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Work in mouse has implicated cilia motility and leftward nodal flow as the mechanism for breaking symmetry. In zebrafish, it is assumed that Kupffer’s vesicle is analogous to the mouse node. However, its architecture is different and the fluid dynamics inside Kupffer’s vesicle is not completely understood. We show that cells lining both the dorsal roof and the ventral floor of Kupffer’s vesicle possess posteriorly pointed cilia that rotate clockwise. Analysis of bead movements within Kupffer’s vesicle shows a net circular flow but the local flow differs in direction depending on the location within the vesicle. Histological analysis suggests that the orientation of the cells at anterior-dorsal region likely direct net flow in the vesicle. Our data suggest that the plane of the circular net flow is tilted with respect to the D-V axis, which may be converted to a local leftward flow in the anterior-dorsal region of the vesicle.
Vertebrates appear outwardly bilaterally symmetric, but have internal asymmetries along the left-right (L-R) axis. This axis is revealed by the asymmetric placement of organs along the midline. Exactly how L-R asymmetry is established during early vertebrate embryogenesis is still under considerable debate, however. The L-R axis is defined after the dorsal-ventral (D-V) and anterior-posterior (A-P) axes (for reviews, Capdevila et al., 2000; Takaoka et al., 2007). The first observable evidence for L-R patterning in all vertebrates is asymmetric gene expression in the lateral plate mesoderm (LPM), which subsequently leads to proper organ laterality in fish, frog, chick, mouse, and rabbit (for reviews, Burdine and Schier, 2000; Bisgrove and Yost, 2001; Speder et al., 2007). However, the event that constitutes the earliest break in symmetry, and how this leads to asymmetric gene expression, still remains an open question (for example see, Tabin, 2005).
It has been shown that cilia in the mouse node rotate in a clockwise direction when viewed from the ventral side (Figure 1, A1). These cilia are tilted toward the posterior (Nonaka et al., 2005; Okada et al., 2005). Therefore, the local flow above individual cilia is rotational (Figure 1, thin, black arrows), but the rightward swing of the cilium close to the cell surface retards the fluid (Figure 1, A2, thick, grey arrow at the node surface). The net flow at the node is from right to left (Figure 1, A2, net flow), driven by the effective stroke of the tilted cilia (Figure 1, A2, thick shaded arrows). This nodal flow is considered to be conserved in Xenopus (Schweickert et al., 2007), medaka, Oryzias latipes, (Tanaka et al., 2005), and rabbit (Tanaka et al., 2005; Blum et al., 2007). Broadly speaking, this linear flow has been proposed as the initial break in L-R symmetry for all vertebrates (Nonaka et al., 1998; Nonaka et al., 2002; Okada et al., 2005).
The zebrafish, Danio rerio, has a transient ciliated organ called Kupffer’s vesicle that is derived from the dorsal forerunner cells (DFCs) (Cooper and D’Amico, 1996; D’Amico and Cooper, 1997). Currently, it is assumed that Kupffer’s vesicle is analogous to the mouse node in terms of left-right patterning (Essner et al., 2002). However, the fluid dynamics inside Kupffer’s vesicle is not completely understood. While the ciliated surface of the mouse node is relatively flat, Kupffer’s vesicle is a hollow sphere containing cilia, projecting both from dorsal roof and ventral floor (Amack et al., 2007; Kreiling et al., 2007). The first report of cilia motility in Kupffer’s vesicle proposes that cilia rotate counterclockwise when viewed from the apical side (Figure 1, B1), which is opposite to what is observed in the mouse node (Kramer-Zucker et al., 2005). This observation was repeated by (Shu et al., 2007), obtained from a dorsal view of Kupffer’s vesicle, but it is unclear which side of the cilium was observed to reach this conclusion. Later analysis of 3D images found that cilia are concentrated in the dorsal-anterior region of the vesicle, suggesting that these cilia cause the dominant counterclockwise flow in Kupffer’s vesicle (Kreiling et al., 2007). However this report did not use live imaging to determine whether this is in fact the case. Although a previous report implicates tilted cilia in creating a strong local leftward flow (Kramer-Zucker et al., 2005), the observed net flow seems circular and counterclockwise at the center of the vesicle (Figure 1, B2, thick arrow) based on bead movement and computational analysis (Essner et al., 2005; Kawakami et al., 2005; Ellertsdottir et al., 2006; Shu et al., 2007, Kreiling et al., 2007). Thus, we undertook a set of studies to determine the accurate fluid flow inside Kupffer’s vesicle and how it is generated, considering the overall shape of the vesicle (Figure 1, B2).
To better understand the flow produced by cilia in Kupffer’s vesicle, we carefully analyzed the morphological structure of the vesicle in zebrafish using Scanning Electron Microscopy (SEM) and histological sections. Our results suggest that the majority of the cilia in Kupffer’s vesicle are tilted toward the posterior of the embryo. Using video microscopy, we demonstrated that all cilia in Kupffer’s vesicle rotate clockwise (when viewed from the apical side of the cell) analogous to what is observed in the mouse node. Additionally, our imaging demonstrates that the bead movements around an individual cilium reflect cilia motility locally, and thus we see beads moving in opposite directions at the dorsal roof and ventral floor. However, the dominant flow in the center of Kupffer’s vesicle is counterclockwise (when viewed from the dorsal side of Kupffer’s vesicle), and is likely dictated by the presence of more ciliated cells on the dorsal roof than on the ventral floor. Our results furthermore suggest that the plane of the circular net flow within the vesicle is tilted, and thus cells in the anterior-dorsal region of the vesicle may experience a local dominant leftward flow. Finally, we have hypothesized that net flow in zebrafish Kupffer’s vesicle is analogous to flow in the mouse node and medaka fish in terms of left-right patterning even though the ciliated cell structures appear to differ in architecture.
To determine the similarities and differences between the mouse node and Kupffer’s vesicle, we first observed the ultrastructure of Kupffer’s vesicle by Scanning Electron Microscopy (SEM). Fixed embryos were divided into two segments; the body part including the dorsal roof of Kupffer’s vesicle (Figure 2, A, C), and the yolk part including the ventral floor of Kupffer’s vesicle (Figure 2, B, D). The dorsal half sphere of Kupffer’s vesicle was located at the end of the notochord (Figure 2, A1, C1, nc, KV) and the ventral half sphere was located as an indention into the yolk (Figure 2, B1, D1, KV). Cilia were observed from the 3-somite stage (Figure 2, A, B) to the 11-somite stage (Figure 2, C, D) on all cells lining the dorsal roof and ventral floor cells of Kupffer’s vesicle. The size of Kupffer’s vesicle became significantly larger at later stages (Figure 2, A2, B2, C2, D2). Likewise, cilia grew longer at later stages (Figure 2, A2, B2, C2, D2, arrows). Consistent data were observed by immunolabeling cilia in Kupffer’s vesicle (Supplemental Table I). By SEM, cell borders were easily observed between the individual cells (Figure 2, C2, D2, arrowheads), suggesting that a monocilium projects from each cell of both the dorsal and ventral sides of the vesicle, consistent with previous reports (Amack et al., 2007; Kreiling et al., 2007). Similar results were obtained from 4-10 somite stages (data not shown). Furthermore, the cilia located in anterior region appeared to be pointed toward the posterior (Figure 2, A3, B3, C3, D3, arrows). We often observed bending cilia in this region, suggesting a stroke pattern similar to what is predicted in the mouse node (Figure 2, A3, C3, yellow arrows). However, unlike cilia in the mouse node, we do not observe the base of cilia to be significantly shifted towards the posterior with respect to the cell using this method (Figure 2, C2-C3, D2-D3). At 7-9 somite stages, the cilia at the dorsal-anterior regions are pointed more towards the posterior and often exhibited bending (Figure 2, E1, E2, yellow arrows). Thus, cilia are likely to point towards the posterior, which has been suggested previously (Kramer-Zucker et al., 2005).
To clarify whether cilia at different locations within Kupffer’s vesicle have the same type of motility, we performed high-speed video microscopy. A water immersion objective lens (x60) was set against the dorsal side of the embryo facing Kupffer’s vesicle (which we refer to as the dorsal view) and focused to different planes within the vesicle (Figure 3, A). Using differential interference contrast (DIC) microscopy, we observed the live motion of cilia rotation at the dorsal roof (top surface closest to the objective, Figure 3, A, line D), the middle plane between the dorsal and ventral sides (Figure 3, A, line M), and the ventral floor (the bottom surface farthest from the objective, Figure 3, A, line V). From a dorsal view, cilia on the dorsal roof rotated counterclockwise (Figure 3, C1-C5, supplemental data; video 1), and ventral floor cilia rotated clockwise (Figure 3, D1-D5, supplemental data; video 2). Cilia at the lateral edges of Kupffer’s vesicle also rotated (supplemental data; video 3). Our earliest observation of cilia motility began at the 3-somite stage (number of the embryos (Nemb)=3, number of the cilia (ncilia)=7), our latest measurement was taken at the 11 somite stages (Nemb=1, ncilia=7). The direction of cilia rotation was consistent from 4-11 somite stages (data not shown).
It is important to note that cilia on all cells in Kupffer’s vesicle rotate clockwise (when viewed from the apical side of the cell), which differs from a previous report (Kramer-Zucker et al., 2005). However, cilia on the dorsal roof were observed from the basal side of the cell, thus the rotation appears different from those on the ventral floor. We assume this is the reason why our data conflicts with the previous conclusion. Ventral floor cilia, however, were viewed from the apical side of the cell, which is analogous to how cilia motility was viewed in the mouse node. If it were possible to view through the zebrafish yolk and ventral floor of the vesicle towards the apical side of the dorsal roof cells, the cilia would then rotate clockwise in video recordings. Thus, all cilia in Kupffer’s vesicle rotate clockwise similar to what is observed in the mouse node (Nonaka et al., 1998; McGrath et al., 2003; Okada et al., 2005; Nonaka et al., 2005). We measured the frequency of cilia rotations at different stages (Table I). The average frequency of cilia motility was not significantly different from the 3 somite stage onward.
Additionally, cilia imaged at the dorsal-anterior region often showed an elliptical trajectory at the tip, suggesting that cilia are pointed toward the posterior when viewed from the dorsal side of Kupffer’s vesicle (Figure 3, E, dotted lines, F1-F5, and supplemental data video 4). If cilia are straight and not tilted, the trajectory of the cilia rotation should be circular (Figure 3, D1, and F5, blue cross). To further analyze how many cilia point towards the posterior, we plotted the total images of cilia motility at 7-8 somite stages and summed the trajectories of cilia at the dorsal roof and ventral floor where the ciliated cells are located within a relatively flat sheet (Figure 3, F1-F5). If these regions of the dorsal roof or ventral floor are as flat as the cellular surface of the mouse node, the angle of cell orientation can be eliminated as a factor in the analysis. The sum of cilia trajectories indicates the direction of the cilia tip. In these regions, we find that more cilia point toward the posterior (Figure 3, F1-F4, red crosses, Table II), compared to cilia not pointing toward the posterior (Figure 3, F1, F2, F5, blue crosses, Table II). More than 60% of the dorsal roof and 70% of the ventral floor of cilia pointed towards the posterior. Although SEM analysis shows bending cilia predominantly in the dorsal-anterior regions, the trajectories of cilia motility reveal posterior pointing of cilia not only at the dorsal roof but also at the ventral floor of the vesicle (Table II). The posterior tilt of the cilia at the ventral floor suggests the direction of fluid flow in this location would be opposite as compared to the dorsal roof.
While all cells in Kupffer’s vesicle possess cilia that rotate clockwise when viewed from apical side, the fact that the dorsal roof cells face the ventral floor cells presents an intriguing issue. As the cilia move simultaneously, the direction of flow at the dorsal roof is predicted to be opposite the direction at the ventral floor. To date, all measurements of flow in Kupffer’s vesicle have been performed by imaging fluorescence beads by confocal or epi-fluorescence microscopy in one focal plane. It is assumed that previous observations have focused where movement of injected beads is the most obvious. To determine if the flow at different levels within Kupffer’s vesicle varies, we injected 0.2 um diameter beads into the vesicle and observed their movements from the dorsal side using DIC microscopy (Figure 3, A, arrow, “dorsal view”). We observed bead movements at three different levels; dorsal roof, middle, and ventral floor (Figure 3, A, line D, M, V, respectively; three independent experiments, Nemb=2 for each, 5, 7, 8, or 10 somite stage, Nemb=4 for 9 somite stage). At the dorsal roof, the majority of beads moved counterclockwise, although occasionally individual beads were observed moving linearly or randomly (Figure 4, A1, supplemental data; video 7). At the ventral floor, beads were often observed moving clockwise (Figure 4, A2, supplemental data; video 8). At the middle level, movements were a randomized mixture of a dominant counterclockwise flow with a few clockwise movements (supplemental data; video 9). In one experiment, images taken at the middle level showed movement of a bubble introduced during injection as counterclockwise along with the dominant bead movements (Figure 4, B1-B8, arrows). In conclusion, both clockwise and counterclockwise motions exist in Kupffer’s vesicle depending on the location, but the dominant flow is counterclockwise as observed from a dorsal view. To insure that bead injections did not interfere with left-right patterning, embryos were injected with beads and fixed at 18-19 somite stages, followed by RNA in situ hybridization analysis for southpaw, lefty1, and lefty2. 13/15 injected embryos had normal left-sided expression of these genes, indicating left-right patterning was not altered by bead injections in this experiment (data not shown).
To better understand how the dominant counterclockwise flow in the middle of Kupffer’s vesicle is generated, we performed a histological analysis. Around the 3 somite stage, Kupffer’s vesicle was visible and already possessed an open interior space (Figure 5, A1, B1). At the 9 somite stage, the structure of Kupffer’s vesicle is more ellipsoid, and the two layers of cells lining the dorsal roof and ventral floor were easily visible (Figure 5, B1, B2, B3). Although it was not obvious from our SEM analysis, histological analysis suggests that the cells lining the dorsal roof appear more numerous than cells lining the ventral floor (Figure 5, B1, B2, B3). The difference was also apparent in DIC images showing cell borders (Figure 3, B1-B2, dotted lines). Furthermore, we observed that the cells were more densely formed at the anterior edge of Kupffer’s vesicle at the 5 and 9 somite stages (Figure 5, B2, B3, brackets). This data is consistent with results from 3-dimentional reconstruction of immunofluorescently labeled cilia at Kupffer’s vesicle, which suggested the presence of more cilia in anterior-dorsal side (Kreiling et al., 2007). Similar to the findings of Kreiling et al, our data also suggests that the dominant counterclockwise flow in the middle of Kupffer’s vesicle is driven by the more numerous cilia at dorsal-anterior roof. In addition, the cell axes in the dorsal-anterior roof were orientated toward the ventral-posterior, and the surface of these cells face into the center of ellipsoid. The orientation of these cells, coupled with the posterior tilt of the cilia, suggests the flow plane in Kupffer’s vesicle is tilted with respect to the D-V axis (Figure 6).
Kupffer’s vesicle is a transient liquid-filled organ in teleosts. In zebrafish, the vesicle has cells on both the dorsal and ventral surfaces (Amack et al., 2007; Kreiling et al., 2007; this report), and is derived from a specific population of endoderm cells called the dorsal forerunner cells (Cooper and D’Amico, 1996; D’Amico and Cooper, 1997). By contrast, Kupffer’s vesicles in medaka and Fundulus heteroclitus, are composed of a single layer of columnar cells only at the dorsal roof (Brummett and Dumont, 1978; Okada et al., 2005), and in medaka these cells do not express the endodermal markers casanova and sox17 (Hojo et al., 2007). This suggests that the characteristics of Kupffer’s vesicle differ within fish species. Thus, it is important to compare these ciliated organs in different species with regard to L-R patterning. In the mouse node, ciliated cells are located on the ventral side of the embryo, and thus the cilia project to the outside (Figure 5, C1) (Vogan and Tabin, 1999). This planar structure of ciliated tissue is analogous to that of fish like medaka, which have only one epithelial layer on the dorsal roof (Figure 5, C2) (Okada et al., 2005; Hojo et al., 2007). However, zebrafish Kupffer’s vesicle is composed of two layers of cells having the cilia both at the dorsal roof and the ventral floor (Figure 5, C3) (Amack et al., 2007; Kreiling et al., 2007). Although previous studies have suggested this to be true using immunohistochemistry and TEM (Kreiling et al., 2007) neither technique distinguishes from where cilia originate. Our SEM analysis presents intact cilia and cell membranes allowing us to definitively state that cells on the ventral floor and dorsal roof each possess monocilia. Furthermore, our SEM and video microscopy demonstrate that numerous cilia within the vesicle are tilted to the posterior. Previously, a section of TEM analysis showed one cilium in zebrafish Kupffer’s vesicle to be tilted 45° relative to the surface of the roof (Kramer-Zucker et al., 2005). Although the posterior tilt of the cilia in the mouse node is thought to be due to a shift of the base of the cilia to the posterior, we could not confirm that this occurs at high frequency in Kupffer’s vesicle. Although the SEM technique has limitations for observing depth inside the curved Kupffer’s vesicle, posterior positioning of the cilia base is not obvious. In addition, in mouse, nodal pit cells appear to be rounded at the surface of the node. This cell shape coupled with the posterior location of the cilia base is thought to result in the tilt of each cilium (for review see Shiratori and Hamada, 2006). Our SEM and histological analyses do not suggest a similar convex shape of the cell surfaces in Kupffer’s vesicle. Instead we observe more flat or even concave shapes of cell surfaces, which would be unlikely to contribute similarly to cilia tilt. Thus we think it is more likely that the combination of cilia bending and cell orientation of Kupffer’s vesicle causes cilia to be pointed towards the posterior.
Recent studies have reported counterclockwise cilia motility in zebrafish when viewed from dorsal side (Kramer-Zucker et al., 2005; Shu et al., 2007). Using our video microscopy technique, we have analyzed cilia motility at different levels within Kupffer’s vesicle. In conclusion, all cilia rotate clockwise when viewed from the apical side of the cells. This is clearly conserved in the mouse node and in the Xenopus gastrocoel roof plate, which have clockwise rotating cilia when viewed from the ventral side (Nonaka et al., 2005; Okada et al., 2005; Schweickert et al., 2007). However, we did not obtain evidence for non-motile cilia, which is different from the mouse node (McGrath et al., 2003).
Work in the mouse has demonstrated that cilia protrude from node cells, and are posteriorly tilted, most likely due to the posterior position of the basal bodies (Nonaka et al., 2005; Okada et al., 2005). The tilt of the cilium results in surface interactions between the cilium and the cell membrane. The surface viscosity of the membrane diminishes the rightward force of the cilia rotation, resulting in a dominant leftward flow in the mouse node (Cartwright et al., 2004; Brokaw, 2005). Structures in medaka and rabbit are considered analogous to the mouse node based on the planar structure of the ciliated layer (Okada et al., 2005; Blum et al., 2007). Although most studies on flow in zebrafish Kupffer’s vesicle have shown counterclockwise movements (Essner et al., 2005; Kawakami et al., 2005; Kramer-Zucker et al., 2005; Ellertsdottir et al., 2006; Shu et al., 2007), most were done by injecting fluorescence beads which were then observed using confocal or epi-fluorescence microscopy. The advantage of DIC imaging is that we can be specific about the location and level within Kupffer’s vesicle by observing the edge of Kupffer’s vesicle. Our live images present detailed locations of beads at the dorsal roof, the ventral floor and in the center of Kupffer’s vesicle, providing the observation of the local flow around an individual cilium and of the strongly dominant flow circulating within the entire vesicle at the same time. Thus, these analyses allow us to determine what constitutes the net flow. Although cilia rotate in opposite directions at the dorsal roof and ventral floor, the dominant flow inside zebrafish Kupffer’s vesicle is counterclockwise (when viewed from the dorsal side of the cells) and correlates with cilia movement on the dorsal roof of the structure. Our histological analysis and DIC image are in agreement with previous data that suggested cilia are more numerous at the dorsal roof (Kreiling et al., 2007). If we focus only on the dorsal roof, the morphological structure of zebrafish Kupffer’s vesicle is similar to other fish such as medaka. Our observations of bead movements in the closed space of zebrafish Kupffer’s vesicle show dominant flow is circular and not linear as reported in mouse. Previous studies suggest that the speed of bead movements from right to left is significantly faster than that from left to right in zebrafish (Kramer-Zucker et al., 2005), but there is still high probability of rightward flow in Kupffer’s vesicle, compared to medaka Kupffer’s vesicle (Okada et al., 2005). It is important, however, to note that the mouse node is initially covered by Reichert’s membrane, and in vivo flow takes place in a closed space. However, under in vitro experimental conditions, Reichert’s membrane is removed and thus return flow is not detected (Cartwright et al., 2007). Considering that zebrafish Kupffer’s vesicle is a closed system, we suggest that the flow we observe is closer to what occurs in vivo in the mouse. However, the question still remains as to how the return rightward flow might influence L-R asymmetry.
In the mouse node, motile cilia are assumed to be located at center with non-motile cilia located around the outer edge (McGrath et al., 2003). If these center cilia project straight towards the outside, the rotational motility would cause a vortex flow on the top of the cilia (Figure 6, A1). Recent observation that the cilia are posteriorly tilted explains how the rotation can result in a dominant leftward flow (Figure 6, A2-A3) (Nonaka et al., 2005; Okada et al., 2005). If cilia at the center of the dorsal roof of Kupffer’s vesicle are projected straight into the vesicle, this would also produce a counterclockwise vortex directly above the cilia. Moreover, cilia on the ventral floor would cause a vortex in the opposite direction (Figure 6, B1) but this is not what we observe. Even if the cilia at the center of the dorsal roof are tilted toward the posterior, dominant flow would be predicted to be leftward at the dorsal side and rightward at the ventral floor (Figure 6, B2). If this is coupled with posteriorly tilted cilia at the periphery of the vesicle, the flow would move from up to down on the left side and down to up on the right side. Overall, this would result in a circular flow that is parallel to the D-V axis when viewed horizontally from the posterior. (Figure 6, B3). In this case, the trajectories of beads would be linear when viewed from the dorsal side, but this is also not what is currently observed.
We did observe that the surface of cells on the anterior-dorsal roof of Kupffer’s vesicle face the center of the ellipsoid and are tilted with respect to the D-V axis of the embryo. If the cilia on these cells were not tilted, the circular plane of flow would no longer be parallel to A-P axis, but would still be a vortex over the tips of the cilia (Figure 6, C1). However, the anterior-dorsal cells do have tilted cilia and therefore, this population of cells will produce the strongest fluid force in a leftward direction (Figure 6, C2). It is important to note that, the architecture we report will also result in the circular flow plane being tilted off the D-V axis due to the angle of the anterior-dorsal roof cilia and cells (Figure 6, C3). Even though rightward flow exists at the posterior-ventral floor, it is likely to have a negligible effect on the anterior dorsal cells. A recent report using a technically distinct imaging approach described micrometer scale observations of cilia-driven flow surrounding a single beating cilium in Kupffer’s vesicle (Supatto et al., 2008). Their technique demonstrated that the rotational axis of the cilium is tilted towards the dorsal direction. These results concur with the predictions we make in our model.
We predict that this overall tilted circular motion of fluid dynamics is the net flow in zebrafish Kupffer’s vesicle. A previous work presented detailed trajectories of bead movements using DIC and fluorescent imaging, in mouse, rabbit, and medaka (Okada et al., 2005). This work described the rapid fluid flow from right to left crossing over the midline and slow rightward counter flow at the posterior end. In addition, it has been suggested that rightward flow is also necessary to generate leftward fluid flow in the mouse node (Buceta et al., 2005). Thus, our model of fluid flow in zebrafish Kupffer’s vesicle is reasonably comparable to the descriptions of flow within the mouse node, and may actually reflect the flow that occurs in vivo within the space generated by Reichert’s membrane over the mouse node.
Accumulating evidence says that cilia-driven leftward flow is the key to establishing correct organ situs in mouse (Nonaka et al., 2002). Obviously, defects in cilia or lack of fluid flow randomizes L-R organ patterning in mouse and zebrafish (Brody et al., 2000; Chen et al., 1998; Nonaka et al., 1998; Marszalek et al., 1999; Okada et al., 1999; Takeda et al., 1999), suggesting that cilia motility is the source of the flow. Previous studies have shown a left-biased intracellular Ca2+ flux, proposed to be important in L-R asymmetry (McGrath et al., 2003; Okada et al., 2005; Sarmah et al., 2005; Tanaka et al., 2005). Although the leftward flow is obvious in the mouse node, it is still questionable how this leftward flow activates left side specific gene expression. One model proposes that the peripheral monocilia which contain Polycystin-2 are non-motile but bend in response to the leftward fluid flow which allows the calcium ion channel to open (McGrath et al., 2003). However, in our model, peripheral cilia are still motile and “bending” affected by flow would occur at any side.
Recently, it has been proposed that nodal flow in the mouse transports nodal vesicular parcels (NVPs, 0.3-5 um in diameter) could deliver morphogens (Tanaka et al., 2005). This theory suggests NVPs must rupture on the left side to specifically affect these cells (Cartwright et al., 2007). Because the size of the beads we use is around 0.2 um in diameter, it is assumed that NVPs would be bigger than beads but smaller than the bubble we imaged inside zebrafish Kupffer’s vesicle. Thus it must be technically possible to trace some movements of NVPs if they exist in the fluid. However, it was difficult to determine if these vesicles exist in the bead experiments. During observation of the cilia motility without beads, we occasionally observed some small vesicles or particle movements, but not frequently enough to record.
We have proposed an angulated circular motion of net flow, which would produce a dominant leftward flow specifically across the cells in the dorso-anterior region of Kupffer’s vesicle. Even though the rightward flow may not be effective on the posterior-ventral region, the circular motion still suggests that morphogens would be recycled. However, it is possible that this regional leftward flow is fast enough to cause mechanical activation, or break morphogen vesicles at the left side. Overall, our current study on the dynamics of flow in Kupffer’s vesicle provides the foundation for future studies to determine how flow contributes to the asymmetric establishment of Nodal signaling in zebrafish.
The zebrafish and procedures used in this study were in accordance with the guidelines and approval of Princeton University IACUC. Wild type fish with low background of left-right defects used for experiments, including AB, WIK, and Tu, are maintained in our facility per standard methods. Developmental stages were determined from somite numbers and other morphological features of embryos (Kimmel et al., 1995).
Embryos were collected at appropriate stages, dechorionated, and fixed with 1% paraformaldehyde (PFA), 2.5% glutaraldehyde (GA) in phosphate buffered saline (PBS) overnight. The body axes and yolks were separated using tweezers, and fixed at 4°C for an additional overnight. After PBS washing, samples were stored in PBS at 4°C for another overnight to remove any excess GA. Embryos were then fixed with 1% osmium in PBS at 4°C overnight. After washing with aqua destillata, dehydration with a graded series of ethanol solutions was performed at room temperature. Specimens were processed through a critical point freeze-dry procedure. After coating with iridium through ion beam sputtering (VCR Group Inc., IBS/TM200S), both ventral floor and dorsal roof of Kupffer’s vesicle were analyzed by scanning electron microscopy (Phillips, SEM XL30).
Embryos were dechorionated in E2 buffer (0.5 mM KCl, 15.0 mM NaCl, 2.7 mM CaCl2, 1.0 mM NaHCO3, 0.7 mM HEPES, adjusted at pH 6.5) after tailbud stage. From 1-10 somite stages, embryos were embedded into 0.9% low-melting agarose and set under the 60x water immersion objective lens. All images were taken with the dorsal roof of Kupffer’s vesicle facing the 60x water immersion objective lens, defined as “dorsal view”. FluoSpheeres®Fluorescent Microspheres were obtained from Molecular Probes (Eugene, OR, USA) and injected for observation of bead movements. Beads were injected into Kupffer’s vesicle at 4 somite stage when the shape of Kupffer’s vesicle first became clear. DIC live images were recorded by video microscopy using an Andor iXon camera (Andor Technology) with speed of 112-166 frames per second (fps) for cilia motility and 50 fps for beads movements. Different views of dorsal roof and ventral floor of Kupffer’s vesicle were obtained by changing the focusing depth. All images were reconstructed for movies at 8 fps for cilia motility and 30 fps for beads movements using MATLAB.
Based on the previous analysis in medaka (Okada et al., 2005), cilia motility images were taken at the dorsal roof and ventral floor regions where the majority of ciliated cells are located within a relatively flat sheet. Cilia motility was traced using discrete Fourier transform analysis and interpreted into power spectrum, based on the intensity of raw images. Only powers in the range of cilia frequencies were summed and plotted into one image to determine the trajectories of cilia movement using MATLAB program. The source code is available on request. The number of posteriorly pointed cilia was manually counted for each dorsal roof or ventral floor image.
Embryos were collected at appropriate stages and fixed in 4% PFA overnight at 4°C. After gradual dehydration into ethanol, embryos were infiltrated and embedded in JB-4 plastic resin according to manufacturer’s instructions (Electron Microscopy Sciences). 4 μm sections were obtained with a microtome (Leica RM2255, Wetzlar, Germany), followed by Hematoxylin & Eosin staining. Images were taken under DIC microscope (Leica DMRA2, Wetzlar, Germany), and photographed with a JenOptik ProgRes C14 digital camera.
Video 1: Cilia located at dorsal roof of Kupffer’s vesicle showed counterclockwise rotations when viewed from dorsal side at 3-somite stage.
Video 2: Cilia located at ventral floor of Kupffer’s vesicle showed clockwise rotation when viewed from dorsal side at 3-somite stage.
Video 3: Cilia located at lateral edge of Kupffer’s vesicle showed beating when viewed from dorsal side at 3-somite stage.
Video 4: Cilia located at anterior-dorsal side of Kupffer’s vesicle showed conical shape when viewed from dorsal side at 7-somite stage. Posterior side of the embryo is to the bottom of the panel.
Video 5: Cilia located at posterior-ventral side of Kupffer’s vesicle when viewed from dorsal side at 7-somite stage. Posterior side of the embryo is to the bottom of the panel.
Video 6: Cilia located at lateral edge of Kupffer’s vesicle showed beating when viewed from dorsal side at 7-somite stage. Posterior side of the embryo is to the bottom of the panel.
Video 7: Counterclockwise movements of beads closest to the dorsal roof of Kupffer’s vesicle when viewed from dorsal side at 5-somite stage.
Video 8: Clockwise movements of beads closest to the ventral floor of Kupffer’s vesicle when viewed from dorsal side at 5-somite stage.
Video 9: Dominant flow was observed by counterclockwise movements of beads and the bubble motion at the middle level of Kupffer’s vesicle when viewed from dorsal side at 9-somite stage. Posterior side of the embryo is to the right side of the panel.
Table I: Measurement of cilia length in zebrafish Kupffer’s vesicle
We thank Dr. Stephan Y Thiberge and Dr. David Tank for supporting assistance with video microscopy, Dr. Nan Yao, Dr. Jane Woodruff and Ms. Margaret Bisher for technical support for the SEM and histological analysis, and Dr. Joseph Goodhouse for assistance with confocal microscopy. We also thank Dr. Jonathan Eggenschwiler, Dr. Elizabeth Gavis, Dr. Gertrud Schupbach, Dr. David Tank and Burdine lab members for helpful discussion, and Ms. Heather McAllister for maintenance of the fish facility. This work has been supported by Johnson & Johnson, Inc. microscopy award/fellowship to NO and RDB and an award to RDB from the National Institutes of Child Health and Human Development (1R01HD048584). The Princeton imaging facility is supported by an NIGMS Center Grant (P50 GM 071508).
Grant information: NICHD:1R01HD048584; NIGMS:P50 GM 071508; Johnson & Johnson, Inc.