Cilia-driven fluid flow in the zebrafish pronephros, brain and Kupffer's vesicle is required for normal organogenesis

Abstract

Cilia, as motile and sensory organelles, have been implicated in normal development, as well as diseases including cystic kidney disease, hydrocephalus and situs inversus. In kidney epithelia, cilia are proposed to be non-motile sensory organelles, while in the mouse node, two cilia populations, motile and non-motile have been proposed to regulate situs. We show that cilia in the zebrafish larval kidney, the spinal cord and Kupffer's vesicle are motile, suggesting that fluid flow is a common feature of each of these organs. Disruption of cilia structure or motility resulted in pronephric cyst formation, hydrocephalus and left-right asymmetry defects. The data show that loss of fluid flow leads to fluid accumulation, which can account for organ distension pathologies in the kidney and brain. In Kupffer's vesicle, loss of flow is associated with loss of left-right patterning, indicating that the 'nodal flow' mechanism of generating situs is conserved in non-mammalian vertebrates.

Full text

1907
Introduction
Cilia and flagella have well-established roles as motile and
sensory organelles in various species; however, their function
in vertebrate organogenesis is only now coming to light. The
identification of mutations in ciliogenic genes as the
underlying cause of several mutant phenotypes and human
pathologies, including polycystic kidney disease, left-right
asymmetry defects and retinal degeneration, has implicated
cilia function in the normal development of several organs
(Ibanez-Tallon et al., 2003).
The establishment of left-right asymmetry is the earliest
embryonic process associated with cilia function. To generate
proper organ laterality, mechanisms must exist that translate
existing anterior-posterior polarity into signals that break the
bilateral symmetry of the gastrulating embryo. The asymmetric
expression of genes such as nodal and southpaw (Bisgrove et
al., 2003; Hamada et al., 2002) and the position of organs such
as the heart on the left side of the trunk reflect the outcome of
early left-right signaling. In humans, abnormalities in organ
situs in the primary ciliary dyskinesia (PCD) syndrome
(Afzelius, 1985) are due to mutations in several different
axonemal dyneins affecting ciliary motility (Ibanez-Tallon et
al., 2003). The inversus viscerum (iv) mouse, which is mutant
in the left-right dynein gene (lrd; Dnahc11 – Mouse Genome
Informatics) encoding an axonemal dynein heavy chain present
in node cilia (Supp et al., 1997), displays randomization of
early gene expression and later organ laterality (Layton, 1976;
Lowe et al., 1996). Cilia paralysis and loss of fluid flow in this
and other mutants (Brody et al., 2000; Chen et al., 1998;
Marszalek et al., 1999; Nonaka et al., 1998) suggested that
nodal fluid flow was the key factor in establishing organ situs.
Artificial reversal of nodal flow has been shown to randomize
the left-right axis, providing further support for the ‘nodal
flow’ hypothesis (Nonaka et al., 2002). McGrath and co-
workers have proposed that two types of cilia exist in the node:
motile lrd-expressing cilia and non-motile polycystin2-
expressing sensory cilia (McGrath et al., 2003). Nodal flow
could generate a morphogen gradient regulating situs, or,
alternatively, mechanosensory ion fluxes mediated by
polycystin2 may be the signal that initiates left-sided gene
expression. Whatever the final signal may be, the
demonstration that dynein-expressing node monocilia exist in
a range of vertebrate embryos (Brummett and Dumont, 1978;
Essner et al., 2002) indicates that cilia-driven fluid flow may
be part of a general mechanism for establishing left-right
asymmetry. Currently, however, ‘nodal flow’ has been directly
demonstrated only in mouse embryos, and it is unclear whether
or not ciliary motion and fluid flow is relevant to other
vertebrates.
Cilia dysfunction has been implicated in polycystic kidney
disease, based on the findings that disruption of ciliogenic and
cilia-associated genes leads to cyst formation in the kidney
(Igarashi and Somlo, 2002; Nauli and Zhou, 2004). The two
proteins associated with human autosomal dominant PKD,
Polycystin-1 and Polycystin-2, have been detected in the renal
Cilia, as motile and sensory organelles, have been
implicated in normal development, as well as diseases
including cystic kidney disease, hydrocephalus and situs
inversus. In kidney epithelia, cilia are proposed to be non-
motile sensory organelles, while in the mouse node, two
cilia populations, motile and non-motile have been
proposed to regulate situs. We show that cilia in the
zebrafish larval kidney, the spinal cord and Kupffer’s
vesicle are motile, suggesting that fluid flow is a common
feature of each of these organs. Disruption of cilia structure
or motility resulted in pronephric cyst formation,
hydrocephalus and left-right asymmetry defects. The data
show that loss of fluid flow leads to fluid accumulation,
which can account for organ distension pathologies in the
kidney and brain. In Kupffer’s vesicle, loss of flow is
associated with loss of left-right patterning, indicating that
the ‘nodal flow’ mechanism of generating situs is conserved
in non-mammalian vertebrates.
Key words: Cilia, Pronephros, Kupffer’s vesicle, Ependymal cell,
Spinal canal, Kidney cyst, Hydrocephalus, Left-right asymmetry
Summary
Cilia-driven fluid flow in the zebrafish pronephros, brain and
Kupffer’s vesicle is required for normal organogenesis
Albrecht G. Kramer-Zucker
1
, Felix Olale
2
, Courtney J. Haycraft
3
, Bradley K. Yoder
3
, Alexander F. Schier
2
and
Iain A. Drummond
1,
*
1
Renal Unit, Massachusetts General Hospital, 149 13th Street, Charlestown, MA 02129, USA
2
Developmental Genetics Program, Skirball Institute of Biomolecular Medicine, New York University School of Medicine, New York,
NY 10016, USA
3
Department of Cell Biology, University of Alabama at Birmingham Medical Center, Birmingham, AL 35294, USA
*Author for correspondence (e-mail: idrummon@receptor.mgh.harvard.edu)
Accepted 4 February 2005
Development 132, 1907-1921
Published by The Company of Biologists 2005
doi:10.1242/dev.01772
Research article
Development

1908
cilium (Pazour et al., 2002; Yoder et al., 2002) as has the
product of the inversin gene, which is mutant in the human
kidney cystic condition nephronophthisis type 2 (Otto et al.,
2003). Cystin, the protein encoded by the mutant gene in the
cpk cystic mouse, is also localized to apical cilia (Hou et al.,
2002; Yoder et al., 2002). In zebrafish, the results of a large-
scale retroviral insertional mutagenesis screen revealed that
several genes associated with cilia assembly were mutated in
fish that developed pronephric cysts (Sun et al., 2004). In
mammalian epithelial cultured cells, cilia are proposed to be
non-motile mechanosensors that initiate signals controlling
tubular epithelial cell proliferation or homeostasis (Nauli et al.,
2003; Pazour and Witman, 2003). Non-motile cilia with
sensory functions have been described in Caenorhabditis
elegans neurons and several mutant strains with altered sensory
behavior have been identified (Perkins et al., 1986). However,
whether kidney cilia are always immotile, or whether they
might play an additional role in kidney tubule fluid movement,
remains an unresolved question.
Our insight into cilia assembly has been significantly
advanced by the discovery of the cellular machinery
responsible for moving particles along the microtubule
scaffold of cilia or flagella, called intraflagellar transport (IFT)
(Rosenbaum and Witman, 2002). The pleiotropic phenotype
observed in animals carrying mutations in IFT genes has
confirmed the importance of cilia in organogenesis and tissue
physiology. A hypomorphic mutation in the mouse polaris
gene (Tg737/IFT88) results in cystic kidney disease,
pancreatic and bile duct hyperplasia, hydrocephalus, and
skeletal patterning defects (Cano et al., 2004; Moyer et al.,
1994; Richards et al., 1996; Yoder et al., 1995). In zebrafish,
the oval mutant is a stop codon in the polaris/IFT88 homolog;
these fish show widespread neurosensory cell death
(Tsujikawa and Malicki, 2004). Three different zebrafish IFT
proteins associated with cystic kidneys were also identified in
a large-scale insertional mutagenesis screen (Sun et al., 2004).
Despite the implication that cilia defects are associated with
mutant phenotypes, the mechanism by which ciliary
malfunction may lead to the various organ pathologies
remains unclear.
To better understand the developmental roles of cilia in
organogenesis, we examined cilia in the pronephric kidney,
the spinal cord and Kupffer’s vesicle (KV, the equivalent of
the mouse embryonic node) of zebrafish larvae and assessed
the consequence that loss of cilia has on the formation and
function of these organs. We found that cilia in all three of
these structures are motile, suggesting that cilia function to
drive fluid flow. Indeed, we show by using injected
fluorescent tracers that this is the case. Disruption of cilia
function in IFT morphant embryos resulted in loss of fluid
flow and subsequent development of kidney cysts,
hydrocephalus and laterality defects. The association between
defects in fluid flow and organ pathology when cilia
biogenesis was perturbed suggests that a common
mechanism, namely loss of fluid flow, leads to fluid backup
and subsequent organ distension, with formation of cysts in
the kidney and hydrocephalus in the brain. Our data also
demonstrate that fluid flow is a conserved feature of
gastrulation-stage midline structures that regulate left-right
asymmetry and, further, that disruption of this flow in
zebrafish also causes abnormalities in situs.
Materials and methods
Zebrafish lines
Wild-type TL or TÜAB zebrafish were maintained and raised as
described (Westerfield, 1995). Dechorionated embryos were kept at
28.5°C in E3 solution with or without 0.003% 1-Phenyl-2-thiourea
(PTU, Sigma) to suppress pigmentation and staged according to
somite number (som) or hours post-fertilization (hpf) (Westerfield,
1995).
Cloning of
polaris
,
hippi
and a fragment of pronephric
axonemal
dynein heavy chain 9
Zebrafish polaris was cloned by RT-PCR based on sequence predicted
from Sanger Center zebrafish genomic DNA sequence (Sanger
Institute). RT-PCR products were subcloned in pCRII TOPO
(Invitrogen) and sequenced. Zebrafish hippi sequence was derived
partially by tblastn searches (Sanger Institute) and used for 5′ and 3′
RACE reactions (Invitrogen). Finally, the coding sequence was
obtained by reverse transcription and nested PCR of wild-type total
RNA (outer primer-set: forward CCCTTTGCGAGTAAAGAGTGT-
TAAATGTGA, reverse CATCATCTGCTGCAAACTAGCCCTCT,
nested primer-set forward CGGGATCCGCCACCATGGCGGAG-
GAGGAAGAG reverse CGGAATTCCGGCGGTGAGTGTGT-
GTTTCAATA) and subcloned into the expression vector pCS2+. The
hippi gene maps to linkage group 2. The nested PCR for murine hippi
was performed based on known sequence in GenBank (NM_028680)
using total RNA from mouse brain (kind gift from Dr Ruth Luthi-
Carter, Neurology, MGH) and the following primer (outer primer-set:
forward GGCGCTGGGGGTCTGAGCA, reverse AAATTGT-
GTTTGGAAATCAATGCAACA, nested primer-set forward CG-
GAATTCGCCACCATGGCCGCCGCGGCCGCG, reverse GCTC-
TAGAGAAGCATGGAAGCCCACGTGTTTA). The amplicon was
subcloned into the expression vector pCS2+.
For isolation of pronephros specific axonemal dynein heavy
chains, 72 hpf. zebrafish embryos were incubated in 10 mmol/l
DTT in egg water for 1 hour at room temperature and then washed
three times with egg water. They were then incubated at 28.5°C
in 5 mg/ml collagenase II in Hank’s saline with calcium
(Worthington) for 4 hours. The larvae were then put in Hank’s saline
and triturated gently five times with a 1000 µl pipette tip, so that the
disintegrated, approximately 20 pronephric duct fragments were
collected by visual identification. Total RNA from the collected
tissue was used for reverse transcription and nested PCR. The
following degenerate primers were used (I=inosine): GTIAT(AC)A-
CICCICTIACIGA (forward primer), GCIGGIACIGGIAA(AG)A-
CIGA (nested forward primer), C(GT)ICCIGC(AG)TAICCIG-
G(AG)TT (reverse primer for reverse transcription and first and
second PCR). A 323 bp fragment was subcloned and 15 clones were
sequenced.
Accession numbers
ift57/hppi, AY956331; dynein heavy chain 9, AY956332;
ift88/polaris, AY956333.
In situ hybridization
Whole-mount in situ hybridization was performed as previously
described (Thisse and Thisse, 1998) with some minor modifications.
For the polaris antisense probe the template (pCRII-TOPO-polaris
1200 bp, flanking primers forward AGCAGGCTGTCAGGA-
CAAGTC and reverse GTTTGAAGTCTCTCTGTCTTAGGT was
linearized with Not1 and the antisense riboprobes were transcribed
using SP6 RNA polymerase. The hippi antisense riboprobes were
generated using a BamHI linearized template and T7 RNA
polymerase. In situ hybridization experiments with southpaw (spaw)
(Long et al., 2003) and pitx2 (Yan et al., 1999) were performed using
standard techniques. Embryos were then mounted in Permount (Fisher
Scientific) and photographed on a Zeiss Axioplan microscope
equipped with a Zeiss AxioCam digital camera.
Development 132 (8) Research article
Development

1909Cilia and zebrafish organogenesis
Morpholino antisense oligonucleotides
Morpholino antisense oligonucleotides were designed either to
target the translation of the mRNA (abbreviation AUG) leading to a
protein knockdown phenotype or to target an exon splice donor site
causing splicing defects of the mRNA (abbreviation SP). For the
design of the antisense oligonucleotides the translation start site and
the splice donor site of the second coding exon were chosen and the
morpholino oligonucleotides obtained from GENE TOOLS, LLC,
Philomath, OR. The following morpholinos were used: polarisAUG
CTGGGACAAGATGCACATTCTCCAT, polarisSP AGCAGATG-
CAAAATGACTCACTGGG, hippiAUG CCTCCGCCATCCCT-
CTCTCTTTCT, hippiAUGmis CCTgCGgCATCCgTCTCTgTTaCT
(5 mismatches in lower case), hippiSP AGTGTTATCGCCT-
CACCAGGGTTCG, dhc9P1SP GATTTACACACCTTGTAGTC-
CATTT. The morpholinos were diluted in 100 mmol/l KCl, 10
mmol/l HEPES, 0.1% Phenol Red (Sigma). The injections
were done using a microinjector PLI-90 (Harvard Apparatus,
Cambridge, MA). The effect of the splice-morpholinos was verified
by RT-PCR from single embryo total RNA with nested primers in
flanking exons yielding a 300-600 bp amplicon. Rescue experiments
were done by co-injection of capped mRNA together with a
morpholino. Capped mRNA was made using mMESSAGE
mMACHINE (Ambion). For the injection of the capped mRNA or
capped mRNA together with morpholino, a microprocessor
controlled nanoliter injector Nanoliter 2000 (World Precision
Instruments, Inc.) was used.
Histochemistry and immunohistochemistry
Embryos were fixed in 2% glutaraldehyde/1.5%
paraformaldehyde/70 mmol/l Na
2
HPO
4
pH 7.2/3% sucrose at 4°C
overnight. After being washed in PBS and taken through an ethanol
dehydration series they were embedded in JB-4 resin (Polysciences
Inc.) and sectioned at 3-5 µm. Slides were stained in Methylene
Blue/Azure II (Humphrey and Pittman, 1974), mounted and
examined using a Nikon immunofluorescence microscope. For
acetylated tubulin staining, the embryos were fixed in Dent’s Fix
(80% methanol/20% DMSO) at 4°C overnight. After gradual
rehydration they were washed several times in 1PBS with 0.5%
Tween20 and blocked in 1PBS-DBT (1% DMSO/1% BSA/0.5%
Tween20) with 10% normal goat serum (NGS) (Sigma) at room
temperature for 2 hours. Primary antibody incubation in 1PBS-
DBT 10% NGS [1:500 monoclonal anti-acetylated tubulin 6-11B-1
(Piperno and Fuller, 1985) (Sigma)] was at 4°C overnight. The
embryos were washed in 1PBS with 0.5% Tween20 and blocked
in 1PBS-DBT 10% NGS at RT for 1 hour and then incubated in
1:1000 goat anti-mouse Alexa 546 (Molecular Probes) in 1PBS-
DBT 10% NGS at 4°C overnight. After rinsing in 1PBS, the
embryos were washed with methanol and equilibrated in clearing
solution (1/3 benzoyl-alcohol and 2/3 benzoyl-benzoate) and
examined using a Bio-Rad Radiance 2000 confocal microscope. Z-
stacks were acquired and used for creation of projections with
extended focus.
Cilia length measurements were performed using Image J 1.32j
(National Institute of Health) in two to four different embryos per
group. Confocal images where individual cilia base and ends could
be discerned (>60 individual measurements) were outlined and the
calculated length recorded. Our measurements may underestimate
cilia length owing to a foreshortening effect caused by viewing some
cilia at an angle.
For double labeling with two monoclonal antibodies, the embryos
were stained as above and the procedure was repeated with 1:20
monoclonal antibody alpha 6F, raised against the chicken alpha1
subunit of the Na+/K+ ATPase (Takeyasu et al., 1988), obtained from
the Developmental Studies Hybridoma Bank, as primary and 1:1000
goat anti-mouse Alexa 633 (Molecular Probes) as secondary antibody
after incubation with goat anti-mouse Fab fragments 1:20 in 1PBS-
DBT at 4°C overnight.
Electron microscopy
Embryos were prepared for electron microscopy by previously
published protocols (Drummond et al., 1998).
High speed videomicroscopy
PTU-treated embryos were put in E3 egg water containing 40 mmol/l
BDM (2,3-butanedione monoxime, Sigma), for 5 minutes to stop the
heartbeat and then changed to 20 mmol/l BDM containing egg water
for observation. The embryos were then analyzed using a 40/0.55
water immersion lens on a Zeiss Axioplan microscope (Zeiss,
Germany) equipped with a high-speed Photron FastCAM-PCI 500
videocamera (Photron LTD). Image acquisition of beating cilia was
250 frames per second and 1088 frames total per take by Photron
FastCAM version 1.2.0.7 (Photron LTD). Image processing was done
using Photoshop 7.0 (Adobe) and movies compiled in Graphic
Converter v.4.5.2 (Lemke Software, Germany). Three-dimensional
illustrations were drawn using Strata3D Software (Strata).
Fluorescent dye/bead injection and fluorescence
videomicroscopy
For urine excretion assays a 5% solution of Tetramethylrhodamin-
conjugated 70 k MW dextran (Molecular Probes) was injected into the
common cardinal vein (CCV) of 3.0-3.5 dpf. embryos anesthetized
with 0.2 mg/ml tricaine (3-aminobenzoic acid ethylester, Sigma) in
egg water, these were then examined using a 40/0.55 water
immersion lens on a Zeiss Axioplan microscope equipped with a MTI
SIT68 fluorescence camera the video was recorded at real time with
a Panasonic PV-8400 tape recorder. Digitalization was done using
SonicMyDVD Version 3.5.2 software (Adaptec), still frames were
captured using QuickTime v.6.5.1 and movies were recompiled by
Graphic Converter (Lemke Software, Germany). For the dye transport
in the central canal, the dye was injected into the brain ventricle of
60 hpf. embryos anesthetized with tricaine. Sequential images were
taken with a Nikon fluorescence microscope. Fluorescent beads of
0.02 µm diameter (Fluospheres (580/605), Molecular Probes) were
dispersed 1:50 in 0.1 mol/l saline with 0.1% Phenol Red (Sigma) and
used for injection into KV of embryos at 8-10 somite stage.
Statistics
The two-tailed Student’s t-test was applied to the quantitative results.
Results
Apical cilia are present in KV, in the central canal of
the spinal cord, and in the pronephric ducts
To survey zebrafish larvae for the presence of ciliated cells
during organogenesis, we performed immunocytochemistry
using an anti-acetylated tubulin antibody. Immunostaining
confirmed an earlier report that apical cilia are present in cells
lining KV at the 8-somite stage (Fig. 1A,D) (Essner et al.,
2002). Also, ependymal cells along the central canal bore
apical cilia at 24 hpf (Fig. 1B,E), as did pronephric duct cells
at 48 hpf (Fig. 1C,F). KV cilia and pronephric duct cilia
showed an ultrastructure consisting of nine peripheral
microtubules and a central pair (9+2 pattern; Fig. 1G,I),
whereas a 9+0 formation was present in ependymal cell cilia
along the central canal of the spinal cord (Fig. 1H). Outer
dynein arms were present in cilia of all three organs (arrows).
All kidney cilia examined were 9+2; no 9+0 cilia were found
in the zebrafish pronephros.
Cilia in pronephros, central canal and KV are motile
The presence of dynein outer arms in the cilia of all three
tissues suggested that these cilia are motile (Smith and Yang,
Development

1910
2004). We therefore examined cilia for motility using high-
speed videomicroscopy. To examine pronephric cilia,
embryos at 2.5 days post-fertilization (dpf) were treated with
butanedione monoxime (BDM) to stop the heartbeat and
circulation in order to eliminate glomerular filtration
pressure. Images were acquired at 250 frames per second and
then replayed in slow motion at 15 frames per second to count
the beat frequency. Under these conditions, cilia beating at a
frequency of 20.0±3.2 Hz (n=34) were observed in all parts
of the pronephros, including the tubules and ducts (see
Movies 1, 2 in the supplementary material). As a similar cilia
beat frequency was observed in embryos not treated with
BDM, we conclude that BDM does not influence cilia
motility. In BDM-treated embryos there was no luminal fluid
flow generated by glomerular filtration, so the observed cilia
movements must represent active beating and not passive
deflection. Rotational cilia movement generated a corkscrew-
like wave pattern in the lumen of the duct directed toward the
cloaca (Fig. 1K). While the majority of pronephric epithelial
cells displayed a single apical motile cilium, a subset of cells
with up to 16 apical cilia could be observed in the midpart of
the pronephric duct.
The cilia in the central canal of the spinal cord were filmed
under the same BDM conditions as above to avoid disturbances
by circulating blood cells. The ependymal cilia were
approximately 2 µm in length and also showed a rotary pattern
of motility (Fig. 1J). The frequency of rotation was
approximately 12 Hz (see Movie 4 in the supplementary
material).
The cilia in KV were similar in length (3 µm) to the
ependymal cilia and rotated in a counterclockwise orientation
(Fig. 1J; see Movie 8 in the supplementary material). In
addition to images of moving KV cilia themselves, cilia
motility could be detected by the movement of small pieces of
debris suspended in the fluid of KV. Debris was observed to
travel in a counterclockwise orbit, interrupted by small
counterclockwise spins corresponding to the radii of circular
cilia beat patterns (see Movie 9 in the supplementary material).
This type of particle and fluid movement was subsequently
confirmed with fluorescent bead injections (see below).
Cilia length is shortened in IFT morphants and oval
mutant embryos
In order to manipulate cilia structure and assess their function
in vivo, we cloned and disrupted the expression of zebrafish
homologs of the IFT proteins of polaris/IFT88/osm-5 and
hippi/IFT57/che-13. The sequence homology and identity
between human, mouse and zebrafish IFT genes are shown in
Fig. S1 in the supplementary material. In addition, we analyzed
the oval mutant (ovl
tz288b
), which carries a point mutation in
the zebrafish homolog of polaris/IFT88/osm-5 leading to a
protein truncation (L260X) (Tsujikawa and Malicki, 2004).
The in situ expression of both polaris(IFT88) and hippi(IFT57)
in 24-48 hpf embryos was ubiquitous with some enrichment
along the pronephric ducts (see Fig. S2 in the supplementary
material) and the brain ventricles, and also around KV (data
not shown).
Using morpholino antisense oligonucleotides (MO), we
disrupted protein function of the hippi and polaris genes. AUG
and SP-morpholinos were designed for both genes. The
effectiveness of SP-morpholinos was verified by RT-PCR using
RNA from single embryos. The results show that morpholino
suppression of mRNA splicing persists for at least 72 hours
(Fig. 2O,P; see Fig. S5 in the supplementary material). Whole-
mount immunostains for acetylated tubulin of 44 hpf. embryos
Development 132 (8) Research article
Fig. 1. Apical cilia are present in Kupffer’s vesicle, the central canal
of the spinal cord and pronephric ducts. Immunostaining of
acetylated tubulin. (A) Apical cilia are present in cells lining
Kupffer’s vesicle at the 8-somite stage (arrowhead) in midline
longitudinal sections. (D) Kupffer’s vesicle; higher magnification
(DAPI nuclear staining in blue). (B) Ependymal cells along the
central canal bear cilia at 24 hpf (arrowheads). (E) Cross section at
44 hpf; cilia arise from all cells of the spinal central canal. (C) Cilia
can also be seen in the pronephric duct at 48 hpf (arrowheads).
(F) Cells double stained for acetylated tubulin and the alpha1 subunit
of the NaK-ATPase confirmed the apical position of the pronephric
cilia. (G-I) EM cross sections of cilia in Kupffer’s vesicle (G) show a
9+2 structure; ependymal cell cilia (H) are 9+0 in structure;
pronephric cilia (I) are 9+2 with clear dynein outer arms (arrows).
Cilia beat pattern: (J) The cilia in the of the spinal central canal and
Kupffer’s vesicle rotate in an counterclockwise orientation. (K) In
the pronephric duct monociliated and multiciliated cells can be
observed. Their cilia beat in rotation like a corkscrew with an
undulating appearance along their longitudinal axis. Mean values for
cilia length and beat frequency are given for comparison. Scale bars:
100 µm in A-C; 10 µm in D-F; 250 nm in G-I. KV, Kupffer’s vesicle;
PND, pronephric duct; SCC, spinal central canal.
Development

1911Cilia and zebrafish organogenesis
were performed and the specimens
examined by confocal microscopy
with extended focus. Wild-type
pronephric tubules and ducts are
ciliated along the entire length of the
nephron. Individual cilia were
visible in the posterior segment of
the pronephric duct (Fig. 2A). In the
pronephros of IFT morphant
embryos and oval homozygous
mutants, severe shortening or
absence of cilia was observed along
the entire length of the pronephric
nephron, from the cloaca (Fig. 2B-
D) to the anterior region of the
pronephric tubules (data not shown).
Cilia length was reduced from wild-
type control 8.8±2.0 µm (n=107) to
polaris MO 2.5±1.9 µm (n=271,
P<0.001) and hippi MO 3.5±2.0 µm
(n=141, P<0.001). In oval
heterozygotes cilia were 10.0±2.5
µm (n=68) in length, while in oval
homozygotes they were 3.7±2.1 µm
(n=104, P<0.001). Ependymal cilia
of the spinal central canal were
similarly shortened or reduced in
number in the morphant and mutant
embryos (Fig. 2F-H) compared with
wild-type controls (Fig. 2E). Length
measurements of the cilia showed
2.1±0.7 µm (n=63) in wild type
versus 0.9±0.5 µm (n=21, P<0.001)
in polaris MO and 1.2±0.8 µm
(n=46, P<0.001) in hippi MO. At
the 8-10 somite stage KV cilia were
also shortened or missing in IFT
morphants compared with controls
(Fig. 2I-K). When present, cilia
length was 3.3±1.1 µm (n=119) in
wild type versus 2.0±0.8 µm (n=25,
P<0.001) in polaris MO and
1.4±0.6 µm (n=15, P<0.001) in
hippi MO. In several instances cilia
appeared largely absent in hippi MO
embryos (Fig. 2K). Although
reduced in length, pronephric cilia
in IFT morphant or mutant embryos
were comparable in structure and
maintained a relatively normal 9+2
microtubule doublet ultrastructure
(Fig. 2L-N).
Cyst formation and hydrocephalus in IFT morphants
and
oval
–/–
embryos
To determine whether the previously described phenotypes
associated with IFT88/polaris loss of function were also
observed when expression of other IFT proteins was disrupted,
we compared the phenotype of embryos injected with
morpholinos targeting both IFT88/polaris and IFT57/hippi.
Morpholinos targeting the translation start site (AUG) or the
splice donor site (SP) of the second coding exon of
IFT88/polaris or IFT57/hippi all led to pronephric cyst
formation, hydrocephalus, ventrally curved body axis and
pericardial edema in 72 hpf embryos (Fig. 3). Edema became
generalized by day 5/6, when most of the embryos died. The
kidney cyst and hydrocephalus were easily recognizable in the
living fish, in which pigmentation was inhibited by PTU and
also by histology (Fig. 3K). Body axis curvature, kidney cysts
Fig. 2. Cilia structure is altered in IFT morphant embryos. A whole-mount confocal
immunostaining of acetylated tubulin in 44 hpf embryos shows apical cilia in the pronephric ducts
(A-D), the spinal canal (E-H) and Kupffer’s vesicle (I-K). At 44 hpf polaris and hippi morphants
(B,C), as well as oval homozygous mutant embryos (D), exhibit severely shortened cilia
(arrowheads) compared with wild type (A). For reference, arrows in A-D indicate the point were
the pronephric ducts merge at the cloaca. (E-H) Cilia (arrowheads) in the central canal (lumen
indicated by arrow in E) of the spinal cord are also shortened or absent in polaris (F) and hippi (G)
morphants and in oval
–/–
embryos (H) compared with wild type (E). Immunostaining of
pronephros/central canal: anterior is to the left, dorsal to the top. Kupffer’s vesicle cilia in polaris
(J) and hippi (K) morphant embryos are greatly reduced compared with control (I). (L,M)
Ultrastructure of the pronephric cilia in wild-type (L), polaris (M) and hippi (N) morphants show a
typical 9+2 microtubule doublet pattern. (O) Molecular analysis of the effectiveness of SP-
morpholinos inducing splice defects: RT-PCR of single embryos generates a 354 bp polaris
fragment in control embryos, bridging part of exon 1 to part of exon 5 at 24, 48, 72 hpf (lane M, 1
Kb Plus DNA Ladder). polarisSP-injected embryos analyzed with the same primer sets at the
same timepoints show a larger amplicon of 457 bp caused by a non-splicing of intron 2, which
encodes a premature stop codon; the lower wild-type band recovers over time. Lower panel: RT-
PCR of
β
-actin of the same RNA samples. (P) RT-PCR of hippi mRNA results in a 553 bp
fragment in control embryos, whereas the amplicon is reduced in size in the hippiSP-treated
embryos, indicating an in-frame deletion of exon 2 and 3 (lower band, 260 bp) or an out-of-frame
deletion of exon 2 only (middle band, 378 bp); there is recovery of the wild-type band over time.
Development

1912
and hydrocephalus were also observed in oval (IFT88/polaris)
homozygous mutant embryos, confirming that the morpholino
phenotypes we observed are due to IFT88/polaris loss of
function (Fig. 3H) (Tsujikawa and Malicki, 2004).
The specificity of the morpholino was tested for the
hippiAUG morpholino, for which an introduction of five
mismatches completely abolished its effects (Fig. 3D). To
further establish the specificity of the morpholino action, we
performed rescue experiments for the hippiSP morpholino by
co-injection of capped RNA made in vitro from zebrafish hippi
cDNA. Zebrafish hippi mRNA injection alone did not cause an
obvious phenotype; injected embryos appeared wild type.
Co-injection of zebrafish hippi mRNA with morpholino
completely rescued 36 out of 44 injected larvae to a wild-type-
like phenotype; histological cross sections confirmed the
absence of hydrocephalus and cyst formation in the co-injected
larvae (see Fig. S3 in the supplementary material). Co-injected
capped murine hippi RNA with hippiSP morpholino showed a
complete rescue in 6 out of 26 embryos. Partial rescue was
observed in 14 out of 26 embryos, with embryos showing a
straight body axis and substantially reduced cyst formation. In
1 out of 26 doubly injected embryos, axis deformity and
hydrocephalus were observed, but no cysts formed and the
remaining five embryos did not show rescue (data not shown).
The data demonstrate that the observed phenotypes are specific
to loss of IFT57/hippi function and further suggest that
IFT57/hippi protein function is in large part conserved in
vertebrates.
Pronephric fluid flow is impaired in the IFT
morphant/mutant embryos and can lead to cyst
formation
The reduction in cilia length in IFT morphant embryos
suggested that cilia motility might also be affected and
contribute to the observed organ phenotypes. Indeed, in the IFT
morphant embryos and oval homozygotes, moving cilia were
rarely detected. The remaining motile cilia in these embryos
appeared to be stumpy and had a faster, uncoordinated
flickering movement (PolAUG 32.2±2.3 Hz, n=8, HippiSP
30.6±4.2 Hz, n=7, significantly different from control 20.0±3.2
Hz, n=34, P<0.001) (see Movie 6, Movie 5 in the
supplementary material). The cloaca-directed, helical wave
pattern of cilia beat observed in wild-type embryos was never
seen in IFT morphant tubules or ducts.
To test if disturbed ciliary motility had an impact on fluid
output from the pronephros, we performed dye excretion
experiments. Tetramethylrhodamine-conjugated dextran (70
kD) injected into the common cardinal vein of living 3.5-day-
old embryos was filtered in the glomerulus and excreted via
the pronephric ducts at the cloaca (Fig. 4C). The time span
after injection until the first visible
urine excretion at the cloaca was
4.5±2.9 minutes (n=12) in wild-type
Development 132 (8) Research article
Fig. 3. IFT morphant phenotype: kidney
cysts and hydrocephalus. Disruption of
polaris function by injection of polarisAUG
(B) and polarisSP (C,I) results in
hydrocephalus (black arrowhead),
pronephric cyst formation (arrow),
pericardial edema (white arrowhead), and
ventrally bent body axis at 72 hpf compared
with non-injected control embryos (A). This
phenotype imitates the oval homozygous
mutant (H), in which the polaris gene is
mutated. Heterozygous embryos are
indistinguishable from wild-type controls
(G). Embryos injected with the control
hippiAUG mismatch morpholino have a
normal morphology (D), whereas hippiAUG
(E) and hippiSP (F) cause a phenotype
similar to the polaris morphants and the oval
homozygous mutant. (J) Wild-type embryo
in longitudinal section. (K) hippiSP
morphant embryo showing severe
hydrocephalus (**) and kidney cyst (*).
(L) hippiSP morphant embryo showing axis
curvature, cysts (arrow) and hydrocephalus
(arrowhead). (M) Histological cross sections
of a 72 hpf control embryo show the midline
fused glomerulus, pronephric tubules and
pronephric ducts on either side.
(N,O) hippiSP morphant embryos at 72 hpf
show a cystic dilatation (*) of the pronephric
tubules with a stretched glomerulus in the
midline. (O) polarisAUG morphants show
kidney cysts (*) and distension of the
pronephric ducts. gl, glomerulus; pd,
pronephric ducts; pt, pronephric tubules.
Development

1913Cilia and zebrafish organogenesis
control embryos. A movie
available in the supplemental
data shows in fast motion how
the fluorescent urine output is
observed from 3-8 minutes post-
injection (see Movie 3 in the
supplementary material). In
the morphant embryos, dye
excretion fell to levels below our
detection limits; no ‘jet’ of
fluorescence at the cloaca was
observed in 9 out of 9
polarisAUG and 9 out of 9
hippiSP morphants, even at
timepoints more than 30 minutes
post-injection (Fig. 4G,K),
compared with a visible
excretion in 22 out of 27 wild
types. To demonstrate that the
failure to detect fluorescent
output was not because of
blocked glomerular filtration,
the embryos were sectioned and
examined for dye passage and
uptake by pronephric epithelial
cells: all embryos showed
endocytic uptake of the filtered
dye by proximal duct cells,
indicating that the fluorescent
dextran was efficiently filtered in
IFT morphant embryos (Fig.
4D,H,L). Similar to control
wild-type embryos, oval
heterozygotes showed dye
excretion starting at 5.3±0.4
minutes (n=2) after injection. By
contrast, two out of five oval
homozygotes did not show dye
excretion, and in the remaining
three embryos, dye excretion
was delayed, being first
detectable at 13.7±5.5 minutes
(n=3) (P=0.1, not significant)
after injection, and the flow of
excreted dye was markedly
reduced. In these embryos, only
the lumen of the common
pronephric duct was visibly
fluorescent (arrowhead), and
there was no ‘jet’ of
fluorescence in the medium
outside the cloaca (arrow) (Fig.
4R). The data indicate that cilia
function is required to maintain normal rates of fluid flow in
the pronephros.
To demonstrate that impaired fluid flow in the zebrafish
pronephros could lead to cyst formation, we mechanically
obstructed the pronephric ducts close to the cloaca (Fig. 4S).
Number 5 Inox tweezers were used to pinch and physically
obstruct the cloaca. This obstruction of fluid flow resulted in
rapid pronephric cyst formation, within 30 minutes. Cystic
distension of the pronephric tubules and the anterior segment
of the pronephric ducts occurred well anterior to the point of
occlusion. Cyst structure was verified in histological cross
sections (Fig. 4T,U). Taken together with our data on
disruption of cilia function, the results suggest that a reduction
in flow rate in the pronephros may lead to back pressure at the
site of fluid input to the pronephros and result in tubule luminal
expansion and cyst formation.
Fig. 4. Fluid flow is impaired by lack of normal cilia movement in the pronephros. Living embryos
were injected with 5% tetramethylrhodamine-conjugated 70 k MW dextran into the circulation. After
passage of the pronephric kidney, the dye was excreted at the cloaca (C, arrow). The images of the
first column (A,E,I,M,P) are transmitted light images. The images in the second column (B,F,J,N,Q)
were taken at 2-3 minutes post-injection, while the images in the third column (C,G,K,O,R) were
captured when maximum excretion was reached. The time after injection that the image was
captured is indicated in the bottom left of each panel. No fluorescent dye excretion via the cloaca
was observed at timepoints >30 minutes in polarisAUG morphants (n=9) or hippiSP morphants
(n=9), whereas in the control embryos excretion was observed in 22 individuals (n=27). On histology
of the same embryos, all showed endocytic uptake of the dye in anterior duct cells (arrowhead)
(D,H,L), indicating that the dye had been filtered via the glomerulus (double arrows) into the cyst
lumens (*). In oval heterozygous embryos, excretion started at 5.3±0.4 seconds (n=2) (O), and 3 out
of 5 oval homozygous embryos showed weak dye excretion at 13.7±5.5 seconds (n=3), whereas 2
did not show a visible output. In A-R anterior is to the left and dorsal is to the top. Mechanical
obstruction of the pronephric ducts close to the cloaca (S, arrow) causes cystic distension of the
anterior pronephric tubules within 30 minutes (T, arrow). The dilated tubules/glomerulus can be seen
in cross sections (U, arrows). Scale bar: 100 µm.
Development

1914
Distension of the brain ventricles is associated with
impaired fluid flow along the central canal of the
spinal cord
To determine whether a similar loss of fluid flow could account
for the distension of the brain ventricles seen in IFT morphant
embryos, we injected 70 kD rhodamine-dextran into the fourth
ventricle at the level of the hindbrain and labeled the
cerebrospinal fluid in order to monitor its transport along the
central canal of the spinal cord by fluorescence microscopy.
The embryos were pretreated with BDM in order to prevent
dye movement by an active circulation. The leading front of
the dye traveling along the central canal was imaged at various
timepoints and transport rate was quantified. In wild-type
controls, the mean velocity of fluid movement in the spinal
central canal was 27.0±1.9 µm/minute (n=4) (Fig. 5A,B),
whereas it was reduced in the polarisAUG morphants to
11.3±3.3 µm/minute (n=5, P<0.001) (Fig. 5D) and in the
hippiSP morphants to 12.0±1.7 µm/minute (n=3, P<0.001)
(Fig. 5F). Identical results were obtained for the oval mutant
(Fig. 5J). Impaired fluid flow probably results in fluid backup
and distension of the brain ventricles and the development of
hydrocephalus.
Disruption of dynein heavy chain 9 expression
partially phenocopies the IFT-morphants
Previous studies of the role of cilia in kidney cystic disorders
have suggested that kidney cilia are non-motile and act to sense
fluid flow. Our results in zebrafish indicate that cilia motility is
a primary factor in maintaining proper lumen size and kidney
function. To distinguish between sensory versus motile cilia
function, we sought to decrease cilia beat rate without causing
gross changes in cilia length or structure. The outer doublet
microtubules and associated dynein arms are critical for the
initiation and propagation of ciliary bending, while the central
pair/radial spokes system serves to regulate beat frequency and
wave form (Smith and Yang, 2004). In order to disrupt cilia
beat rate and pattern, we isolated an axonemal dynein heavy
chain by RT-PCR using degenerate primers and RNA from
isolated pronephric ducts of 72 hpf zebrafish larvae. A cDNA
encoding the P1 domain, the primary ATP binding site that is
essential for dynein motor function, of a dynein heavy chain
homologous to human dynein heavy chain 9 (DYH9,
AAF69004) was obtained. Injection of a morpholino
(dhc9P1SP) targeting the splice donor site of the P1-domain
coding exon resulted in mis-splicing of dhc9 mRNA,
producing either an out-of-frame deletion of the P1 ATP-
binding domain or truncation after the P1 domain (see Fig. S5
in the supplementary material). The reduction of dhc9 function
partially phenocopied the IFT morphants/mutants (Fig. 6A)
with injected embryos showing hydrocephalus and pericardial
edema. Histological sectioning revealed distension of the
pronephric tubules (Fig. 6G) and dilated pronephric ducts (Fig.
6H,I). The movement of the cilia in dhc9P1SP larvae was
significantly slower than in wild-type control embryos: wild
type 20.0±3.2 Hz (n=34) versus dhc9P1SP 14.7±3.9 Hz (n=15,
P<0.001) (see Movie 7 in the supplementary material). The
cilia beat frequency was also reduced in the central canal of
the spinal cord with wild type 12.3±3.4 Hz (n=18) versus
dhc9P1SP 7.3±1.9 Hz (n=6, P<0.001). The rate of dye
transport in the spinal canal was also reduced in dhc9P1SP
morphants (Fig. 6C) compared with wild type (Fig. 6B): wild
type 27.0±1.9 µm/minute (n=7) versus dhc9P1SP 20.6±2.6
µm/minute (n=7, P<0.001). Excretion of circulating 70 kD
rhodamine-dextran by the pronephros could be detected in only
5 out of 19 morphant embryos (Fig. 6L,M), and these embryos
showed a delayed onset of 12.8±5.9 minutes (n=5) versus
4.5±2.9 minutes (n=12) in wild-type controls (P<0.01) (Fig.
6J,K). The embryos used for this experiment had sufficient
circulation (circulating blood cells) despite pericardial edema.
The data indicate that a reduction in beat frequency without
changes in cilia structure is sufficient to cause the phenotypes
associated with IFT protein loss of function.
Development 132 (8) Research article
Fig. 5. Fluid flow is impaired by lack of normal cilia movement in
the central canal of the spinal cord. BDM-pretreated embryos were
injected with 5% tetramethylrhodamine-conjugated 70 k MW dextran
into the fourth brain ventricle and dye distribution along the central
canal of the spinal cord was recorded at various timepoints; shown
are 2 and 40 minutes post-injection. Control (A,B) and oval
heterozygous (G,H) show a distribution of the dye up to an anterior-
posterior level of the tip of the yolk extension at 40 minutes
(arrowheads) (B,H), whereas polarisAUG morphant (C,D), hippiSP
morphant (E,F) and oval homozygotes (I,J) show reduced dye
migration. Anterior to the left. Scale bar: 100 µm.
Development

1915Cilia and zebrafish organogenesis
Impaired fluid flow in KV is associated with laterality
defects
The presence of motile cilia in KV suggested that the fluid
enclosed by the KV epithelium is ‘stirred’ and that, similar to
the role of ‘nodal flow’ in the mouse, fluid movement might
also play a role in establishing proper organ laterality in
zebrafish. We visualized fluid movement in KV by injecting
small fluorescent beads into the lumen of KV of 8-10 somite
embryos. When viewed dorsally by videomicroscopy, injected
beads moved in a counterclockwise direction (Fig. 7A-D, see
Movie 10 in the supplementary material). Larger bead
aggregates tended to collect in the center of KV, where they
rotated in place in a counterclockwise direction. As noted
above, cilia beating and the movement of small pieces of debris
in KV was also in a counterclockwise direction (see Movie 9
in the supplementary material). To test whether fluid motion
in KV required intact cilia, we examined injected bead
movements in IFT88/polaris and IFT57hippi morphant
embryos. The absence of normal cilia in KV of IFT morphant
embryos completely abolished bead movement (Fig. 7E-H).
The data indicate that counterclockwise cilia beating drives
fluid in a counterclockwise flow pattern inside the confined
space of KV.
It has been suggested that in order to achieve right-to-left
fluid flow in the mouse node, only a portion of the circular cilia
beat must be involved in fluid
propulsion (Cartwright et al., 2004). If
cilia were, for instance, tipped toward
the posterior, then a cilium would
extend into node fluid only on the right-
to-left portion of the cycle. On the return
left-to-right portion of stroke, the cilium
would move close to the cell surface and
produce only a small drag on node fluid.
We therefore examined KV cilia by electron microscopy for
signs that the cilia might be polarized in an anterior-posterior
orientation. As shown in figure 7P, cilia and their associated
basal bodies on the dorsal aspect (‘roof’) of KV were observed
to be tipped to the posterior approximately 45° relative to the
surface of the roof. We also observed that the cell membrane
and cytoplasm adjacent to the cilium insertion appeared to be
a site of active vesicle fusion (Fig. 7P).
To test whether loss of cilia-dependent fluid flow in KV
resulted in laterality defects, we assayed expression of
two conserved left-right genes, the nodal-related protein
southpaw (spaw) and the bicoid-related transcription factor
pitx2, which is downstream of nodal. At mid-somite stages
(15-23) southpaw is expressed in the left lateral plate
mesoderm (LPM) (Long et al., 2003). pitx2 is also expressed
in the same location at similar stages (Campione et al., 1999;
Yan et al., 1999). Observed expression patterns for the IFT
morphant embryos are shown in Fig. 7I-O. In IFT morphant
embryos, right-sided or bilateral expression of southpaw was
significantly increased, and in 33% of polarisAUG embryos
southpaw expression was missing (Fig. 7R). Right-sided
pitx2 expression was increased in polarisAUG-injected
embryos compared with wild type, and the frequency of
absent signal was also increased. Significantly, hippiSP-
injected embryos showed a complete absence of pitx2
Fig. 6. Dynein heavy chain 9 knockdown
morphants show abnormal cilia movements
and phenotypic changes similar to the IFT
morphants. A morpholino targeting the
splice-donor site of the exon coding for the
P1-domain of dhc9 causes kidney cysts,
hydrocephalus (arrow) and axis curvature
(A). Sequencing RT-PCR of aberrant splice
products (A, inset) revealed non-splicing of
the adjacent intron with a premature stop
codon (upper band) and an out-of-frame
deletion of the P1-domain coding exon
(lower band). The transport of injected
fluorescent dye along the central canal of
the spinal cord (B,C) is impaired in dhc9SP
morphants (C) versus control (B).
Histologically, Dhc9P1SP morphant
embryos show distension of the tubules
near the glomerulus (G) and dilated ducts
(H) compared with wild-type control (D,E).
The dilatation of the duct can also be seen
in frames taken from Movie 7 (see
supplementary material) (F, wild type; I,
Dhc9P1SP morphant). (J-M) Dye excretion
via the urine was not detected in dhc9SP
morphants (arrows in L,M) versus control
(J,K).
Development

1916
expression and a near-complete absence of southpaw
expression (Fig. 7R).
Discussion
The recent convergence of studies of kidney cystic disease,
left-right asymmetry defects, retinal degeneration and flagella
formation has led to the idea that defects in the formation or
function of cilia may underlie pathologies observed in all these
conditions. However, despite the focus on cilia as a central
organelle in these phenotypes, it has been unclear what exactly
cilia do to support normal organ development and function
and how loss of this organelle can lead to such pathology.
Our analysis of cilia during zebrafish organogenesis has
demonstrated that cilia in the zebrafish pronephros, in the
central canal of the spinal cord and in KV are motile. Beating
cilia were found to induce fluid flow in all three organs. Lack
of proper cilia formation due to inhibition of IFT88/osm-
5/polaris or IFT57/che-13/hippi expression was associated
with loss of fluid movement and resulted in pronephric cyst
formation, left-right asymmetry defects and hydrocephalus.
We conclude that back pressure from blocked flow and
subsequent fluid accumulation may account for organ
distension pathologies in the brain and kidney, while the loss
of fluid flow in KV may result in absence of a mechanosensory
signal regulating organ laterality.
Development 132 (8) Research article
Fig. 7. Impaired fluid flow in Kupffer’s vesicle is
associated with defects in laterality. Embryos at the
8-10 somite stage were dechorionated and
fluorescent beads were injected into Kupffer’s
vesicle. Control embryos showed a rotating
movement of bead aggregates in a counterclockwise
orientation (arrowheads in A-C) when viewed
dorsally. Relative timepoints in seconds are
indicated in the bottom right of each panel. The
wall of Kupffer’s vesicle is indicated by dotted
lines. (D) Superposition and enlargement of frames
(A-C) with an additional transmitted light frame
showing the counterclockwise direction of
movement. None of the injected morphant embryos
(polarisAUG and hippiSP) showed this
phenomenon (E-H). Abnormal expression of the
laterality markers pitx2 and spaw in polarisAUG
and hippiSP embryos (I-O,R). In situ experiments
were performed on 14-somite (spaw) and 20-somite
(pitx2) embryos. Dorsal views of the lateral plate
mesoderm are shown with the different expression
patterns seen in polarisAUG embryos. southpaw
was expressed on the left (I), right (J) or bilaterally
(K), or in many cases absent (L). pitx2 shows the
same patterns (M, left-sided; N, right-sided; O,
absent expression), with the exception of bilateral
expression. Sagittal section electron micrograph (P)
of the roof of Kupffer’s vesicle showing, a single
cilium and associated basal body. (Q) Diagram of
the micrograph in P, detailing how the angle of the
basal body [approximately 45° to the posterior (P)]
would result in the cilium projecting into Kupffer’s
vesicle on the right-to-left portion of the
counterclockwise rotary beat. (R) Frequency of
laterality defects in polaris and hippi morphant
embryos. Expression of pitx2 and southpaw was
randomized in polarisAUG embryos, while hippiSP
embryos showed significantly higher numbers of
embryos with no expression of southpaw and pitx2
(control, n=25; polarisAUG/pitx2, n=29;
polarisAUG/southpaw, n=36; hippiSP/pitx2, n=40;
hippiSP/southpaw, n=57). In embryos lacking
laterality signals in the lateral plate mesoderm, gene
expression was nevertheless maintained in the
tailbud (spaw) and Rohon-Beard neurons (pitx2).
Scale bar: 10 µm. a, anterior; d, dorsal.
Development

1917Cilia and zebrafish organogenesis
Kidney cilia and cyst formation
Kidney cysts are the result of grossly expanded kidney tubule
lumens. In human diseases such as autosomal dominant
polycystic kidney disease, large numbers of cysts lead to kidney
fibrosis and end-stage renal failure. A role for cilia in this
disorder is implied from the variety of cilia-associated proteins
that, when mutated, can cause tubules to become cystic (Barr
et al., 2001; Blacque et al., 2004; Fan et al., 2004; Kim et al.,
2004; Morgan et al., 2002; Murcia et al., 2000; Mykytyn et al.,
2004; Otto et al., 2003; Pazour et al., 2000; Pazour et al., 2002;
Qin et al., 2001; Sun et al., 2004; Taulman et al., 2001; Yoder
et al., 2002). Cell culture studies of PKD1 and PKD2, the genes
responsible for autosomal dominant polycystic kidney disease,
suggests that they act together in epithelial cells to mediate
calcium entry upon flow-induced cilium deflection (Nauli et al.,
2003; Praetorius and Spring, 2001). This model of cilia function
proposes that the cilium acts as a passive sensor of tubule lumen
mechanics and flow, providing a feedback signal that somehow
limits lumen diameter. Our observation that zebrafish
pronephric cilia are motile expands the repertoire of functions
that kidney cilia can serve. Our results suggest that in more
primitive kidneys, and perhaps at the earliest stages of kidney
development, cilia can function as a motile ‘fluid pump’ to drive
fluid through the nephron. Our results are consistent with an
early report demonstrating that ciliated nephrons in the
amphibian Necturus can generate hydrostatic pressures of up to
4.0-5.7 cm H
2
O (White, 1929). In the elasmobranch (e.g.
dogfish, skate) kidney, cells bearing multiple 9+2 cilia similar
to those we describe in zebrafish, have been proposed to
transport mucus secreted by duct cells and keep the ducts patent
(Lacy et al., 1989).
Mammalian kidney cilia are not thought to be motile and
instead are proposed to serve a sensory function. Nonetheless,
some correspondence of our results to the metanephric kidney
may be seen in the context of early mammalian development
and human disease. In the human fetal kidney, bundles of 9+2
cilia have been observed in electron micrographs
(Zimmermann, 1971) in kidney tubule lumens. Bundles of 9+2
cilia in the tubule lumen have also been observed in the adult
human kidney under pathological conditions (Duffy and
Suzuki, 1968; Hassan and Subramanyan, 1995; Katz and
Morgan, 1984). Some primary cilia dyskinesia cases report an
association between cilia motility dysfunction and cystic
kidneys (Ibanez-Tallon et al., 2003) that, although rare, suggest
that loss of cilia motility may also be important in some human
cystic disorders. Obstruction of fluid flow has been identified
as a cause of a specific type of human glomerular cyst
formation occurring during fetal development (Potter, 1972;
Woolf et al., 2004). One perspective that could take into
account these observations in both fish and humans would be
that the cilia that form first in early mammalian kidney
development may be motile, recapitulating the cilia behavior
we see in the more primitive fish pronephros. As development
proceeds, cilia motility may be lost and cilia take on a new
sensory function in the mature mammalian kidney. Implicit in
this model is the idea that cyst formation, as a result of cilia
dysfunction, could be caused by multiple mechanisms in fetal
versus adult kidneys, and in pronephroi versus mature
metanephroi. As more refined models of cystic gene defects
are developed, e.g. conditional gene knockouts, these
speculative ideas can be rigorously tested.
Increased cell proliferation has been also cited as a
mechanism of cystic expansion in human disease (Nadasdy et
al., 1995; Nauli et al., 2003) and as an initiating stimulus in
some mouse models of cystic disease (MacRae Dell et al.,
2004). Although cell proliferation could play a role in cyst
expansion or progression in zebrafish, we have found no
evidence of an increase in cell number in zebrafish cysts.
Currently, a role for cell proliferation as an early, initiating
event in cyst formation in mouse models of ADPKD
(polycystin1 and polycystin2) and IFT mutants has yet to be
established with quantitative data. It is likely that kidney cysts
can arise from several different primary cellular defects,
including increased proliferation, loss of cilia function and
general cell dedifferention (reviewed by Arnaout, 2001). In this
view, the initial stimulus for cyst formation may vary
depending on the gene mutated.
In zebrafish, complete obstruction of the pronephric duct
caused tubular distension within minutes, indicating that
blocking fluid flow is sufficient for cyst formation. In IFT
mutants/morphants it is likely that flow is reduced, but not
completely blocked. Cyst formation in these larvae occurs
more slowly over a period of hours after hatching. The double
bubble pronephric cyst mutant, for instance, which we have
recently found to be defective in cilia formation (T. Obara and
I.A.D., unpublished), has a patent pronephric duct lumen based
on serial sectioning, and forms cysts between 2 and 2.5 dpf
(Drummond et al., 1998). Also, while the excretion rate in IFT
morphants and oval homozygotes was not sufficient to generate
a jet of fluorescent urine, dye fluorescence was visible in the
common pronephric duct lumen, indicating that the duct
remains unobstructed. It is striking that complete obstruction
initiates cyst formation only in the anterior pronephric tubules
and not, for instance, along the length of the pronephric duct.
The anterior pronephric tubules and glomerulus is also the
initial site of cyst formation in all zebrafish cyst mutants
reported (Drummond et al., 1998; Sun et al., 2004). Only
several hours after initial anterior cyst formation is observed
does the duct lumen begin to expand, for instance as we report
here for the dhc9 morphant. It is possible that the anterior
tubule/glomerulus may be the most labile structure in the
forming pronephros at the time when voluminous fluid flow
begins (at hatching?) and thus most distensible by fluid
pressure. It is notable that many zebrafish cyst mutants show a
curled body axis (Drummond et al., 1998; Sun et al., 2004). It
is unlikely that the reduction in flow/cyst formation we observe
is a secondary consequence of body curvature, because many
mutants exist with ventral axis curvature that do not develop
cysts in the kidney, and initiation of cyst formation occurs prior
to the development of axis curvature (Drummond et al., 1998).
Motile cilia in the brain and hydrocephalus
Retention of cerebrospinal fluid in the brain ventricles by
malabsorption or impaired drainage causes a distension of the
brain ventricles or hydrocephalus. Our results demonstrate that
motile cilia in the spinal canal are necessary to maintain normal
cerebrospinal fluid distribution and that impaired fluid flow
results in a backup of fluid in the central canal and brain
ventricles. Our results are consistent with previous
observations that human ependymal cilia are motile
(Worthington and Cathcart, 1963; Worthington and Cathcart,
1966). In addition, patients with PCD suffer from
Development

1918
hydrocephalus in addition to respiratory syndromes associated
with loss of lung cilia function. Mice with mutations in
mdnah5, hfh4 and polaris (IFT88/tg737) (Ibanez-Tallon et al.,
2003) all exhibit hydrocephalus. While our work was in review,
Ibanez-Tallon and co-workers demonstrated that in mouse
mdnah5 mutants the movement of beads injected into the brain
ventricle was impaired, further implying a role for motile cilia
in the hydrocephalus seen in these animals (Ibanez-Tallon et
al., 2004). Although additional driving forces for fluid flow
along the central canal may exist (for instance, fluid secretion
and reabsorption), motile cilia appear to be crucial for normal
cerebrospinal fluid flow rates.
The 9+0 cilia are thought to be immotile as they lack the
central microtubule pair normally associated with motile cilia.
Ependymal cilia in zebrafish have a 9+0 axonemal
microtubular pattern and yet are motile, indicating that the
presence of a central microtubule pair is not a prerequisite for
motility. This is similar to the mouse node, where 9+0 cilia
beat in a rotary fashion (Nonaka et al., 2002). The presence of
dynein arms on the outer microtubule pairs may be a better
predictor of whether a cilium is sensory (immotile) or motile.
A central pair in 9+2 cilia may have more relevance to the cilia
wave form. The beat pattern of 9+2 cilia has been described as
a planar waveform (O’Callaghan et al., 1999; Shimizu and
Koto, 1992; Smith and Yang, 2004). Zebrafish ependymal cilia
and mouse node cilia beat in a simpler rotary pattern (McGrath
et al., 2003).
Motile cilia in KV and laterality defects
Previous work in the mouse has demonstrated that cilia
function and nodal flow are required in the establishment of
left-right asymmetry (Bisgrove et al., 2003; Hamada et al.,
2002). The ciliated epithelium of the mouse ventral node has
been shown to cause fluid flow in a right-to-left direction across
its surface (Nonaka et al., 1998; Sulik et al., 1994). The
direction of this fluid flow seems to be crucial, as inverting the
direction causes situs inversus, and no flow causes
randomization of the left right axis (Nonaka et al., 2002). Fluid
flow may also be important in determining situs in humans, as
evidenced by the random organ situs seen in patients suffering
from PCD, in which cilia motility is impaired (Afzelius, 1985;
Ibanez-Tallon et al., 2003). In zebrafish and other teleosts, KV
is the functional equivalent of the mouse node (Brummett and
Dumont, 1978; Essner et al., 2002). Early morphological
studies in Fundulus heteroclitus showed clearly that the cells
of the dorsal ‘roof’ of KV are uniformly ciliated (Brummett
and Dumont, 1978). Support for a role for KV in left-right axis
determination was demonstrated recently by the finding that
the T-box transcription factor no tail is required for the
morphogenesis of KV and no tail mutant embryos exhibit
randomized left-right axes (Amack and Yost, 2004).
Importantly, selective suppression of no tail function in the
dorsal forerunner cells, the progenitors of KV, specifically
inhibits KV development in the absence of other embryonic
defects and leads to randomization of the left-right axis
(Amack and Yost, 2004). We show that, like the mouse node,
the zebrafish KV is a ciliated structure and a site of dynamic
cilia-driven fluid flow. We observe that flow occurs in a
circular, counterclockwise direction. However, some aspects of
our data would also suggest that the primary propulsive force
is in a right-to-left direction, similar to the mouse node. First,
cilia anchored in the roof of KV are tipped toward the posterior.
As suggested by previous modeling studies (Cartwright et al.,
2004), cilia beating in a counterclockwise direction at this
angle would be predicted to extend into KV fluid on the right-
to-left stroke and pass along the cell surface on the return left-
to-right stroke. The predicted result of this beat pattern would
be active propulsion in the right-to-left (extended) stroke and
substantially less propulsion as the cilium glides over the cell
surface. In addition to the ultrastructural evidence, support for
this idea can be seen in the movies of KV bead-injected
embryos, where bead movement appears to be faster in the
right-to-left direction. In Movie 10 (in the supplementary
material), the bead aggregate makes four trips around the
periphery of KV. The average time for right-to-left transit
(2.6±0.13 s.e.m. seconds) is significantly less than for left-to-
right transit (3.8±0.15 s.e.m. seconds). While it is clear that
more detailed quantitative studies will be required to
extrapolate on these observations, the results suggest that KV
fluid may be driven in a right-to-left direction; the left-to-right
movement completing the circular pattern may be passive
return flow. A passive return flow might also be expected to
occur in vivo in the mouse node (Cartwright et al., 2004). This
is because the node in mouse is also a closed structure, i.e. the
ciliated node surface is covered and enclosed by Reichert’s
membrane in the embryo (Nonaka et al., 1998). In most flow
studies to date, Reichert’s membrane is first removed to gain
access to the node surface (Nonaka et al., 1998).
Two hypotheses have been put forward to suggest a
mechanism implicating fluid flow in left-right axis
determination. In simple terms, the alternatives are that: (1) a
morphogen gradient is established by right-to-left fluid flow
(Nonaka et al., 1998; Okada et al., 1999); or (2) fluid flow per
se is sufficient to provide a mechanical signal that breaks
left-right symmetry, possibly by stimulating non-motile,
mechanosensory cilia and subsequent intracellular calcium
signaling (McGrath et al., 2003; Tabin and Vogan, 2003).
Given that the KV is a closed vesicle and fluid flow inside the
vesicle is circular, it seems unlikely that the role of fluid flow
would be to drive a morphogen to one side of the zebrafish
embryo, although models for such an effect of cilia have been
proposed (Cartwright et al., 2004). In our view it is more likely
that counterclockwise fluid flow is sensed to generate an
asymmetric signal; however, at present the underlying
mechanisms are unknown. We have also observed laterality
defects in polycystin2 knockdown zebrafish embryos (Obara et
al., unpublished), similar to that seen in the mouse. polycystin2,
as a member of the TRP mechanosensory ion channels, may
play a role in transducing KV fluid flow in the zebrafish.
Our results are consistent with the idea that cilia in the KV
fulfill a function analogous to cilia in the node region of mouse,
i.e. they generate a leftward flow that induces left-side specific
gene expression. However, because IFT proteins and cilia have
also been implicated in other processes (Huangfu et al., 2003;
Tsujikawa and Malicki, 2004), it cannot be formally excluded
that the left-right defects are caused by mechanisms unrelated
to nodal flow. For example, in the mouse, IFT proteins have
been implicated in hedgehog signaling (Huangfu et al., 2003),
and hedgehog signaling has been implicated in left-right
patterning (Zhang et al., 2001). Hence, some IFT mutants
might affect left-right development by disrupting hedgehog
signaling in the mouse. This scenario is less likely in zebrafish,
Development 132 (8) Research article
Development

1919Cilia and zebrafish organogenesis
because loss of hedgehog signaling does not lead to left-right
defects (Chen et al., 2001). The widespread expression of IFT
genes in early embryos also leaves open the possibility that
IFTs could function in dorsal forerunners or other cells and
tissues that form before KV. As our experiments do not identify
the stage at which IFT proteins act in the process of
establishing left-right asymmetry, we cannot rule out a role for
hippi or polaris prior to the formation of KV. Loss of IFT
function could conceivably have additional effects on KV-
associated gene expression that precedes left-sided southpaw
expression and that could influence the competence of KV to
generate a left-sided signal. While we have not ruled this out,
the observation that in IFT morphants, southpaw and pitx2
expression was not affected in tissues other than the lateral
plate mesoderm indicates that loss of IFT function does not
have widespread effects on the expression of these two
markers. It is reasonable to expect that multiple, redundant
mechanisms may act to establish left-right asymmetry, some of
which may function to maintain, propagate or amplify other
signaling systems. At present, we favor a direct role of IFT
proteins and cilia in left-right patterning by generating flow in
KV as one such signaling system that is now amenable to
experimental manipulation in the zebrafish.
In summary, by analyzing the phenotypes of zebrafish IFT
protein morpholino knockdowns and an IFT88 point mutant
(oval), and by disrupting expression of a dynein heavy chain,
we show that cilia-driven fluid flow is crucial for the early
development of zebrafish embryos. Fluid flow in KV correlates
with determination of the left-right axis and its impairment
causes laterality defects. Compromised fluid flow along the
central canal of the spinal cord correlates with backup of fluid
in the brain ventricles, leading to hydrocephalus. In the
pronephros, cilia motility is required for high rates of flow.
Disruption of pronephric fluid flow leads to cyst formation in
zebrafish. These results should serve to refocus attention on
biological fluid dynamics as one common mechanism
underlying various disorders of epithelial tissue structure and
function.
The authors would like to thank members of the Drummond lab,
Stephanie Wiessner, Jinhua Zhao, Tomoko Obara and Narendra
Pathak, for the helpful discussions and support. We also thank Dr
Jarema Malicki for providing the oval mutant; Mary McKee and
Dennis Brown for assistance with electron microscopy; Dirk
Hentschel for contributing to the development of dextran injection
methods; Richard Bouley, Valerie and Nicolas DaSilva of the MGH
Membrane Biology Group for helpful discussions; Margaret Boulos,
Humberto Urquiza, Amy Doherty, Marcellino Pina and Eric Stone for
aquaculture support; and the other members of the Developmental
Biology Lab at the Massachusetts General Hospital for critical input
into this work. Genomic sequence used in this work was produced by
the Zebrafish Sequencing Group at the Sanger Institute and can be
obtained from ftp://ftp.ensembl.org/pub/traces/zebrafish.
This work was supported by NIDDK grants DK53093 and
DK54711 to I.A.D., DK65655 to B.K.Y. and NIH 5RO1 GM56211
to A.F.S. A.F.S. is an Irma T. Hirschl Trust Career Scientist and an
Established Investigator of the American Heart Association. C.J.H. is
supported through a T32 training grant (DK07545) to Dr D. Benos.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/132/8/1907/DC1
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