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Developmental Biology
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Evolution of Developmental Control Mechanisms
The formation and positioning of cilia in Ciona intestinalis embryos in relation to thegeneration and evolution of chordate left–right asymmetry
Helen Thompson a, Michael K. Shaw b, Helen R. Dawe c, Sebastian M. Shimeld a,⁎a Department of Zoology, University of Oxford, South Parks Road, Oxford, OX1 3PS, UKb Sir William Dunn School of Pathology, University of Oxford, South Parks Road, Oxford, OX1 3RE, UKc Biosciences, College of Life and Environmental Sciences, University of Exeter, Stocker Road, Exeter, EX4 4QD, UK
In the early mouse embryo monocilia on the ventral node rotate to generate a leftward flow of fluid. This nodalflow is essential for the left-sided expression of nodal and pitx2, and for subsequent asymmetric organ patterning.Equivalent leftfluidflowhas been identified in other vertebrates, including Xenopus and zebrafish, indicating it isan ancient vertebrate mechanism. Asymmetric nodal and Pitx expression have also been identified in severalinvertebrates, including the vertebrates’ nearest relatives, the urochordates. However whether cilia regulatethis asymmetric gene expression remains unknown, and previous studies in urochordates have not identifiedany cilia prior to the larval stage, when asymmetry is already long established. Here we use Scanning and Trans-mission Electron Microscopy and immunofluorescence to investigate cilia in the urochordate Ciona intestinalis.We show that single cilia are transiently present on each ectoderm cell of the late neurula/early tailbud stageembryo, a time point just before onset of asymmetric nodal expression. Mapping the position of eachcilium on these cells shows they are posteriorly positioned, something also described for mouse node cilia. TheC. intestinalis cilia have a 9+0 ring ultrastructure, however we find no evidence of structures associated withmotility such as dynein arms, radial spokes or nexin. Furthermore the 9+0 ring structure becomes disorganisedimmediately after the cilia have exited the cell, indicative of cilia which are not capable of motility. Our resultsindicate that although cilia are present prior to molecular asymmetries, they are not motile and hence cannotbe operating in the same way as the flow-generating cilia of the vertebrate node. We conclude that the ciliamay have a role in the development of C. intestinalis left–right asymmetry but that this would have to be in asensory capacity, perhaps as mechanosensors as hypothesised in two-cilia physical models of vertebrate cilia-driven asymmetry.
Many animals have asymmetric positioning of organs which differ-entiates the left side of the organism from the right. In humans, thefailure to establish correct organ positioning results in heterotaxy(Kosaki and Casey, 1998) which is a symptom of a number of geneticdiseases including primary ciliary dyskinesia (PCD), also known asimmotile cilia syndrome. PCD has a 50% incidence of situs inversus, thecomplete mirror image reversal of organs, as well as defects in respira-tory cilia and sperm motility (Afzelius, 1976; Noone et al., 1999).
Morphological asymmetry in mice is preceded by molecular asym-metry, with the left-sided expression of nodal observed adjacent tothe mouse node (Collignon et al., 1996; Lowe et al., 1996). Left-sidednodal expression is later transferred from the node to the Lateral PlateMesoderm, which is essential for downstreammorphological asymme-tries (Brennan et al., 2002; Kawasumi et al., 2011), and is followed by
. Shimeld).
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asymmetric expression of the nodal antagonists lefty-1 and −2 (Menoet al., 1996) and the transcription factor pitx2 (Ryan et al., 1998).
Scanning Electron Microscopy (SEM) of mouse embryos from thisdevelopmental time shows the presence of cilia, ~5 μm in length and0.3 μm in diameter (Hirokawa et al., 2009), on the mouse ventralnode, with one cilium projection per cell (Sulik et al., 1994). Themicrotubule-based cytoskeleton known as the axoneme can providestrength and motility for cilia, and cilia are generally separated intotwo groups on the basis of axoneme structure; motile (9+2 withdynein arms, radial spokes and nexin) and immotile (9+0 withoutdynein arms). However the cilia at the mouse ventral node do notcategorise to either type, instead having an unusual 9+0 with dyneinarms structure (Fig. 1A) (Hirokawa et al., 2006).
Cilia at the mouse ventral node are motile and beat in a vorticalfashion which generates a net leftward fluid flow across the node(Nonaka et al., 1998). The flow, termed nodal flow, generated bycilia is able to create a net flow as a result of the posterior positionand 40°±10° tilt of each cilium on the cells of the node (Hashimotoet al., 2010; Hirokawa et al., 2006; Nonaka et al., 2005; Okada et al.,2005). Disruption to nodal flow or the positioning of cilia leads to
Fig. 1. Mechanisms and asymmetric gene expression involved in left–right asymmetry across the holozoa, including Ciona intestinalis. A: The ultrastructure of motile cilia (9+2),primary cilia (9+0) and nodal cilia (9+0 with dynein arms). Motile cilia have a 9+2 structure in which the nine doublet microtubules surround a central pair whereas primarycilia do not possess a central pair. Additionally, motile cilia also possess dynein arms, radial spokes and nexin connections which are essential for motility. Nodal cilia have the samestructure as primary cilia but also possess dynein arms which provide the motor to enable the cilia to rotate. B: The Holozoan phylogeny of selected genomes and including thepresence of asymmetric localisation of genes important in the development of the left–right axis (asymmetric localisation through experimentation is depicted with a tick). Thepresence and asymmetric localisation of the genes has not been tested in some species (−) or is absent (cross). Some species which are known to require lateral flow for thegeneration of left–right asymmetry are indicated with a green box and Y. Experimentation has shown that lateral flow is not present on the chick (red jungle fowl) or pig. Comparedto chordates the purple sea urchin has reversed asymmetric expression, as do some molluscs. Relevant references for gene expression are (Bisgrove et al., 1999; Boorman andShimeld, 2002; Campione et al., 1999; Cheng et al., 2000; Duboc et al., 2005; Grande and Patel, 2009; Ishimaru et al., 2000; Levin et al., 1995; Logan et al., 1998; Lowe et al., 1996; Lustiget al., 1996; Meno et al., 1996; Mita and Fujiwara, 2007; Nagai et al., 2010; Okada et al., 2005; Onai et al., 2010; Rebagliati et al., 1998; Ryan et al., 1998; Soroldoni et al., 2007; Yu et al.,2007). C: SEM of a C. intestinalis embryo within a chorion with emanating follicle cells (fc). D: A C. intestinalis larvae. E: A dorsal view of the head of a C. intestinalis larva, showing theasymmetric positioning of the ocellus (oc) and otolith (ot) in the sensory vesicle (sv). Scale bars=100 μm (C); 50 μm (D); 20 μm (E).
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randomised and bilateral expression of normally left-sided expressedgenes, and compromised downstream left–right development. Whilea role for cilia in the development of left–right asymmetry is nowgenerally accepted, the precise mechanism whereby nodal flow setsup the left–right axis remains controversial. Two main hypotheseshave been invoked to explain this: the chemical hypothesis and thephysical hypothesis. The former is based on the idea that a morphogenis transported to the left of the node in nodal vesicular parcels (NVPs)by the nodal flow, hence producing a concentration gradient in thecavity of the ventral node (Nonaka et al., 1998; Okada et al., 2005).The latter assumes that the leftward flow is mechanically detected byimmotile cilia on the periphery of the ventral node, resulting in an intra-cellular Ca2+ cascade (Fliegauf et al., 2007; Hamada, 2008; Hirokawa etal., 2009; McGrath et al., 2003; Shiratori and Hamada, 2006;Vandenberg and Levin, 2010).
Nodal flow is conserved in some other vertebrates, with cilia drivenlateral fluid flow identified in the homologous organiser structuresin rabbit, Xenopus and zebrafish, but has not been identified in
invertebrates. However the asymmetric expression of nodal and pitxhas been found in invertebrate chordates, echinoderms and molluscs,suggesting it is of ancient evolutionary origin (Fig. 1B). Here, we focuson the vertebrates nearest invertebrate relative, the urochordates(Delsuc et al., 2006), to test whether cilia may be involved in the regu-lation of left–right asymmetry outside the vertebrates.
Urochordates, together with cephalochordates and vertebrates,constitute thephylumChordata. At somepoint in their lifecycle all chor-dates share the same distinctive characteristics including a notochord,dorsal neural tube, endostyle and postanal tail. The twomost frequentlystudied urochordate systems are the ascidians Ciona intestinalis andHalocynthia roretzi, both sessile filter feeders as adults but which duringembryogenesis develop a motile tadpole larva with clear similarities tovertebrates. The C. intestinalis larva has left–right asymmetry, withasymmetric positioning of two sensory pigment spots and adjacentregions of the brain, while in the adult there is an asymmetrically foldedgut (Fig. 1C–E) (Boorman and Shimeld, 2002).Nodalhas been describedin H. roretzi (Morokuma et al., 2002), C. intestinalis (Mita and Fujiwara,
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2007) and a third ascidian,Molgula oculata (Osada et al., 2000).Nodal isfirst observed in a pair of lateral blastomeres in C. intestinalis embryosand later in six vegetal blastomeres at the 32 cell stage, where it issymmetrically expressed (Imai et al., 2004; Morokuma et al., 2002).Asymmetric expression is seen by the tail-bud stage of development(Morokuma et al., 2002). The pro-orthologue of Pitx2, Ci-Pitx, is alsoexpressed asymmetrically at the tailbud stage (Boorman and Shimeld,2002). This expression precedes the first morphological signs of left–right asymmetry, as seen in the development of vertebrate left–rightasymmetry. The regulation of this asymmetric gene expression is essen-tially unknown, though one study has implicated H+/K+-ATPasefunction (Shimeld and Levin, 2006). No study has ever reported thepresence of cilia on ascidian embryos prior to the larval stage; henceany role for cilia in early ascidian development remains completelyunknown.
Here, we seek to resolve the apparent inconsistency in conservationof left–right patterning mechanisms, which has molecular mechanismsdeeply conserved but the presence of motile cilia found in some verte-brates, but not others such as chick and pig (Gros et al., 2009), and theiruse in symmetry-breaking experimentally supported only in mice. Weaim to test whether cilia are present on the developing C. intestinalisembryo prior to asymmetric gene expression, and if they are motileand could hence generate a leftward flow. Using SEM and immunofluo-rescence we show cilia are present on every ectoderm cell of C. intesti-nalis embryos through a tightly defined period of development justpreceding asymmetric gene expression. We show that the cilia arepositioned at the posterior of each cell, as they are at the mouse node.However, Transmission Electron Microscopy (TEM) shows the cilia tohave a 9+0 structure without dynein arms and with collapsingaxonemes upon exit from the cell. We hence conclude that, while thetiming and positioning of cilia suggest they may have a role in left–right development, this is not via generation of fluid flow.
Materials and methods
Ciona intestinalis animals and embryos
Adult C. intestinalis were collected from Northney and SparkesMarinas, Portsmouth, UK, and maintained in a closed sea water systemat 16 °C. Oocytes and sperm were dissected and mixed for 15 min inRoom Temperature (RT) filtered sea water (FSW). The sperm wereremoved and the oocytes washed with RT FSW. The fertilised oocyteswere left to develop at 12 °C. Developing embryos were collected andde-chorionated every 30 min as described (Mita-Miyazawa et al.,1985). After de-chorionation the embryos were washed twice in FSWand then immediately fixed for Electron Microscopy (EM). For TEM,embryos were also fixed within their chorion, and proved to have iden-tical cilia structure to those that had been chemically de-chorionated.
Electron microscopy (EM)
Embryos were fixed by addition of 2.5% glutaraldehyde in 150 mMcacodylate buffer (pH7.0) with 1.6% NaCl and 0.01% picric acid to FSWat RT and stored at 4 °C for 1–2 days. After the primary fixation thesamples were washed in cacodylate buffer (0.2 M, pH 7.4) and post-fixed with 1% osmium tetroxide in 100 mM cacodylate buffer (pH7.0) followed by several rinses in cacodylate buffer. The samples forSEM were dehydrated with ethanol to 100%, were critically point-dried then mounted on aluminium SEM stubs using double-sidedcarbon tabs. After sputter-coating with gold, the samples were exam-ined in a JEOL 6390 JSM SEM. Samples for TEM were enbloc stainedwith 0.5% aqueous uranyl acetate for 16 h at 4 °C in the dark. Followingdehydration, the material was embedded in resin (Agar 100 resin). 50–70 nm sections were cut, stained with 0.2% lead citrate and viewed on aFEI Technai 12TEM.
Antibody staining
The primary antibody used was anti-acetylated alpha tubulin cloneC3B9 (Woods et al., 1989). Embryos were fertilised as previously statedand allowed to develop at 18 °C for 7 h until they reached the neurulastage. Embryos were fixed in −20 °C MeOH for a minimum of 2 hbefore they were processed as described (Hudson and Lemaire, 2001).Primary antibody was used at 1:2 (C3B9; (Woods et al., 1989)) in Tx-PBS (0.1%) and incubated for 1 h at RT. Embryos were washed for atleast 10 min 3 times in Tx-PBS (0.1%) at RT, and then the secondaryantibody (Alexa-fluor 488-conjugated goat anti-mouse (MolecularProbes)) was used at 1:500 in Tx-PBS (0.1%) before at least 10 min 3times in Tx-PBS (0.1%). Finally, DAPI was added at 1:1000 in Tx-PBS(0.1%), washed for at least 10 min 3 times in Tx-PBS (0.1%) andmounted in Vectashield anti-fade (Vectashield, Vector LaboratoriesLtd., Peterborough, UK). The embryoswere then viewedon a Zeiss Axio-scope and the images processed inMetamorph (Universal Imaging) andfigures assembled in Adobe Photoshop CS4.
Cilia positional measurements
Embryos were fixed and imaged on a JEOL JSM 6390 SEM as de-scribed above. Photographs were taken of each embryo and eachembryo was aligned along its A–P axis, then all the cells parallel to thesurface of the specimen were traced in Adobe Illustrator and the posi-tion of the base of the cilium recorded with a blue circle. The centre ofeach cell was calculated and the length of the A–P axis of the cellthrough the centre recorded. The length through the centre of the cellat which the cilium was situated along the A–P axis was measuredand converted into a percentage of the total cell A–P axis. The centreof the each cell was located and the distance from the anterior to thecentre of the cell marked as 0o. The angle from this point to the ciliumwas measured and recorded. Statistics used ANOVA and analyses wereconducted in R 2.12.2 (R Foundation for Statistical Computing, Vienna,Austria).
Results
Absence of cilia on Ciona intestinalis embryos prior to the early tail budstage
We first used SEM to examine the external surface of C. intestinalisembryos at different stages of development for cilia. Early 8–16 cellembryos (Fig. 2A) are covered in microvilli of ~1.0 μm in length(Fig. 2B, arrows), which emanate from a rough, crinkled cell surface.Much longer, thin projections, up to 8.5 μm in length, were also presentbetween some cells of the developing embryo (Fig. 2C). Later 8–16 cellembryos (Figs. 2D, E) also had crinkled surface texture covered inmicrovilli and projections between cells up to 12 μm long (Fig. 2F). 32cell, 44 cell and 110 cell embryos (Fig. 2G–I) had smoother cell surfacesand an absence of the microvilli observed at earlier stages. Projectionssimilar to those seen in earlier embryos were observed between cellsof the 32 cell embryo (arrowhead) (Fig. 2G) but not in any of theother embryos. Some remnants of chorion remain on the later embryos(Fig. 2H, I). No cilia were observed on any embryo at any of thesestages. The gastrula (Fig. 3A) and early to mid neurula (Fig. 3B, C)stage embryos have a smooth surface texture and no evidence ofmicro-tubule structures. Due to the de-chorionation process some residualpieces of chorion and test cell remain on the cells but should not beconfused with microvilli (Fig. 3A). Cells at these stages also lack cilia.
Transient cilia appear on ectodermal cells at the tail bud stage
At the early tailbud stage (Fig. 3D inset) the posterior region of theembryo had begun to lengthen and bend ventrally, while the anteriorregion of the embryo had expanded to create a visible head region. At
Fig. 2. The surface texture and microvilli structures of early Ciona intestinalis cleavage stage embryos. A: SEM image of an embryo at the early 8–16 cell stage with the chorion removed.The embryo is under-going cleavage from an 8 to 16 cell embryo. B: Higher magnification of the cell surface of the developing embryo. The surface appears crinkled and microvilli arepresent (arrows). C: Higher magnification of the area between individual cells. Thin projections are present between some cells (block arrows). D: SEM image of an embryo at the late8–16 cell stage with the chorion removed. E: A closer image of a dividing cell with furrows on the embryo. F: The thin projections (block arrows) observed in earlier stage embryoswere also present on the later stage embryo. G, H, I: SEM images of 32 cell, 44 cell and 110 cell embryos with the chorion removed. Small, thin projections can be seen between somecells of the embryo (block arrows). Some chorion (ch) debris remains on the cells of the 44 cell and 110 cell embryos. Scale bars=20 μm (A, D, G, H, I): 5 μm (D, E, F): 2 μm (B, C).
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this stage all the cells of the ectoderm possessed a single ciliumprojecting from their apical surface (arrows) (embryos n=54). Thecilia were on average 0.43 μm (SD=0.02; n=48) in length (Fig. 3D).Small cilia might be confused with other projections, hence to confirmtheir identity we used staining with anti-acetylated α-tubulin(which labels microtubules in cilia), and DAPI (which stains cell nuclei)(Fig. 3E, F). Focusing through the embryo showed that anti-acetylatedα-tubulin staining was only found around the outside of the embryothat is on the ectoderm. The ectodermal projections thought to be ciliawere stained with anti-acetylated α-tubulin, and measured approxi-mately 0.5 μm in length (embryos n=9).
Cilia persisted from the early to mid tailbud stage but shortly afterthis developmental time point the cilia disappeared. Cilia were absenton all the cells of the late tailbud stage embryos (embryos n=21)(Fig. 3G–I). There was also no evidence of projections between thecells of the embryo. By the late tailbud stage the neuropore hadcompletely closed and the cell shape changed from round to rectangular(Fig. 3I). Cilia were also not found at this stage.
C. intestinalis cilia are polarised relative to the AP axis
At the mouse node, cilia are polarised relative to the AP axis andthis orientation, driven by planar cell polarity signalling, is essentialfor the generation of lateral fluid flow and hence left–right pattern.
In C. intestinalis SEM and immunofluorescence data show that surfacecilia appeared briefly at the early to mid tailbud stage. In comparisonto previous studies, surface cilia just preceded the first signs of molec-ular asymmetry, the left-sided expression of nodal and Pitx (Boormanand Shimeld, 2002; Mita and Fujiwara, 2007; Morokuma et al., 2002).This prompted us to hypothesise that the cilia might be part of a con-served mechanism of left–right patterning, a prediction of which isthat they should be similarly polarised relative to the AP axis inorder to generate lateral fluid flow.
To quantify the position of the cilia we recorded and measured thedistance along each cell's A–P axis and the angle from the centre ofthe cell (n=441) each corresponding cilium was positioned on eachtailbud embryo (n=54) (Fig. 4A–C). The cilia location distance alongthe A–P axis was converted to a percentage of the total cell A–P lengthto incorporate the difference in individual cell size. This processconfirmed that cilia are positioned predominantly in the posterior halfof the cell. In 432 (98%) of the cells the cilium projected posterior tothe centre-point (Fig. 4D) and 368 (83%) of cilia were positioned at anangle of between 136 and 225° from the centre-point (Fig. 4E). Nocilia emanated from the anterior 40% of the cell (Fig. 4D) or betweenthe angles of 0–45° and 316–360° (Fig. 4E).
We also examined whether the cilia were positioned differentlywithin the posterior half of the cell at different embryo locations. Theembryo was divided into nine regions; the Face of the embryo (right,
Fig. 3. The presence of cilia on the tailbud stage Ciona intestinalis embryo through SEM, immunofluorescence and DAPI staining. A, B, C: SEM images of C. intestinalis embryos from thegastrula stage to the mid-neurula stage with the chorion removed. The blastopore (bp) and neuropore (np) is indicated on the gastrula and right lateral (B) and dorsal view (C) of themid-neurula stage embryos. Some chorion (ch) debris remains on the cells of the gastrula embryo. D: SEM image of a left lateral and ventral side of an early tailbud embryowith chorionremoved (inset). A higher magnification image of the central ventral side (red box) of the embryo in D inset showing a single cilium projection from each cell (arrows). E: Immunoflu-orescence and DAPI staining of a tailbud C. intestinalis embryo. F: Highermagnification image of the ventral side of the head region of the tailbud embryo in E. Staining shows primary cilia(white arrows), visualised using anti-acetylated α-tubulin (green). Cell nuclei are visualised with DAPI (blue). G, H, I: SEM image of tailbud embryos with the chorion removed whichdemonstrate no evidence of cilia. Scale bars=20 μm (A, B, C, D (inset) E, G, H, I): 10 μm (D); 5 μm (F).
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ventral and left) and Position along the A–P axis (1, 2 and 3) and thecilia position and angle on each cell recorded. The different Positionand Face percentages and angles were merged and statistically testedby using an ANOVA test incorporating the subtle differences in embryostage as a random effect. The location of the cell on the embryo had noeffect on the angle of the cilium from the centre of the cell (F4,426,p>0.05). Primarily the location of the cell does not affect the positionof the cilium along the A–P axis. However there was a significant differ-ence in the position of cilia on cells located on the left side of the embryoin the centre of the ectoderm, when compared to cilia on cells inother locations (F4,426, pb0.01). Thus, we conclude the cilia of theC. intestinalis early tailbud stage embryo are positioned at the posteriorof each cell, similar to those of themouse ventral node. This positioningis consistent with the generation of lateral fluid flow, as seen atthe mouse ventral node (Antic et al., 2010; Hashimoto et al., 2010;Nonaka et al., 2005).
C. intestinalis ectoderm cilia have a non-motile structure
Cilia are generally grouped into two types;motile and immotile, andthis can be determined via analysis of their ultrastructure by TEM. The
data described above support the hypothesis that C. intestinalis maybe using polarised ectoderm cilia to regulate left–right asymmetry. Forthis to be mechanistically conserved with vertebrates, the cilia wouldneed to be generating a lateral asymmetric force and hence be motile.To test this we therefore used TEM to assess the ultrastructure ofC. intestinalis cilia to ask if the structure was consistent with a possiblerole in motility. Serial sections of embryos (n=4) were cut to identifythe ultrastructure of cilia (n=76) along their entire length. The basalbody of the cilium was found within the cytoplasm of the cell and hasa ring nine triplet microtubule structure (Fig. 5C14–16), with transi-tional fibres extending into the cell cytoplasm (double headed arrows).A nine doublet microtubule ring with connections between the micro-tubule doublets and the ciliary membrane, indicative of the transitionzone, was observed for 140 nm as the cilia exited the cell (Fig. 5C13,white arrows). Immediately after the cilium had exited the cell the ax-oneme had a nine doublet ring structure (Fig. 5A12), but within 70 nmof this the nine doublet microtubules were no longer in an organisedring structure (Fig. 5A1–11). Towards the distal tip of a cilium thedoublet microtubules became more disorganised and some doubletsbecame singlet microtubules (Fig. 5A1–11). The nine doublet microtu-bules first showed evidence of a loss of doublet structure ~210 nm
Fig. 4. Polarisation of cilia on the developing Ciona intestinalis embryo to the posterior of each cell. A: A High magnification SEM image of an early tailbud embryo showing the pres-ence of one cilium per cell on the outer ectoderm. Low magnification image of the embryo in the inset shows the anterior (a) and posterior (p). The red boxed area is shown at ahigher magnification in A. B: A corresponding cartoon of A with an outline of each cell with the location of each cilium anchor (red) and direction of each cilium projection (blue)(Maisonneuve et al., 2009). C: A diagram to show the method used to determine the percentage along the embryo and cell A–P a cilium was positioned. D: A graph to demonstratethe location of each cilium projection as a percentage along the A–P axis of each individual cell (n=441) on 54 early tailbud stage embryos. In 432 of cells (98%) the cilium projectsposterior to the centrepoint (c, yellow line) of the cell. No cilia project from the anterior 40% of any cell. All the cilia (represented by a red circle) aligned along the centre A–P axis ofthe cell are positioned posterior to the centre point (yellow c). E: Each embryo was separated into 45º segments. A radar chart to show the frequency of cilia (n=441) at eachdegree from the centre of the cell. 98.9% of cilia were positioned between 90 and 270° (n=436) from the centre. F: On the left is a cartoon representation to show the segregationof areas on an early tailbud embryo by position (anterior, central, posterior; 1, 2, 3 respectively) and face (right, left, ventral; R, L, V respectively) to generate a comparison of celllocation to the position of the cilia. The right panel shows overlaid plots of cell outline and cilia position colour coded by cells from these regions. Scale bars=20 μm (inset A): 5 μm(A, B); 2 μm (C).
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after the cilium has exited the cell (n=9) (Fig. 5A4, 5). On all the ciliarecorded (n=76) the axoneme accessories required for motility werenot observed, including no evidence of a central pair, radial spokes ornexin connections between doublets.
We hence show that these cilia have a structure most similar tovertebrate non-motile primary cilia (Allen, 1965; Currie andWheatley, 1966; Dahl, 1963; Wheatley, 1967). They are not similarto either motile cilia, or to the unusual mouse nodal cilia, and defini-tively lack the structural characteristics of motility. We conclude thatthey are not motile.
Discussion
While there is a general consensus that the role of nodal and pitx inspecifying left–right asymmetry is conserved at least through thedeuterostomes, and possibly in some protostomes too (Fig. 1), there isconsiderable doubt as to the antiquity of the symmetry-breakingmech-anism ormechanisms that lie upstreamof this. In some vertebrates ciliahave been implicated in this process, a hypothesis supported by bothexperimental data and their helical structure. However cilia have notbeen implicated in regulating left–right asymmetry outside the verte-brates, and indeed previous studies of the vertebrates’ nearest living
relatives, the urochordates, did not find cilia at the right time andplace to be involved in this process (Satoh and Deno, 1984), with thefirst cilia identified well after the onset of molecular and morphologicalasymmetry.
To re-address this question, SEM was conducted on early (8 cell tolate tailbud) C. intestinalis embryos to search for surface structuresthat could be cilia. Early cleavage embryos did not have cilia, thoughwe did note two contrasting cell surface textures, smooth andcrinkled. This phenomenon has been attributed to the position ofthese cells within the cell cycle, and has previously been described inH. roretzi, with microvilli present just before cell division and cytokine-sis but absent after the cell divides (Satoh and Deno, 1984). Anothercommon feature in cleavage embryos were long, thin projections,from2 to 12 μmin length, that extend between cells. Similar projectionshave been previously described in H. roretzi, where they were termed‘cytoplasmic processes’ (Satoh and Deno, 1984). This is the first timethey have been shown to exist in C. intestinalis and, since C. intestinalisand H. roretzi belong to widely divergent groups of urochordates,this suggests this may be a general character of the development ofascidian urochordates. Depending on their structure and compositionit is possible they have a role in communication between cells in earlydevelopment.
Fig. 5. The ultrastructure of Ciona intestinalis early tailbud stage cilia. A: The ultrastructure of a cilium from a early tailbud stage embryo from the tip (1) to immediately prior to thecilium entering the cell (12). Microtubule doublets are not positioned in a ring structure but becomes more disorganised as the sections move towards the distal tip of the cilium.Numbers indicate adjacent serial sections along the C. intestinalis neurula stage embryo. B: Mean measurements from the 75 cilia analysed from the early tailbud stage C. intestinalisembryo are shown in a schematic which demonstrates the disorganisation of the nine doublet microtubules towards the distal tip of the cilium. The transition zone and basal bodysections are from a different cilium from the same embryo and illustrate the ultrastructure of the proximal region of the cilium. C: The ultrastructure of a cilium from the transitionzone (13) to the basal body (16). The basal body has the standard triplet microtubule organisation with transition fibres (double arrows) emanating from the triplets. The transitionzone has nine doublet microtubules which are attached to the membrane through projections (white arrow). Scale bars=200 nm (3–16); 100 nm (1–2).
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Cilia on Ciona intestinalis embryos
Cilia were not observed on C. intestinalis embryos prior to the lateneurula stage. However, as the embryo transitioned through mid tolate neurula, a single cilium was seen to project from some cells. Bythe early tailbud stage, every external cell had a single cilium. Previ-ous SEM studies of ascidian embryos have not identified such ciliadespite relevant stages being examined (Nicol and Meinertzhagen,1988; Satoh, 1978; Satoh and Deno, 1984). It could be that differentspecies differ in this respect; however we suspect that our use offixation specifically for the preservation of fragile microtubule struc-tures has enabled these cilia to be identified and that the presenceof cilia at these stages is a hitherto unappreciated feature of ascidiandevelopment.
Timing and position of cilia on Ciona intestinalis embryos
For cilia to be involved in regulating C. intestinalis left right asymme-try in the same way as in the mouse, they would have to precede theonset of asymmetric nodal expression. The timing of the emergence ofcilia on the C. intestinalis embryo is consistent with such a role. Ciliaare first present at the late neurula/early tailbud stage of developmentand persist for approximately 1 h until the mid-tailbud stage. Duringthis same window asymmetric Ci-Nodal and Ci-Pitx expression werefirst observed (Boorman and Shimeld, 2002; Mita and Fujiwara, 2007;Morokuma et al., 2002). Hence the cilia could regulate asymmetricgene expression. Posterior positioning of cilia is a feature associatedwith nodal cilia and the establishment of left–right asymmetry and isrequired for the rotational movement of the cilia to generate a leftward
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flow (Hashimoto et al., 2010; Hirokawa et al., 2006; Nonaka et al., 2005;Okada et al., 2005). It is a conserved characteristic in the homologousorganiser structures of the rabbit, zebrafish and Xenopus (Essner et al.,2005; Feistel and Blum, 2006; Schweickert et al., 2007). C. intestinalisectoderm cilia are similarly polarised.
The location of the C. intestinalis cilia, on all ectoderm cells, is topo-logically different from those on the mouse node or equivalent orga-niser structures in other vertebrates. However we note that currentdata suggests C. intestinalis (and other ascidians) have lost the orga-niser, instead relying on segregation of maternal determinants andshort range inductive signalling to establish the body plan (Lemaire,2011; Yu et al., 2007). A corollary of ascidians losing the organiserwould be a need to change the location of asymmetry generation. Inthis context we also note that the tissue in which asymmetric nodalexpression is seen is different between vertebrates (adjacent to theorganiser, then left lateral plate mesoderm) and ascidians (leftectoderm), but matches the respective locations of the cilia. It ispossible therefore that the ectoderm has taken over this role of theorganiser in ascidians. We also found that there was a small butsignificant difference in cilia position on the left and right sides of theembryo. While this is intriguing, it does not mean cilia are regulatingasymmetry, as it is possible both cilia position and gene expressionare regulated by an additional, unidentified signal.
Ciona intestinalis cilia structure and motility
To address the potential motility of C. intestinalis cilia, their structurewas analysed by TEM and compared to that of other cilia, includingthose from the mouse ventral node and surface epithelia. The majorityof cilia had a 9+0 axoneme structure immediately after the cilia haveexited the cell however this rapidly disintegrated, with some doubletsbecoming singlets. There was also no evidence that additional struc-tures, such as the dynein arms or radial spokes essential for motility,were present on the cilia of the neurula/tailbud C. intestinalis embryos.
Although cilia have not been observed on early stages of C. intesti-nalis development prior to these experiments, cilia are present on thelarvae in numerous tissues (Crowther and Whittaker, 1994; Dilly,1969; Katz, 1983; Manni et al., 2005; Torrence and Cloney, 1982)and a comprehensive TEM analysis of the structure of these ciliawas recently conducted (Konno et al., 2010). The cilia structuresobserved were grouped into six different categories; (1) 9+2with dynein arms and radial spokes, (2) 9+2 without dynein arms,(3) 9+2 with electron dense bridges to the cilia membrane, (4) 9+0with an electron dense area in the centre of the axoneme, (5) 9+0without dynein arms or radial spokes and (6) short axoneme cilia-likeprotrusions. The only type of cilia we observed on the late neurula/early tailbud stage embryos were similar in structure to group 5on the larvae. However the role of these cilia on the larval stageembryos has not been elucidated so cannot provide clues as to therole of neurula/tailbud stage cilia.
The disorganised 9+0 structure is a phenomenon that has beendescribed repeatedly over the last 50 years but is still not well under-stood (Allen, 1965; Dahl, 1963). These studies reported numerouscilia, in the rat cerebral cortex and human inner retinal neurons andretinal pigment epithelium, which have a distinct 8 peripheral and 1central doublet ultrastructure (8+1), with a peripheral doublet of a9+0 cilium displaced to the centre. This ultrastructure arrangementis never found as a basal body but begins approximately 0.5 μm distalto the cell surface. The apparent collapse of the axoneme could occurbecause there is an absence of connections between doublets whichmaintain the 9+0 ring structure (Dahl, 1963). More recently, theincursion of one or more microtubule doublets into the axonemecore was described in mammalian epithelial cell lines and the amas-tigote Leishmania mexicana, the parasite responsible for Leishmaniasis(Gluenz et al., 2010). The absence of the central pair, radial spokesand nexin as well as the disorganised axoneme architecture has led
to the suggestion that this structure could facilitate a sensory role,through increased bending capacity or transportation of signallingmolecules (Gluenz et al., 2010).
Potential roles of Ciona intestinalis cilia as mechanosensors or sites ofsignalling
If the C. intestinalis cilia are not motile, what other functions couldthey perform? In other organisms non-motile cilia play numeroussensory roles, for example in mechanosensation and chemosensationand indeed some models of mouse nodal flow function invokemechanosensory cilia generating asymmetry by deforming in theflow generated by other cilia (McGrath et al., 2003). Although, as dis-cussed above, it has been hypothesised that cilia with this conserveddisorganised structure could have a sensory role, direct evidence forthis is lacking and we cannot either conclude or exclude that theC. intestinalis cilia are mechanosensory.
Vertebrate primary cilia also have a role in intercellular signallingvia the Hedgehog (Hh) and Wingless (Wnt) signalling pathways.Components of the Hh signalling pathway localise to the cilia(Corbit et al., 2005; Han et al., 2008; Haycraft et al., 2005; Liu et al.,2005; May et al., 2005; Ocbina and Anderson, 2008; Rohatgi et al.,2007) and primary cilia have been implicated in Wnt signalling al-though the mechanistic link between the two is not known (Berbariet al., 2009; Veland et al., 2009). In C. intestinalis embryos, two hedge-hog genes have been described, Ci-hh1 and Ci-hh2 (Takatori et al.,2002). Only Ci-hh2 is expressed at the correct developmental timepoint for a role in signalling via ectoderm cilia, but is only expressedin a row of internal cells in the tail region running along theanterior-posterior axis (Takatori et al., 2002). It is unlikely, therefore,that these cilia play a role in the receipt of Hh signalling. Wnt signal-ling component genes are also not expressed at the correct time/place(Ikuta et al., 2010). We therefore consider it unlikely that Hh andWntsignalling are operating in the C. intestinalis cilia at this stage ofdevelopment.
Finally, one study has suggested that H+/K+-ATPase activity,which is known to regulate vertebrate asymmetry, also regulatesasymmetry in C. intestinalis (Shimeld and Levin, 2006). The timingof H+/K+-ATPase activity in C. intestinalis matches the timing of theappearance of cilia. H+/K+-ATPase activity has been implicated ex-perimentally in asymmetry in several other species, including Xeno-pus, chick and sea urchin (Aw et al., 2008; Duboc et al., 2005;Hibino et al., 2006; Levin et al., 2002; Raya et al., 2004), although itsprecise role is unknown, as is its potential interface with cilia-drivenlateral flow. Likewise we do not know whether the two are function-ally related in C. intestinalis, though the coincidence in timing of ciliaand H+/K+-ATPase activity raises the possibility that both are part ofa conserved mechanism.
Conclusions: The evolution of ciliary mechanisms in theestablishment of left–right asymmetry
Not all vertebrates use lateral flow driven by cilia to regulateasymmetry, however its discovery in mammals, Xenopus and zebra-fish suggests it is ancestral for at least the jawed vertebrates. Inthese vertebrates, cilia are positioned close to or on the organiser.However in C. intestinalis, which lacks a true organiser (Lemaire,2011), cilia are positioned universally over the embryo. Althoughthe cilia are not motile, they could be linked with other mechanismsin the establishment of left–right asymmetry: for example, there iscurrently debate as to how lateral flow is converted to downstreammolecular and morphological asymmetry, with the ‘physical hypoth-esis’ requiring sensory cilia which are able to bend to detect the later-al flow as mechanosensors (McGrath et al., 2003). The disorganisedcilia structure in C. intestinalis is compatible with this (Gluenz et al.,2010). Our data raise the possibility that both cilia and H+/K+-
222 H. Thompson et al. / Developmental Biology 364 (2012) 214–223
ATPase activity may be involved in regulating asymmetry in C. intes-tinalis, implying both mechanisms predate the evolution of the verte-brates. Study of these mechanisms in amphioxus is needed todetermine if this is primitively shared by all chordates.
Acknowledgements
We thankMichael Bonsall for assistancewith the statistical analysesand the BBSRC for financial support.
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