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{at}receptor.mgh.harvard.edu)
Accepted 4 February 2005
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SUMMARY |
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Key words: Cilia, Pronephros, Kupffer's vesicle, Ependymal cell, Spinal canal, Kidney cyst, Hydrocephalus, Left-right asymmetry
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Introduction |
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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
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.
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Materials and methods |
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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
CCCTTTGCGAGTAAAGAGTGTTAAATGTGA, reverse CATCATCTGCTGCAAACTAGCCCTCT, nested
primer-set forward CGGGATCCGCCACCATGGCGGAGGAGGAAGAG reverse
CGGAATTCCGGCGGTGAGTGTGTGTTTCAATA) 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 AAATTGTGTTTGGAAATCAATGCAACA, nested
primer-set forward CGGAATTCGCCACCATGGCCGCCGCGGCCGCG, reverse
GCTCTAGAGAAGCATGGAAGCCCACGTGTTTA). 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)ACICCICTIACIGA (forward primer), GCIGGIACIGGIAA(AG)ACIGA (nested forward primer), C(GT)ICCIGC(AG)TAICCIGG(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
AGCAGGCTGTCAGGACAAGTC 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.
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 AGCAGATGCAAAATGACTCACTGGG,
hippiAUG CCTCCGCCATCCCTCTCTCTTTCT, hippiAUGmis
CCTgCGgCATCCgTCTCTgTTaCT (5 mismatches in lower case), hippiSP
AGTGTTATCGCCTCACCAGGGTTCG, dhc9P1SP GATTTACACACCTTGTAGTCCATTT. 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
Na2HPO4 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 1xPBS with 0.5% Tween20 and blocked in
1xPBS-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
1xPBS-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 1xPBS with
0.5% Tween20 and blocked in 1xPBS-DBT 10% NGS at RT for 1 hour and then
incubated in 1:1000 goat anti-mouse Alexa 546 (Molecular Probes) in
1xPBS-DBT 10% NGS at 4°C overnight. After rinsing in 1xPBS,
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 1xPBS-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 40x/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 40x/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.
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Results |
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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 (ovltz288b), 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 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).
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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 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.
|
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.
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|
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 expression and a near-complete absence of
southpaw expression (Fig.
7R).
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Discussion |
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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 H2O (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
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, 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.
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ACKNOWLEDGMENTS |
---|
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.
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Footnotes |
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Supplementary material for this article is available at http://dev.biologists.org/cgi/content/full/132/8/1907/DC1
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