1/2 are regulated by Fgf signaling has not been determined but lgl1/2 could be a possible link between Fgf and Notch signaling, which needs further exploration.
Apical constrictions are induced by myosin regulatory light chain activity that is phosphorylated by Rho associated coiled-coil protein kinase (ROCK; [Aguilar-Cuenca et al., 2014; Bresnick, 1999; Sawyer et al., 2010]). In the neural tube of mice and frogs the scaffolding and actin-binding protein shroom3 is important for the apical localization of ROCK and neurulation (Haigo et al., 2003; Hildebrand, 2005; Hildebrand and Soriano, 1999; Nishimura and Takeichi, 2008). In contrast to previously published studies, our experiments did not reveal a function for shroom3 in lateral line rosette formation (Durdu et al., 2014; Ernst et al., 2012). Most importantly, neuromast cells apically constrict normally in the absence of shroom3 in NICD embryos (Figure 5V–V’, arrows). In chicken otocyst invagination also occurs independently of shroom3 providing further evidence that apical constriction is achieved via different mechanisms in different contexts (Martin and Goldstein, 2014; Sai et al., 2014).
Notch signaling specifies support cells by inhibiting the proneural gene atoh1a, which is required for the acquisition of a neurogenic cell fate ([Bermingham et al., 1999], Figure 4J,L and Figure 9C). Accordingly, constitutive activation of Notch by overexpression of NICD leads to the loss of atoh1a-positive hair cell progenitors, whereas the loss of Notch signaling in mib1 primordia causes an increase in atoh1a and delta-positive hair cell progenitors (Figure 1L’’–N’’, [Itoh and Chitnis, 2001]). Downregulation of atoh1a in mib1 mutants modestly rescued e-cadherin expression and improved proneuromast cohesion, suggesting that atoh1a inhibits e-cadherin (Matsuda and Chitnis, 2010). A role for neurogenic genes in epithelial morphology was also described in Drosophila, where the loss of these genes causes delamination of cells from epithelia (Hartenstein et al., 1992). Nevertheless, it is unlikely that the NICD phenotype is caused by the loss of the central hair cell precursor and loss of atoh1a expression because: (a) re-expression of Atoh1a in NICD primordia does not rescue neuromast size (Figure 4—figure supplement 1E’’,F) (b) apc and hs:atoh1a primordia show an increase in e-cadherin, even though atoh1a is robustly expressed, arguing that atoh1a does not inhibit e-cadherin expression (Figures 7CC, 8CC’’; Figure 7—figure supplement 1A,B), (c) the loss of atoh1a/b by morpholino injection into wildtype embryos does not affect e-cadherin expression and does not lead to larger neuromasts (Matsuda and Chitnis, 2010; Nechiporuk and Raible, 2008). On the contrary, our data show that ectopic atoh1a causes a significant increase in neuromast size. Interestingly, Atoh1a also upregulates Notch target her4 expression (Figure 4—figure supplement 1B,C,F). This suggests that neuromast size in hs:atoh1a embryos is possibly regulated by Notch signaling acting downstream of Atoh1a dependent Delta ligand expression. During zebrafish inner ear development Atoh1b is required for the initial activation of her4 (Radosevic et al., 2014). Whether Atoh1a regulates Notch in the lateral line still needs to be further investigated.
The above experiments show that, even though NICD primordia loose hair cells, which could lead to signaling changes in neighboring cells, the loss of hair cells is not the cause of organ size increase of NICD proneuromasts.
Interestingly, during proneuromast formation and maturation signaling between central hair cell precursors and surrounding support cells changes slightly between the first forming proneuromast and the trailing, more mature proneuromasts (Figure 9B and C; [Matsuda and Chitnis, 2010]). In the first forming proneuromast fgf3/10a is expressed in all cells, whereas in trailing, more mature rosettes these ligands are restricted to a central cell. This restriction is controlled by Notch signaling that inhibits the proneural gene atoh1a in support cells (Figure 9C). In contrast, even though Fgf signaling is active in all cells in the first forming proneuromast, the downstream targets atoh1a and deltaa/b are immediately restricted to a central hair cell precursor. Likewise, wnt10a is only expressed in one central cell. The restriction of atoh1a to a central cell contrasts findings in flies where atoh1 is first broadly expressed, marking a prosensory field and only subsequently becomes restricted during cell specification (Jarman et al., 1995; Millimaki et al., 2007). The mechanisms by which Notch signaling immediately inhibits atoh1a, delta and wnt10a expression in support cells and confines them to a central cell are still unknown. Another difference between signaling in the first forming and trailing proneuromasts has previously been described. Fgf signaling is induced in all cells of the first forming proneuromast by Wnt ligands, possibly wnt10a (Figures 8K, 9A,B). In more trailing proneuromasts, Fgf signaling depends on atoh1a, rather than Wnt signaling (Figure 9C, [Matsuda and Chitnis, 2010]). Likewise, atoh1a expression becomes Fgf-independent and self-regulatory (Matsuda and Chitnis, 2010; Millimaki et al., 2007). Even though, Fgf and Notch signaling are particularly high in support cells because of lateral inhibition, some Fgf and Notch signaling occurs even in central cells, as evidenced by her4 expression (Figures 1J and 5S, not shown in Figure 9C), as well as notch3, albeit at lower levels (Figure 1I and Figure 5—figure supplement 1E). Possibly, this low level of Notch signaling ensures that also central cells maintain their apical constrictions.
The expression of Fgf ligands in a central cell in a forming neuromast suggests that this cell acts as a signaling center that organizes neighboring cells into a rosette (Harding and Nechiporuk, 2012; Lecaudey et al., 2008; Nechiporuk and Raible, 2008). Indeed, loss of Fgf signaling causes existing neuromasts to fall apart and a failure of new neuromasts to form (Lecaudey et al., 2008; Lush and Piotrowski, 2014; Nechiporuk and Raible, 2008). Durdu et al., reported that Fgf signaling in trailing proneuromasts depends on Fgf ligands being secreted into a dorsal lumen and that only central cells that are apically in contact with the lumen activate the Fgf target pea3 (Durdu et al., 2014). They concluded that Fgf signaling and/or Fgf-dependent rosette formation is important for cell migration and the periodicity of organ deposition (Durdu et al., 2014; Lecaudey et al., 2008). However, lumina only form in the trailing region of the primordium and are not required for Fgf-mediated formation of the first proneuromast (Figure 9B). Our results demonstrate that Fgf signaling is dispensable for rosette formation in the presence of ubiquitous Notch signaling. This finding begs the question why fgf10a production is restricted to a central cell that limits the number of Fgf-responding and Notch-activating cells. A likely explanation is that restricting Fgf signaling and thereby Notch signaling is important to control organ size.
An interesting question is why in NICD primordia two larger rosettes form, rather than three smaller ones as in their siblings. We hypothesize that the Notch-driven increases in cell adhesion and cortical tension change the morphology of the organ to achieve minimal surface tension (Heisenberg and Bellaïche, 2013). Measurements and manipulations of physical forces are needed to test if physical forces determine organ size control and the number of organs that can self-organize in a defined tissue such as the lateral line primordium.
Together, our experiments demonstrate that Notch signaling is an essential part of the feedback loop between Wnt and Fgf signaling that coordinates cell migration with sensory cell specification and organ morphogenesis. Importantly, we show that Notch signaling acts cell-autonomously and downstream of Fgf signaling in sensory organ rosette formation via the regulation of cell adhesion and tight junction molecules. Therefore, Notch is involved in neuromast rosette self-organization that is analogous to rosettes and that precedes lumen formation in pre-implantation mouse embryos (Bedzhov and Zernicka-Goetz, 2014). Similarly, rosettes self-organize during C. elegans gastrulation but are also characteristic for some brain tumors (Pohl et al., 2012; Wippold and Perry, 2006). We speculate that Notch signaling might be involved in these other developmental events as well. Therefore, our results do not only inform lateral line biology but also contribute to our understanding of morphogenesis of other organs and tissues and how rosettes form in certain brain tumors.
Tg(cldnb:lynGFP)zf106 (Haas and Gilmour, 2006), Tg(Tp1bglob:eGFP)um13 (Parsons et al., 2009), Tg(UAS:myc-Notch1a-intra)kca3 (Scheer and Campos-Ortega, 1999), Tg(hsp70l:Gal4-VP16)VU22 (Shin et al., 2007), apcmcr (Hurlstone et al., 2003), mindbomb, mib1ta52b (Itoh and Chitnis, 2001), Tg(hsp70l:dkk1b-GFP)w32 (Stoick-Cooper et al., 2007), Tg(hsp70l:dnfgfr1-EGFP)pd1 (Lee et al., 2005), Tg(hsp70:ca-fgfr1)pd3 (Marques et al., 2008), Tg(hsp70:atoh1a)x20 (Millimaki et al., 2010), Tg(UASDNshroom3) (created by injecting plasmid: pT2dest(bidirectional UAS)-dsred-shrm3DN(520-874)) received from B. Link (Clark et al., 2012; Kwan et al., 2007), Tg(ubi:Zebrabow) (Pan et al., 2013), p53zdf1, pactm101b, cdh2hi3644 kind gift from Anand Chandrasekhar. To generate the Tg(−8.0cldnB:gal4vp16)psi8, we used the zebrafish Tol2 kit (Kwan et al., 2007). The 8 kb claudinB promoter fragment (Haas and Gilmour, 2006) was cloned into the 5’ entry vector and combined via a Gateway reaction with the Gal4-VP16 middle entry vector of the Tol2 kit. The DNA was injected into one-cell stage zebrafish embryos to generate transgenic fish. To generate the TgBAC(cxcr4b:H2A-GFP)p2 transgenic line, the BAC clone DKEY-169F10 was modified in two ways by recombineering. First, the Tol2 sites and the cryaa:dsRed transgenesis marker were inserted into the BAC backbone (Fuentes et al., 2016). Second, a cassette consisting of H2A-GFP flanked by 791 bp and 1042 bp of homology upstream of cxcr4b exon 2, and downstream of the cxcr4b stop codon, respectively, was inserted to replace the cxcr4b coding sequence in cxcr4b exon 2 (amino acid 6–358, the last amino acid before the stop codon) using seamless galK-mediated recombineering (Warming et al., 2005). This transgene expresses the first five amino acids from cxcr4b exon 1 fused to H2A-GFP from the cxcr4b promoter. The final BAC transgene was characterized by SpeI and EcoRI restriction digestion, sequencing of PCR amplicons of the modified locus, and BAC-end sequencing. The DKEY-169F10 BAC library were obtained from ImaGenes GmbH, Germany. BAC transgenes were purified with the nucleobond BAC 100 kit (Clontech). We co-injected 1 nl of 50–250 ng/ml BAC transgene DNA, and 40 ng/ml Tol2 mRNA into the lifting cell of the zygote of 0- to 20-minute-old embryos. The Tol2 mRNA was transcribed from pCS2FA-transposase (Kwan et al., 2007) using the mMessage Machine SP6 Transcription Kit (Thermo Fisher). At 4 days post fertilization (dpf), the rate of mosaic expression of the fluorescent protein in the lens was scored. Clutches with 50% or more embryos showing mosaic fluorescent protein expression in the lens were raised to adulthood. Stable transgenic larvae were identified by out-crossing adults injected with the cxcr4b:H2A-GFP BAC transgene, and by raising larvae positive for the fluorescent transgenesis marker in the lens at 4 dpf. We screened 100 or more embryos from eight adults injected with the cxcr4b:H2A-GFP BAC transgene for fluorescent transgenesis marker expression in the lens and identified three founder fish. The full names of the transgenic lines identified are TgBAC(cxcr4b:H2A-GFP; cryaa:dsRed) p1 through p3. All experiments were performed according to guidelines established by the Stowers Institute IACUC review board.
In situ hybridization procedures were performed as described (Kopinke et al., 2006). The following probes were used: lef1, pea3, sef, axin2, fgf3, fgf10, fgfr1, dkk1b (Aman and Piotrowski, 2008), wnt10a (Lush and Piotrowski, 2014), deltaa, deltab, deltac, deltad (Itoh and Chitnis, 2001), dGFP, notch3, her4.1, atoh1a (Jiang et al., 2014), s100t (Venero Galanternik et al., 2015), shroom3 (Ernst et al., 2012), epcam (probe cb6 from ZIRC), celsr2 (Wada et al., 2006; Siddiqui et al., 2010), cldne kind gift from Ashley Bruce. The following primers were used to clone other probes: yap1 (Fw 5’-CGACTTTCCTTGAAAACGGT-3’ and Rv 5’-AAGGTGTAGTGCTGGGTTCG-3’), stk3 (Fw 5’-GCAGTGCTTCCTTAAACTCCAAAC-3’ and Rv 5’-GCAGGAATCTAGAGTAAGATGCAG-3’), amotl2a (Fw 5’-TGGAGAAGGTGGAAAGGATG-3’ and Rv 5’-GCTGGGCTCTTCTGAATCAC-3’), shroom1 (Fw 5’-CGTCTATGATGGGCAAACCT-3’ and Rv 5’-GGCAGGTCGTATGAGATGGT-3’), shroom2a (Fw 5’-AACAAGCAAACCCAATGGAG-3’ and Rv 5’-CTTTGGAGGCGAGTTGTAGC-3’), cdh1 (Fw 5’-TGGAAGAACAAGGACGCTCT-3’ and Rv 5’-TCTCAGGGACAGATGCAGTG-3’). The following probes were cloned into pPR-T4P vector using AA18 and PR244 primers: cldnb (Fw 5’-CATTACCATCCCGAAACGAAAAAGCATGGCATC-3’ and Rv 5’- CCAATTCTACCCGAGAGGCTGTTTCAAACGTGG-3’), cd9b (Fw 5’-CATTACCATCCCGTTGTGTTCACACACTCGCTG-3’ and Rv 5’- CCAATTCTACCCGACAACAGGACAACCACTCGC-3’).
The following primers were used to identify Tg(hsp70:ca-fgfr1)pd3 embryos by PCR: Fw 5’- GCAGCCTGACAGGACTTTTC-3’ and Rv 5’-GATCCGACAGGTCCTTTTCA-3’.
All embryos were injected at the one cell stage. The splice-blocking yap1 morpholino (5’-AGCAACATTAACAACTCACTTTAGG-3’) was injected at a concentration of 10 ng/2 nL (Skouloudaki et al., 2009). Translation blocking E-cadherin morpholino (5’-ATCCCACAGTTGTTACACAAGCCAT-3’) was injected at a concentration of 8.4 ng/2 nL (Babb and Marrs, 2004).
Embryos for Phalloidin staining were fixed for 30 min in 4% paraformaldehyde (PFA), washed 3 times for 5 min with 0.8% TritonX in PBS (PBT) and incubated for 2 hr at room temperature with Phalloidin (Invitrogen, 1:40 in 2% PBT). For immunohistochemistry embryos were fixed in 4%PFA at 4°C for the following antibodies: c-Myc (9E10) Santa Cruz Biotechnology (sc-40) (1:500), Anti-Acetylated Tubulin (T679) Sigma-Aldrich (1:500), mouse anti-ZO1 (Zymed; 61–7300) (1:200). Glyo-Fixx (Thermo Scientific, UK) at 4°C was used as a fixative for these antibodies: Di-pERK1/2 (Diphosphorylated ERK-1 and 2 antibody-mouse (M8159) Sigma-Aldrich) (1:500), Anti – ROCK – 2a (CT), Z – FISH- rabbit (AS-55431) (1:50), Mouse Anti-E-Cadherin (610182) BD-Biosciences (1:500), and Bouin’s fixative (Polysciences) was used for Phospho-Myosin Light Chain 2 (Ser19) Antibody (#3671 Cell Signaling (1:20)).
Samples were fixed in 2.5% glutaraldehyde and 2% paraformaldehyde in PBS for 1 hr at room temperature (then stored in 4°C until processing), followed by a secondary fixation in 1% aqueous osmium tetroxide with potassium ferricyanide overnight at 4°C. Samples were then dehydrated in a graded series of ethanol with propylene oxide as a transitional solvent, and infiltrated in Epon. Sections were cut on a Leica UC6 ultramicrotome in the range of 50 nm to 70 nm thickness and post stained with 5% uranyl acetate in 70% methanol and Sato’s lead stain. Sections were viewed in a FEI Technai Spirit BioTWIN TEM and imaged with a Gatan Ultrascan digital camera.
Samples were fixed in 2.5% glutaraldehyde and 2% paraformaldehyde in PBS for 1 hr at room temperature (then stored at 4°C until processing) and treated with tannic acid, osmium, thiocarbohydrazide, and osmium (TOTO) as described in Jongebloed et al. (1999). Samples were then dehydrated in an ethanol dilution series, critical point dried in a Tousimis Samdri 795 CPD, mounted on stubs and coated with gold palladium. Samples were imaged in a Hitachi TM-1000 tabletop SEM.
Heat-shock induction was done at various developmental time points, different temperature combinations and time intervals as indicated in the text. After the heat-shock, embryos were allowed to develop at 28.5°C.
BrdU incorporation, Hydroxyurea and Aphidicolin treatment (inhibition of proliferation) was performed as described in (Aman et al., 2011). Rat anti-BrdU (Accurate; OBT0030G) was used at a dilution of 1:500. To visualize nuclei embryos were placed into 0.1 ng/mL DAPI (Invitrogen). All chemical inhibitors were diluted in DMSO (final concentration of 1%) to concentrations as indicated in the text. Used drugs were purchased from Tocris, USA: Fgfr1a inhibitors PD173074 and SU5402, MAPK/ERK inhibitor PD0325901, GSK-3β inhibitor BIO and γ-secretase inhibitor DAPT.
Time-lapse imaging was performed as described (Lush and Piotrowski, 2014).
Individual cells were reconstructed in 3D using Bitplane’s Imaris software. Cells were reconstructed manually by drawing the cell boundary in every single z-slice using Imaris’ surface function. After the cell boundary was drawn in every z-slice, a surface was created, and the cell volume was exported for analysis.
For illustrative purposes, the cell surfaces were exported, oriented with the cell constriction upward, and placed side by side for comparison. Every attempt was made to export a view of the cells at the same scale, and at an angle, which shows the most representative perspective.
Rosette constrictions for each cell were analyzed using the ratio of two area measurements in 2D. First we measured the area of the bounding box of the smallest discernable constriction. Second we measured the area of the bounding box of the largest part of the cell. We then used the ratio of those two measurements to determine how much the cell is constricting on the apical side with respect to the bulk of its structure.
Rosette constrictions were analyzed using the Imaris surface function. Because so much membrane from multiple cells comes together at a single point, constrictions are the brightest feature in the image. A global threshold was set, selecting the brightest portion of the image. Smaller insignificant features were eliminated using a size threshold, leaving only the largest and brightest rosette constrictions.
Cell counting was performed in Fiji software where every DAPI positive nuclei was counted by hand using a Wand tool. An average number of cells between different samples was calculated and plotted with the standard error bars. Additionally, Student’s t tests were performed in Microsoft Excel software to assess the significance between the samples.
For the RNASeq experiment we cut tails from 36 hpf Tg(cldnb:lynGFP) embryos and isolated primordium and neuromast cells by FACS. Reads were mapped to the Danio rerio genome version danRer10 from the University of California, Santa Cruz (UCSC) using TopHat version 2.0.13, with gene annotations from Ensembl 84. RNASeq analysis and Gene Ontology (GO) Enrichment analysis was performed in the R environment using the bioconductor packages edgeR, topGO, and biomaRt. Reads counted on Ensembl transcripts from UCSC using HTSeq version 0.6.1 were analyzed with edgeR to generate P values to form comparisons between wildtype and NICD embryos. Of the 32,105 genes detected, only 19,643 genes with a sum across all samples of at least three reads per million were considered for the analysis. P values were adjusted for multiple hypothesis testing using the Benjamini–Hochberg method. The fragments per kilobase of transcript per million mapped reads (FPKM) values were generated using Cufflinks version 2.2.1. To define differentially expressed genes in the NICD embryos, genes were selected with the following criteria: log2 ratio >0.5 with p value <0.01 and log2 ratio <−0.5 with p value <0.01, resulting in 187 genes up- and 502 genes downregulated, respectively. The necessary GO IDs were obtained from Ensembl using biomaRt. Of the 19,643 expressed genes, 11,757 were associated with GO terms. Therefore, only 117 of the 187 genes and 323 of the 502 genes were used in the GO analysis.
The whole-genome sequence data from this study have been submitted to the NCBI Gene Expression Omnibus (GEO; http://www.ncbi.nlm.nih.gov/geo/), accession number GSE86571.
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Tanya T WhitfieldReviewing Editor; University of Sheffield, United Kingdom
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.
Thank you for submitting your article "Proliferation-independent regulation of organ size by Notch signaling" for consideration by eLife. Your article has been reviewed by three peer reviewers, and the evaluation has been overseen by Tanya Whitfield as the Reviewing Editor and Marianne Bronner as the Senior Editor. The following individual involved in review of your submission have agreed to reveal their identity: Ajay B. Chitnis (Reviewer #2).
The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.
This manuscript provides a detailed assessment of the role of Notch signaling in establishing neuromast size in the zebrafish lateral line. Previous studies from the Chitnis lab (Matsuda et al. 2010) described how loss of Notch signaling in mib1 mutants triggers problems in the morphogenesis of the lateral line primordium, including instability of deposited neuromasts and eventual disintegration of the primordium. In contrast to what was shown in this previous study, Kozlovskaja-Gumbriene et al. from the Piotrowski Lab have now shown, using a transgenic line expressing an active form of Notch (NICD), that broad activation of Notch in the primordium formation results in larger and fewer proneuromasts in the migrating primordium. This key observation now reveals a previously unappreciated, more direct role for Notch signaling in determining morphogenesis of epithelial rosettes. While this study confirms many observations made previously by Matsuda et al., the direct comparison of both gain and loss of function effects of Notch signaling allows them to go further by demonstrating the role of Notch signaling in inhibiting Wnt signaling and in acting downstream of FGF signaling to determine morphogenesis of epithelial rosettes.
The study also demonstrates that Notch has a direct role in determining formation and stability of epithelial rosettes, and now provides a clearer explanation for the early phenotype in Mib1 mutants associated with deposition of smaller unstable neuromasts. The study also reveals an important additional role played by Notch ligands expressed in the central Atoh1a expressing cell in both nascent and maturing proneuromasts. In addition to preventing neighboring cells from expressing atoh1a, activation of Notch contributes to formation of stable apical constrictions in neighboring cells. Contrary to previous studies that suggested FGF signaling-dependent shroom3 expression is required for morphogenesis of epithelial rosettes, this study shows that Shroom's role is dispensable.
For the most part, the reviewers found that the authors' conclusions are well supported by the data, and that the study provides an important mechanistic advance for the field. However, the reviewers had a number of comments where additional detail or discussion could help to clarify the findings. Most of these suggestions require relatively minor changes to the text or schematic diagrams. In a few places, additional quantitative analysis would help to support the conclusions. In addition, there were some places where it was felt that the conclusions were over-stated, and should be toned down.
1) Ensure descriptions of cell shape are accurate, and distinguish between quantitative and qualitative data (rev. 1).
2) State expected outcomes for the hypotheses that are being tested (rev. 1).
3) Include quantitative data to address the comment about mosaicism from reviewer 1, or tone down the conclusions from this section.
4) Consider changing the title (rev. 2).
5) Add a caveat about the control of expression of Ecad by Atoh1a (rev. 2).
6) Discuss or rebut the suggestion that control of wnt10a expression may be indirect (rev. 2).
7) Clarify the schematic of the 1st proneuromast (rev. 2).
8) Address and clarify the relationship between primordium and neuromast size in the NICD overexpression experiments (rev. 3).
9) State exactly how the cell counts were done, and provide additional data to clarify the relationship between NICD, Yap1 and primordium cell number, if possible (rev. 3).
10) Ensure that the schematics are close representations of the data shown (rev. 3).
11) Make the manuscript shorter and more concise where possible (rev. 3).
Detailed versions of the reviews are appended below for further information.
1) In the subsection “A reduction in Notch leads to loss of cell-cell adhesion and apical constrictions”, Figure 6: The data presented in Figure 6 are difficult to interpret. First, the rows of isolated cells in panels A-C are difficult to relate to their counterparts in the intact images. Cell shapes in L1 appear similar in NICD and mib embryos but are quite distinct from wt, yet are described as normal. Cell constrictions in L2 are said to be lost in mib mutants. This is evident in some cells (e.g. blue and pink) but less so in others. And the pink cell in L1 in NICD also appears to lack an obvious apical constriction. Somehow interpretations in this part of the paper seem highly subjective. Note, Figure 6D is labeled to direct attention to Video 5, whereas the text mentions Video 4. Figure 6E-H marks "constriction volume", but it is not clear what this means – and is this quantitative as the term suggests, or strictly qualitative as the images suggest? In the last sentence it is stated that the phenotypic defects in mib mutants become progressively more severe "because of maternal rescue". It would make more sense to say the phenotype worsens as maternal stores become depleted.
2) In the subsection “Notch-positive cells self-organize into rosettes”, Figure 7: Mosaic analysis is often a powerful means of distinguishing between competing hypotheses, but it is important to clearly articulate expected outcomes that might support one hypothesis over the other. In this case expectations are not discussed, nor is it clear whether data support the "signaling center" hypothesis or the "altered cell adhesion" hypothesis. With either model, one might expect that the number of wt cells recruited into a neuromast might vary depending on the number of transgenic cells present. Yet there was no attempt to correlate neuromast size with the percentage of wt vs. transgenic cells. Moreover, it is concluded that wt cells were "never recruited to make neuromasts larger than they would normally be". The basis for this is unclear. The image in Figure 7B" appears to show a larger than normal neuromast that includes a mix of both wt and NICD cells, in apparent contradiction of the conclusion. Size measurements and cell counts (wt & NICD) are needed to make a convincing argument. Other data in Figure 7 are more convincing. On a technical note, the high mag image in 7G' is inverted horizontally relative to the low mag image in 7G.
While overall the paper is excellent and reveals another example of a context in which Notch has a more general role in promoting an epithelial morphology, there are a few details related to interpretation or emphasis with which I did not agree. These issues are enumerated below.
1) I thought the title of the paper, which emphasizes a role for Notch signaling in organ size formation is misleading. While in the gain of function experiments many more cells can be recruited to become part of an epithelial rosette, as noted above, this gain of function phenotype does not represent that pattern of Notch activation or epithelial rosette formation seen under normal circumstances, where formation of rosettes is beautifully coordinated by a central cell that becomes the source of FGFs and Notch ligands. Instead, I believe the paper provides excellent evidence for a role for Notch signaling in not only preventing neighboring cells from expressing atoh1a but also in determining formation of stable epithelial rosettes by activating Notch in the neighbors. The size of these rosettes may, under physiological conditions, still be determined by FGF signaling-though, now as we have learnt from this study, not only by shroom or MAPK dependent mechanisms as previously described but in addition by Notch signaling. As the authors themselves state, by determining Notch3 expression, FGF signaling may still play a critical role in determining which how many cells are recruited to become part of the rosette by Notch signaling. The gain of function experiments with NICD do, however, show how in the absence of all patterning determined by interactions between Wnt and FGF signaling systems, Notch signaling can independently determine a particular configuration of epithelial rosettes in the primordium.
2) In the subsection “Notch signaling upregulates cell-cell adhesion molecules and tight junction genes”, second paragraph: the authors show that e-cadherin expression is promoted by Notch signaling. While data in this paper supports this conclusion, it remains possible that in addition to being promoted by Notch signaling, its expression is inhibited by Atoh1a as suggested by Matsuda et al., since knock-down of atoh1a leads to some recovery of e-cadherin expression in mib1 mutants.
3) "To test if Notch signaling restricts wnt10a expression we treated apc mutant embryos with the γ-secretase inhibitor DAPT (Figure 9M', N'). Indeed, wnt10a is now expressed in many more trailing cells demonstrating that Notch inhibits wnt10a, a feedback interaction not previously described in the primordium (Figure 8A). This negative interaction also explains why wnt10a is restricted to a single central cell in the first forming proneuromast of a wildtype primordium (Figure 9M)".
While this observation superficially supports the assertion of the authors that Notch signaling restricts Wnt10a expression, I feel what is happening here might be more complicated and only indirectly related to Notch signaling. One problem is that the broad spread of wnt10a expression seen in apc mutants is only seen following 20 hours of exposure to DAPT. Does it spread with a shorter treatment? If not, I suspect that the expansion of wnt10a expression is the indirect effect of something that takes a while to develop. One possibility is that it is related to progressive slow expansion in the number of atoh1a expressing cells. As Atoh1a inhibits expression of the FGF receptor, it would therefore permit Wnt signaling in this cell, which is so close to the source of Wnt signals. This may explain why transiently or in an oscillatory pattern Wnt10a is expressed in a spot just adjacent to the leading cloud of wnt10 expression, where it is expressed under normal circumstances. Before the authors can conclude that the restricted pattern is determined by lateral inhibition they should determine if knock-down of atoh1a prevents the expansion. Alternatively, they can skip the experiment and suggest this alternate hypothesis.
4) The schematic for signaling in the 1st proneuromast (most recently formed?) has a number of problems. Notch3 is not being activated by DeltaA/D but by Ras-Mapk which confuses transcriptional regulation of Notch3 and actual activation by its ligand. If there are details the author feels are not well known at this stage they should consider dropping the schematic of early proneuromast or revise it to make it accurate. I feel what’s most clear from their paper is what is presented for the maturing proneuromast.
This paper studies the role of increasing Notch signalling on lateral line development. How organ size is regulated by balancing cell proliferation and cell differentiation is an important question and the lateral line has been a good experimental model to address these questions. The authors deepen into them, and here they propose that in contrast to Wnt signaling that increases neuromast cell numbers by increasing cell proliferation, Notch regulates organ size independently of cell proliferation and by regulating cell adhesion and rosette number in the primordium. The authors combine several approaches and provide an extensive analysis of Notch effects. The work provides interesting, but not fully novel data and some of the conclusions are precipitated and not supported by their data. The link between the effects on cell adhesion and NMs size is not clear. On a side, there are several functional experiments addressing NICD effects on NMs size. On another set of experiments, the authors address the effects of NICD on rosette formation analyzing the primordium without showing the effect on deposited NMs size. In overall, the paper is very long, difficult to follow and without a clear-cut idea of the Notch role on NMs size.
1) Role of NICD in proliferation
The authors make strong emphasis in demonstrating that Notch overactivation increases neuromast size independent on effects on cell proliferation. The authors claim that the size of NICD NM is independent of primordium size (figure legend, Figure 1—figure supplement 1) and this is not fully supported by their data. When overexpressing NICD early by combining cdnB:gal4 with UAS:NICD, the size of the primordium is bigger (while ganglion smaller) and in this condition, the L1 instead of containing approx 35 cells contains 110. Through development, as pLLP decreases its size, the NM size also decreases, indicating that the size of the primordium has an influence on NM size. If NICD is overexpressed after 25 hpf, the L1 or L2 NM only increases its size from 35 cells to 45-55. The authors suggest that as the size of the primordium in this condition is not bigger (due to the effect of ganglion-primordium fate switch) but the NM size is still bigger, the size of the primordium is not the cause of NM increase. However, in this later condition, the effect over NM size is much milder, indicating that if NICD is overactivated early, this has a very strong effect of first neuromasts deposited. How the authors explain that, if the size of the primordium has no effect on NM size, the early effects are much stronger than in the hs:NICD. It is not clear why all the data is shown in embryos in which NICD has been manipulated early instead of hsp:NICD to overcome possible effects of initial primordium size. The authors should also provide data on the size of the primordium in hs:NICD experiments.
Again, the authors state "whereas the increase in the size of NICD neuromasts is controlled by a proliferation-independent process within the primordium".
The difference in the proliferation status in NICD NMs and apc NMs is clear but the authors also show that the BrdU index at the NICD primordium is higher than controls and similar to apc mutants (Figure 3H, although stated that does not change). Could the increased proliferation at the primordium affect deposited NMs size?
The authors also show that the neuromast size in NICD is independent of yap1, while there is a strong reduction in primordium cell numbers in NICD-yap MO embryos (Figure 2G), the size of the NMs is not significantly reduced in this condition. In this particular experiment the deviation bars are higher, thus this mild reduction might become significant if increasing the number of embryos analysed. This is important because is one of the major conclusions of the paper. I suggest to repeat this experiment and add more embryos. Throughout the paper is not clear, which NMs are counted for cell numbers and at which stage the counting is done.
Figure 1P shows that the relative difference of size of wt and NICD NM is reduced overtime, however in Figure 3E this difference is kept. Please explain incongruence. As part of this, it is stated in the text that NICD neuromasts do not significantly grow in cell number (Figure 3E) but instead the NICD NM size increases over time as shown in this figure. How this happens if cell proliferation decreases so drastically in NM (Figure 3D)?
In the schematic representation of the primordium shown in Figure 4, the NICD primordium is not bigger. However, Figure 6B (primordia shown are at different stages!) shows a much bigger primordium in NICD. In particular, the size of the leading part is bigger (not depicted in the drawing). Again, the changes in size of the primordium and in particular the ratio of leading and trailing cells might be relevant for the neuromast size of NICD embryos. This is very little discussed or addressed.
2) Role of NICD in cell fate specification
This section is confusing. If by cell fate the authors refer to the switch between the 4 cell types defined in the primordium, it is clear that there is not a major cell fate switch. The authors conclude that NICD causes the allocation of support cells into fewer but larger proneuromasts and this is clear. But, as the authors mention later, within the primordium, the first forming proneuromast and more trailing proneuromasts are in different stages of cell fate specification due to different activities of FGF signaling. Since Notch affects FGF and Wnt signaling, it is possible that the proneuromast lost is converted to the fate of the other proneuromasts. It is possible that the disappearance of a proneuromast is due to the cell signaling effects and changes of proneuromast commitment stage and not due to increased recruitment of Rock2a that would be secondary. Please discuss this.
3) Cell adhesion:
The authors claim that the main cause of increased NICD neuromast size is their increased adhesivity, more cells sticking together, but not effects on rosette formation are observed when blocking some cell adhesion molecules. In addition, the conclusion is also taken that interneuromasts cells cluster more together in NICD, but these are cells not related to the focus of the study. The conclusion is strong for the little evidences on the cell adhesion and I would suggest to tune-down the conclusions. As in NICD more cells compose a proneuromast, it is obvious that the E-cadherin staining is going to be larger (Figure 7E). Moreover, in the NICD primordium one of the rosette disappears making the proneuromast larger, but if cell adhesivity is increased, why not more rosettes are formed instead? The authors suggest that increased cell adhesion, affects the morphology of the organ to achieve minimal surface tension. If data from microsurgery experiments to measure of surface tensions in wt and NICD cells are available, this would be an interesting addition, but is not essential.
4) Notch and FGF/wnt signaling
The epistatic data showing that Notch is downstream of Wnt and FGF signaling is clear. In parallel Notch negatively feeds-back in FGF signalling by reducing fgf10a, fgf3 expression (not clear about pea3). On the other hand, NICD reduces the levels of Lef1, sef1, dkk1b and wnt10a at the leading region. The effects on Wnt signalling are quite strong and previous reports have analysed the influence of the disruption of Wnt in proliferation and neuromast deposition. The authors make strong emphasis showing that the effects on NMs size is different that the phenotype caused by apc mutants. However, since NICD modifies the balance of FGF/Wnt signalling, it is not clear that Notch and Wnt are not linked in the regulation of primordium size and differentiation. In NICD embryos, what happens if Wnt signaling is raised by crossing with apc mutants? If this is known, please discuss.https://doi.org/10.7554/eLife.21049.028
- Tatjana Piotrowski
The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.
We would like to thank the Piotrowski lab members and Drs. Krumlauf, Gibson, Li and Trainor for stimulating discussions and Dr. Mark Lush and Joaquin Navajas Acedo for critically reading the manuscript. We are thankful to Boris Rubinstein for a help with mathematical analysis. We are grateful to Dr. Ajay Chitnis for mib1ta52b mutant, Dr. Bruce Riley for Tg(hsp70:atoh1a)x20, Dr. Virginie Lecaudey for the shroom3 morpholino and in situ probe, Dr. Brian Link for the UASDNshroom3 plasmid, Dr. Ashley Bruce for the claudinE in situ probe, Dr. Anand Chandrasekhar for the cdh2hi3644 fish line and Xin Gao for help with the RNASeq analysis. We would also like to thank Helena Boldt for excellent technical support. We are also grateful to Mark Miller for help with illustrations and the Stowers Aquatics facility for excellent help with fish husbandry. We are particularly thankful to the Stowers Institute for Medical Research for Funding.
Animal experimentation: All experiments were performed according to guidelines established by the Stowers Institute IACUC review board (IACUC protocol 2014-0129)
- Tanya T Whitfield, Reviewing Editor, University of Sheffield, United Kingdom
- Received: September 2, 2016
- Accepted: December 23, 2016
- Version of Record published: January 13, 2017 (version 1)
© 2017, Kozlovskaja-Gumbrienė 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.